Bio Prospecting for Hyper Lipid Producing Micro Algal Strains for Sustainable Biofuel Production...

Bioresource Technology 102 (2011) 57–70

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Bioresource Technology

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Bioprospecting for hyper-lipid producing microalgal strains for sustainablebiofuel production

T. Mutanda a, D. Ramesh a, S. Karthikeyan b, S. Kumari a, A. Anandraj c, F. Bux a,*

a Institute for Water and Wastewater Technology, Durban University of Technology, Durban 4001, South Africab Tamil Nadu Agricultural University, Coimbatore 641 003, Tamil Nadu, Indiac Department of Nature Conservation, Mangosuthu University of Technology, Durban 4026, South Africa

a r t i c l e i n f o a b s t r a c t

Article history:Received 30 March 2010Received in revised form 9 June 2010Accepted 17 June 2010Available online 10 July 2010

Keywords:BiofuelBioprospectingMicroalgaeSamplingStrain identification

0960-8524/$ - see front matter � 2010 Elsevier Ltd. Adoi:10.1016/j.biortech.2010.06.077

* Corresponding author. Tel.: +27 31 3732597; fax:E-mail address: [email protected] (F. Bux).

Global petroleum reserves are shrinking at a fast pace, increasing the demand for alternate fuels.Microalgae have the ability to grow rapidly, and synthesize and accumulate large amounts (approxi-mately 20–50% of dry weight) of neutral lipid stored in cytosolic lipid bodies. A successful and econom-ically viable algae based biofuel industry mainly depends on the selection of appropriate algal strains. Themain focus of bioprospecting for microalgae is to identify unique high lipid producing microalgae fromdifferent habitats. Indigenous species of microalgae with high lipid yields are especially valuable in thebiofuel industry. Isolation, purification and identification of natural microalgal assemblages using con-ventional techniques is generally time consuming. However, the recent use of micromanipulation as arapid isolating tool allows for a higher screening throughput. The appropriate media and growth condi-tions are also important for successful microalgal proliferation. Environmental parameters recorded atthe sampling site are necessary to optimize in vitro growth. Identification of species generally requiresa combination of morphological and genetic characterization. The selected microalgal strains are grownin upscale systems such as raceway ponds or photobireactors for biomass and lipid production. Thispaper reviews the recent methodologies adopted for site selection, sampling, strain selection and identi-fication, optimization of cultural conditions for superior lipid yield for biofuel production. Energy gener-ation routes of microalgal lipids and biomass are discussed in detail.

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1. Introduction

The depletion of fossil fuel reserves has caused an increase indemand and price of diesel. The uncertainty in their availabilityis considered to be the important trigger for researchers to searchfor alternative sources of energy, which can supplement or replacefossil fuels (Harun et al., 2010; Mata et al., 2010). In recent years,research has been directed to explore alternate fuels from variousbiological renewable sources. Biodiesel is an alternative to dieselfuel, which is produced from oils via transesterification. It is non-toxic, biodegradable and has the potential to replace the conven-tional diesel fuel. The use of biodiesel will ultimately leads toreduction of harmful emissions of carbon monoxide, hydrocarbonsand particulate matter and to the elimination of SOx emissions,which can also help in reducing the greenhouse effects and globalwarming. Presently, biodiesel is produced from different crops,such as, soybean, rapeseed, sunflower, palm, coconut, jatropha,karanja, used fried oil and animal fats (Spolaore et al., 2006; Khanet al., 2009). There will be certain limitations in the use of these oils

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as alternate fuels because of its food demand, life span, lower yield/ha, higher land usage and higher price inter alia (Mata et al., 2010).It is necessary to search for non food based alternate feedstocks forbiodiesel production. Selection of biodiesel feedstock is based onhigher yields, short duration, lower production cost and less landusage. Among the various biodiesel feedstocks, the microalgae oilhas the potential to replace the conventional diesel fuel.

In order to avert fuel shortages in the future, a substantialamount of financial resources (more than 300 million dollars) hasbeen set aside to facilitate basic research in phycology to enableresearchers in the tropics and subtropical regions to search andcollect microalgae for evaluation of their feasibility for biofuel pro-duction (Sheehan et al., 1998). Microalgae are desirable for biofuelproduction as compared to plants because of the following rea-sons: (1) microalgae have fast growth rates, high biomass yield po-tential using non-fresh water streams as substrate, (2) microalgalbased biofuels do not interfere with food security concerns, (3) bio-fuels generated from microalgal lipids have less emissions and con-taminants as compared to petroleum based fuels therefore reducedgreenhouse gas emissions and (4) microalgae require non-arableland for their cultivation and can utilise industrial flue gas as car-bon source and moreover it can be harvested daily (Chisti, 2007;

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Table 1Potential of microalgae as primary PUFA resources (Spolaore et al., 2006).

PUFA Potential application Microalgal producer

Docosahexaenoic acid (DHA) Infant formulas; nutritional supplements; aquaculture Crypthecodinium, chizochytriumEicosapentaenoic acid (EPA) Nutritional supplements, aquaculture Nannochloropsis, Phaeodactylum Nitzschia, Pavlovac-Linolenic acid (GLA) Infant formulas; nutritional supplements SpirulinaArachidonic acid (AA) Infant formulas; nutritional supplements Porphyridium

58 T. Mutanda et al. / Bioresource Technology 102 (2011) 57–70

Greenwell et al., 2009; Griffiths and Harrison, 2009; Rodolfi et al.,2009; Mata et al., 2010).

To date, the main focus of research in the field of biofuels frommicroalgae has been centred on downstream aspects such asbioreactor designs, biomass and lipid production from microalgae,biomass harvesting techniques and the chemistry of biofuel pro-duction. Microalgal bioprospecting encompasses searching andcollection of unique microalgal strains from different aquatic envi-ronments for exploiting the potential applications of value addedproducts such as polyunsaturated fatty acids (Olaizola, 2003;Spolaore et al., 2006) (Table 1). A lot of literature is available onthe mass production and sustainable use of microalgae for bio-diesel production and little emphasis placed on an in-depth studyof microalgal bioprospecting. Therefore, the main objective of thisreview paper is to report current strategies focusing on biopro-specting for microalgae with the main aim of producing biofuels.This manuscript investigates current developments in microalgaebioprospecting. Protocols and procedures employed for successfulmicroalgal bioprospecting are presented and described in-depth.Microalgal sampling, storage conditions and isolation and strainselection procedures are also discussed in detail.

2. Microalgae

Microalgae are unicellular microscopic (2–200 lm), polyphy-letic, noncohesive, artificial assemblage of CO2 evolving, auto-trophic organisms which grow by photosynthesis and are theeukaryotic representatives although the prokaryotic cyanobacteriaare frequently included with the algae (Greenwell et al., 2009). Al-gae are also defined as thallophytes (plants lacking roots, embryos,vascular system, stems and leaves) that have chlorophyll-a as theirprimary photosynthetic pigment and lack a sterile covering of cellsaround the reproductive organs (Brennan and Owende, 2010). Al-gae can either be autotrophic or heterotrophic; the former requireonly inorganic compounds such as CO2, salts and a light energysource for growth; while the latter are nonphotosynthetic there-fore require an external source of organic compounds as well asnutrients as an energy source (Brennan and Owende, 2010). Theevolutionary history and taxonomy of microalgae is complex dueto constant revisions as a result of new genetic and ultrastructuralevidence. The main criteria for categorising microalgae are pig-mentation, life cycle and basic cellular structure (Brennan andOwende, 2010). Microalgae are classified into two prokaryotic divi-sions (Cyanophyta and Prochlorophyta) and nine eukaryotic divi-sions (Glaucophyta, Rhodophyta, Heterokontophyta, Haptophyta,Cryptophyta, Dinophyta, Euglenophyta, Chlorarachniophyta andChlorophyta). However according to Khan et al. (2009), the most

Table 2The four most important groups of algae in terms of abundance (Khan et al., 2009).

Algae Known species (approx.) Storage mater

Diatoms (Bacillariophycea) 100,000 ChyrsolaminarGreen algae (Chlorophyceae) 8000 Starch and TAGBlue–green algae (Cyanophyceae 2000 Starch and TAGGolden algae (Chrysophyceae) 1000 TAGs and carb

important groups of algae in terms of abundance are: diatoms,green algae, blue–green algae and golden algae (Table 2). Thereis potential for further exploitation of these organisms for produc-tion of value added products and biofuels.

3. Sampling

Microalgal collection is mainly influenced by environmentalfactors (both biotic and abiotic), parameters measured onsite, typeof aquatic system and sampling equipment. The collection methodadopted is crucial for success, because damaged or dead cells maylead to failure. Therefore the temporal and spatial collection strat-egy should be adopted to cater for any succession that can occur atthe sampling site (Anandraj et al., 2008; Bernal et al., 2008).

For successful biofuel production using microalgae as feedstockfor biomass and lipid accumulation, the crucial step is to search,collect and identify hyper-lipid producing strains. Selection offast-growing, productive strains, optimized for the local climaticconditions are of fundamental importance to the success of any al-gal mass culture and particularly for low-value products such asbiodiesel. According to Borowitzka (1997), it is also important toevaluate harvesting costs at the time of choosing the species.Low-cost harvesting requires large cell size, high specific gravitycompared to the medium and reliable autoflocculation. The sche-matic outline of the procedures involved in upstream to down-stream processing of microalgal lipids is presented in Fig. 1.

3.1. Sampling environments

The mega biodiversity of microalgae entails them as suitablecandidates for biofuel production. It is estimated that there are be-tween one and ten million algal species, most of them being mic-roalgae. Microalgae require light, CO2, appropriate pH, suitablesalinity, macronutrients (nitrates and phosphates), vitamins andtrace elements for their growth (Chisti, 2007; Brennan andOwende, 2010). These nutritional and environmental requirementsare variable under natural conditions.

Microalgae are found in diverse environmental conditions andhabitats such as lacustrine, brackish, freshwater, hyper saline,wastewater maturation ponds, dams, rivers, marine and coastalareas. These natural ecosystems have immeasurable value assources of hyper-lipid producing microalgae and it has been re-ported that many microalgal species tolerate brackish and salinewaters. Table 3 depicts lipid content of some microalgal strains col-lected from different aquatic environments. Due to selection pres-sure and changing environmental conditions, there are a widerange of microalgal species worldwide found in extreme environ-

ial Habitat

in (polymer of carbohydrates) and TAGs Oceans, fresh and brackish waters Fresh waters Different habitats

ohydrates Fresh water

Isolation of microalgae using traditional and advanced techniques

Assessment of Growth/ Physiology

Qualitative and Quantitative analysis of lipids

Evaluation in open and closed systems

Recommendation for mass production

Enrichment of culture sample

Strain maintenance and Storage in repository

Collection of microalgal samples from different aquatic environments

Biofuel production

Purification

Strain Identification by microscopic and molecular tools

Fig. 1. Steps involved and outcome of bioprospecting of microalgae for biofuels production.

T. Mutanda et al. / Bioresource Technology 102 (2011) 57–70 59

ments (Rodolfi et al., 2009). In any habitat, microalgae have beenshown to have successional tendencies due to variable nutrientavailability, inclement weather and seasonal variations (Bernalet al., 2008). In bioprospecting it is important to collect microalgalsamples temporally and spatially so as to determine if there areany successional tendencies in the habitat. According to Anandrajet al. (2008), microalgal biomass has shown clear temporal andspatial patterns during the heterogeneous conditions of the openand closed phases in estuaries. The microalgae are found as amixed consortium and the population dynamics of the microalgaein any habitat is complex (Bernal et al., 2008). Different types ofmicroalgal strains require different habitats, hence the diversityof environmental conditions where microalgae can be collected.

In tropical and subtropical countries, inland dams, lakes and riv-ers have seasonal variations in ambient water temperature ranging

from 10 to 30 �C. Lower winter temperatures promote the growthof benthic microalgal strains such as Senedesmus sp. whereas sum-mer temperatures enhance the growth of the Chlorophytes such asChlorella strains. The average pH in these habitats also varies from3 to 8.5 depending on the prevailing edaphic conditions and agri-cultural practices in the area. In addition, the nutritional composi-tion of the aquatic system is a major factor for microalgae to thrivein these habitats. The major macronutrients required for microal-gal growth are phosphates, nitrates and ammonium and theseare required in suitable concentrations to promote lipid synthesisby the microalgae (Celekli and Yavuzatmaca, 2009; Chen et al.,2009; Hsieh and Wu, 2009).

Brackish aquatic systems constitute a mixture of seawater andfresh water and this usually occurs at the river mouth on the coast-line. The ecological and physico-chemical parameters in this habi-

Table 3Habitats and oil content of some microalgae.

Microalga Oil content(% dry weight)

Habitat

Botryococcus braunii 25–75 Fresh water/estuaryMonodus subterraneus UTEX 151 16.1 FreshwaterChlorella vulgaris CCAP 211/11b 19.2 FreshwaterChlorococcum sp. UMACC112 19.3 FreshwaterScenedesmus sp. F&M-M19 19.6 FreshwaterScenedesmus sp. DM 21.1 FreshwaterChlorella sp. 28–32 FreshwaterNeochloris oleoabundans 35–54 Fresh waterCrypthecodinium cohnii 20 MarineTetraselmis sueica 15–23 MarineMonallanthus salina >20 MarineDunaliella primolecta 23 MarinePhaeodactylum tricornutum 20–30 MarineIsochrysis sp. 25–33 MarineNannochloris sp. 20–35 MarineCylindrotheca sp. 16–37 MarinePavlova salina CS49 30.9 MarineSkeletonema sp. CS252 31.8 MarineChaetoceros muelleri F&M-M43 33.6 MarinePavlova lutheri CS182 35.5 MarineChaetoceros calcitrans CS178 39.8 MarineNitzschia sp. 45–47 MarineNannochloropsis sp. 31–68 MarineSchizochytrium sp. 50–77 Marine

Modified from Chisti (2007), Rodolfi et al., (2009).


tat are mainly a result of the dissolved CO2, O2 and dissolvedsalts due to eddies and turbulence created as the two water bodiesmix. A number of microalgal strains prefer brackish conditionsbecause of the nutritional composition of the aquatic system andthe warmer temperatures (Woelfel et al., 2007). Most of the micro-algae found in brackish conditions are found in suspensionas a consequence of the rapid water movements (Anandraj et al.,2008).

Hypersaline environments (halophilic), thermophilic springsand maturation ponds are ideal extreme environments for the iso-lation and selection of lipid producing microalgae with novel prop-erties. Isolates collected from these aquatic habitats are robust andhave rare and unique characteristics and therefore possibly betteradapted to specific conditions (Sheehan et al., 1998). The water col-lected from hypersaline aquatic bodies can be used for media for-mulation for microalgal growth in photobioreators and racewayponds under controlled laboratory conditions. For the purpose ofbioprospecting, it is imperative to screen as many aquatic environ-ments as possible to increase the potential to isolate and select un-ique hyper-lipid producing microalgae.

3.2. Parameters to measure on site

It is critical to measure some parameters on the collection sitein order to simulate these conditions when culturing specimensunder laboratory conditions. Elementary factors influencing micro-algal growth are measured on the sampling site. These include abi-otic factors such as light (quality and quantity), water temperature,nutrient concentration (nitrates and phosphates), dissolved O2, dis-solved CO2, pH, and salinity. Technically it is impractical to mea-sure biotic factors on site such as pathogens (bacteria, fungi andviruses) and any competitors in the habitat. These can be analysedmicroscopically and using conventional microbiological methodsin the laboratory after collecting the water samples. However if re-sources and time are available it is desirable to measure the oper-ational factors such as shear produced by mixing, dilution rates,depth and water velocity i.e. if the water is flowing. The global

positioning system (GPS) coordinates of the sampling locationmust be recorded for future reference and resampling (Woelfelet al., 2007).

3.3. Sampling equipment and procedure

The microalgal sampling and selection process is well estab-lished although it requires specialised equipment and may be timeconsuming (Anandraj et al., 2007). The equipment required formicroalgal sampling includes a knife, mesh net (0.7 lm mesh),scooping jar, vials for collecting samples, scalpels, dissolved CO2

and O2 analyser with a data logger, light meter, GPS, salinity meter,multi probe system (measuring pH, temperature, turbidity, con-ductivity and light intensity simultaneously). Heavy duty equip-ment includes a suitable vehicle for rough terrain with enoughspace for the collected samples.

There is no definite sampling procedure documented in litera-ture though researchers can follow simple and cheap methods ofcollecting microalgal samples. Ideally samples can be collectedfrom the natural substrata by chipping, scrapping, and by brushingfrom rock surfaces and bottom sediments. The brushing methodwas reported to be effective and reproducible method of collectingmicroalgal cells and also that it does not damage them (MacLulich,1986). Sampling in deep freshwater lakes and dams requires sys-tematic sampling whereby water samples are scooped from atleast three depth levels to the bottom of the lake or dam. This willallow for collection of microalgae which prefer different lightintensities. Bottom sediments are also major habitats of benthicmicroalgae and therefore should be collected together with piecesof detritus and mud. Stringent regimes and protocols needto be exercised when sampling. Therefore an all encompassingsampling regime is essential to isolate microalgae from aquaticenvironments.

4. Isolation and purification techniques

4.1. Culture media

Various culture media have been developed for isolation andcultivation of microalgae (Table 4). Some of them are modificationsformulated after detailed study on the nutrient requirement of theorganism. In case of marine algae, though artificial sea water mediais common, good growth of algae can be achieved by adding smallquantities of nonpolluted natural seawater (less than 1–4%) to theartificial seawater (Andersen, 2005). Schreiber solution consistingof a mixture of nitrate and phosphate was devised, based on theminimum requirement for the two elements shown by a diatomculture. Soil extracts were added to Schreiber’s medium for grow-ing the green dasycladalean Acetabularia and unicellular and ben-thic marine algae. Earlier algal media were devised to includeantibiotics, vitamins, trace metals and later with EDTA, a metabol-ically inert chelator, to replace organic chelators such as citrate(Andersen, 2005). Likewise modified media supporting growth ofa wide range of microalgae can be formulated by carefully manip-ulating major nutrients (Scott et al., 2010). Antibiotics can beadded to the growth medium to discourage growth of contaminat-ing cyanobacteria and other bacteria. Addition of germanium diox-ide inhibits the growth of diatoms. Axenic cultures can be obtainedby treating isolated algae to an extensive washing procedure, and/or with one or more antibiotics. In addition, bacterial contamina-tion in microalgal purification can be prevented by employing aprocedure involving treatment with a detergent and phenol (Abuet al., 2007). Therefore media composition is vital and most impor-tantly, media has to be inclusive of essential components that are

Table 4Common media used for microalgal strains from different aquatic environments.

Media Freshwater Marine Brackish Suitable for Reference(s)

AF6 medium, modified + + � Euglenophyceae, volvocalean algae, xanthophytes,many cryptophytes, dinoflagellate and green ciliate;specific for algae requiring slightly acidic medium

Watanabe et al. (2000)

AK medium � + + Broad spectrum marine algae Barsanti and Gualtieri (2006)Beijerinck medium + � � Freshwater Chlorophyceae Barsanti and Gualtieri (2006)BG-11 + + + Freshwater soil Cyanophyceae Barsanti and Gualtieri (2006)Bolds basal medium + � � Broad spectrum medium for freshwater

Chlorophyceae, Xantophyceae, Chrysophyceae, andCyanophyceae; unsuitable for algae with vitaminrequirements

Barsanti and Gualtieri (2006)

COMBO Medium � + + Cyanobacteria, cryptophytes, green algae, anddiatoms

Kilham et al. (1998)

Diatom Medium, modified + Freshwater diatom Cohn et al. (2003)DY-III medium + � � Freshwater Chrysophyceae Barsanti and Gualtieri (2006)ESAW medium � + + Broad spectrum medium for coastal and open ocean

algaeBerges et al. (2001)

K medium � + + Broad spectrum medium for oligotrophic marinealgae

Barsanti and Gualtieri (2006)

L1 medium � + + For oligotrophic (oceanic) marine phytoplankters Guillard and Hargraves (1993)Medium F � + + Broad spectrum medium for marine algae Jeffrey and LeRoi (1997)Medium G + + ND Broadspectrum medium Blackburn et al. (2001)MNK medium � + ND General medium for marine algae especially

coccolithophoresNoël et al. (2004)

ND, not determined; +, can be used; �, cannot be used.


found in the natural environment to support and not exclude thegrowth of isolates of interest.

4.2. Conventional methods

4.2.1. Single-cell isolationPerhaps the most common method for single-cell isolation is by

micropipette, although automation is more advantageous. Micro-pipette isolation is usually performed with a Pasteur pipette or aglass capillary having a straight or bent or curved tip. The goal ofmicropipette isolation is to pick up a cell from the sample, depositit without damage into a series of sterile droplets, until a single al-gal cell, free of all other protists, can be confidently placed into theculture medium. Subsequently, the sample can be examinedmicroscopically in a glass or plastic dish, in a multiwell plate, oron a microscope slide. However, the microalgal droplets can beplaced on agar to reduce evaporation but this step depends onthe size of the cells. Furthermore, the single cell can be pipettedand discharged into the sterile rinsing droplet and before the cellcan settle, it should be picked up and transferred. Skill of the tech-nique is important not to shear or damage the cell. For flagellates,cessation of swimming sometimes indicates damage. For diatoms,broken frustules can refract light differently than for intact cells.Leakage of protoplasm is an obvious sign of severe damage. Roger-son and co-workers, (1986) employed repeated introduction andejection of cells, suspended in a 1% crude papain solution, intoand from a micropipette to generate ca. 10% naked cells of Coscin-odiscus asteromphalus. These naked cells were reisolated via micro-pipette into fresh medium. The traditional method of micropipetteisolation can be successfully employed with certain precautions.Ultraclean droplets for rinsing are necessary, because the tiny cellscannot be easily distinguished from particles, especially whenworking with seawater.

4.2.2. Isolation using agar platesIsolation of cells on agar plates is also an old and common

method. It is the preferred isolation method for many coccoid algaeand most soil algae, not only for ease of use but also because axeniccultures can often be directly established without further treat-ment. For successful isolation onto agar, the alga must be able to

grow on agar as streak or pour plates (Brahamsha, 1996). In mostcases, the concentration of agar is not an important factor, assum-ing the agar is between 0.8% and 1.5–2.0%. Some algae do not growon the surface of agar plates, but they do grow embedded in agar.

4.2.3. Atomized cell spray techniqueA fine or atomized spray of cells can be used to inoculate agar

plates. The technique can vary, but in general a liquid cell suspen-sion is atomized with forced sterile air so that cells are scatteredonto the plate (Andersen, 2005). For best results, the spray canbe administered in a sterile hood or clean environment so that air-borne bacteria and fungi are not dispersed onto the plate. Theplates are incubated, and after colony formation, selected cellsare removed and inoculated.

4.2.4. Dilution techniquesThe goal of the dilution method is to deposit only one cell into a

test tube, flask, or well of a multiwell plate, thereby establishing asingle-cell isolate (Andersen, 2005). If the approximate cell concen-tration is known, then it is easy to calculate the necessary dilutionso that, on the basis of probability, a small volume contains a singlecell. The technique can be altered in several ways like dilution withculture medium, distilled water, seawater, filtered water from thesample site, or some combination of these. Also, ammonium, sele-nium, or another element can be added to some isolation tubes orcell wells to specifically select for species that require these nutri-ents (Andersen, 2005).

4.2.5. Gravimetric separationGravity separation can be effective for separating larger and

smaller organisms, and the two primary methods are centrifuga-tion and settling. Gravity is perhaps more frequently used to con-centrate the target organisms rather than to establish unialgalcultures. Usually, the goal is to separate the larger and heavier cellsfrom smaller algae and bacteria. Gentle centrifugation for a shortduration brings large dinoflagellates and diatoms to a loose pellet,and the smaller cells can be decanted. Centrifugation with use ofdensity gradients (e.g., Silica sol, Percoll) has been used to separatemixed laboratory cultures where each species was separated into asharp band (Andersen, 2005). Settling is effective for non-swim-


ming large or heavy cells. Both centrifugation and settling tech-niques are effective for concentrating larger cells, but single-cellisolation is nearly impossible and hence combined with othertechniques.

4.2.6. Media enrichmentEnrichment cultures have long been used as a preliminary step

towards single-cell isolations. Common enriching substances in-clude culture medium, soil water extract, or nutrients like nitrate,ammonium, and phosphate or a trace metal. One strategy of bacte-ria-free algal cultures is to make the medium as acidic as possiblewithout killing the alga. Peat moss can be substituted for soil–water extract when enriching for desmids and some other algaefrom acid habitats. Organic substances, such as yeast extract, case-in, or urea can be added in low concentrations when isolatingosmotrophic algae. Tiny amounts of various fruits and vegetableswere added to the culture in trying to isolate Oxyrrhis in a mixedculture of phytoplankton and it was discovered that unfilteredlemon juice, and subsequently extracts from lemon rind, led tosuccess, because of the availability of ubiquinone or plastiquinone(Andersen, 2005).

Natural samples are often deficient in one or more nutrients,but in nature algae survive, because bacterial action, grazing, anddeath of organisms recycle those nutrients. Sampling reduces recy-cling, and nutrient stress can cause death to the target species.Sometimes enrichments can also be detrimental, if the target spe-cies is rare and unable to compete with weedy species. The enrich-ing substance can be varied and usually added in minimumquantity and in stages. Selective culturing is an exceptional toolfor enrichment culturing of hyper-lipid producing microalgae.Since the goal is to isolate microalgae growing at a high CO2 con-centration, then an enrichment culture can be aerated with 1–5%CO2 and thus select for species with high CO2 tolerance (de Moraisand Costa, 2007; Ramanan et al., 2010). Although conventionalmethods for isolation has sufficed, there are limitations and there-fore continued need for developing novel methods for isolation ofmicroalgae.

4.3. Advanced methods

Algal cultures may be unialgal which means they contain onlyone kind of alga, usually a clonal population (but which may con-tain bacteria, fungi, or protozoa), or cultures may be axenic mean-ing that they contain only one alga. Streaking and spraying areuseful conventional techniques for single-celled, colonial, or fila-mentous algae growing on agar surface. Many flagellates, how-ever, as well as other types of algae must be isolated by single-organism isolations using advanced techniques. Among algal cellmanipulation techniques, the techniques for cell separation andisolation with high specificity are limiting because cell popula-tions are frequently heterogeneous and the cells are suspendedin a solution of different chemicals, biomolecules, and cells. Com-pared with traditional separation methods, microfabricated de-vices have small working volume and subsequently reducedthroughput.

4.3.1. MicromanipulationA micromanipulator allows the selection of single cells from li-

quid culture. This would mean that a single cell can be selectedfrom an enriched environmental sample and grown in liquidmonoculture as well as plates. This will afford a time saving of�60% over the current serial plating technique. This equipment isstipulated in literature as the ideal tool for algal screening and iso-lation (Kacka and Donmez, 2008; Moreno-Garrido, 2008). The suc-cessful application of micromanipulation techniques requires theexpertise and experience. It requires the handling of an Inverted

microscope or stereo microscope with magnification up to 200�.Phase contrast or dark field optics is an advantage. Capillary tubesor haematocrit tubes of approximately 1 mm diameter � 100 mmlong are used for picking individual cells (Godhe et al., 2002;Knuckey et al., 2002).

4.3.2. Flow cytometryFlow cytometric analysis permits the investigator to perform a

rapid and quantitative version of the experiments that could other-wise be performed by fluorescent microscopy. Fluorescence-Acti-vated Cell Sorting (FACS) allows the process to be taken one veryimportant step further. Cells with a specific characteristic or in-deed a combination of characteristics can be separated from thesample for further analysis or growth. Electronic cell sorting forisolation and culture of dinoflagellates and other marine eukary-otic phytoplankton was compared to the traditional method ofmanually picking cells using a micropipette (Sinigalliano et al.,2009). Trauma to electronically sorted cells was not a limiting fac-tor, as fragile dinoflagellates, such as Karenia brevis (Dinophyceae),survived electronic cell sorting to yield viable cells. The rate of suc-cessful isolation of large-scale (>4 l) cultures was higher for man-ual picking than for electronic cell sorting (2% vs. 0.5%,respectively). However, manual picking of cells is more labor-intensive and time consuming. Direct electronic single-cell sortingwas more successful than utilizing a pre-enrichment sort followedby electronic single-cell sorting. The appropriate recovery mediummay enhance the rate of successful isolations. Seventy percent ofisolated cells were recovered in a new medium, which was opti-mized for axenic dinoflagellate cultures. The greatest limiting fac-tor to the throughput of electronic cell sorting is the need formanual post-sort culture maintenance. However, when combinedwith newly developed automated methods for growth screening,electronic single-cell sorting has the potential to accelerate the dis-covery of new algal strains (Sinigalliano et al., 2009).

4.4. Other methods

Immunological and non-immunological methods to separatecells of interest are available. Conventional cell separation can becarried out by immunoreactions of membrane protein with thecapturing antibodies because the type of integrated proteins is spe-cific for their function. Immunological technique is a mainstay ofcommercialized cell separation methods such as the fluores-cence-activated cell sorting (Takahashi et al., 2004); magnetic-acti-vated cell sorting (Han and Frazier, 2005); affinity based cellsorting (Chang et al., 2005). One of the advantages of this methodis high specificity and selectivity. However, the disadvantage isthat the immunologically isolated cells may suffer from damagesand overall separation system involves high cost and complicatedprocesses such as immunoreactions and elution of cells from thecapturing antibodies.

On the other hand, non-immunological methods are relativelyfast and simple techniques. They include techniques such as dielec-trophoresis (Doh and Cho, 2005); hydrodynamic separation (Shev-koplyas et al., 2005); aqueous two-phase system (Yamada et al.,2004) and ultrasound separation (Petersson et al., 2004). These ex-ploit an interactive physical property of cell with the surroundingmedia. However, a disadvantage is its low specificity for cell sepa-ration, as cells do not show remarkable differences between eachcell type with the exception of immunological properties. In thismethod, the type of cells is determined and separated accordingto their cell size, shape and other physical properties. The advanta-ges and disadvantages of the microalgal purification techniquesdescribed in this section are summarised in Table 5.

Table 5Advantages and disadvantages of microalgal purification techniques.

Techniques Advantages Disadvantages Reference(s)

Pringsheim’s micropipettemethod

Single cells can be successivelytransferred and purified

Laborious and time-consuming method requiringconsiderable manual skills.

Melkonian (1990)

The method often fails with small non-flagellatecells, which are more difficult to recognise duringserial transfersSome delicate flagellates are easily damaged duringsuccessive micropipette transfers.

Agar plating (or spraying) Relatively easy Cannot be used with most flagellate taxa which failto grow on solid substrates

Serial dilution Relatively easy Unsuccessful when the numerical ratio betweenalgae and bacteria is unfavourable.

Brahamsha (1996)

Differential centrifugation Less damaging to sensitive cells Costly methodFiltration Less damaging to sensitive cells and usually

gives better separation of algae from bacteriathan differential centrifugation.

It is problematical with small algal cells and withcells secreting mucilage because of bacteriaembedded in the mucilage which may also clogfilters.

Melkonian (1990)

Use of antibiotics Relatively easy Damage the alga as well as leads to increasedresistance levels in contaminating bacteriaLow cost

Flow cytometry Precise and rapid method Require considerable costs for equipment and itsoperation

Sensen et al. (1993)

Simultaneous measurements of individualparticle volume, fluorescence and light scatterproperties

Require multi-user or central facilities.

Highly suitable for separating bacteria fromalgae to establish axenic algal cultures

Axenic cultures are difficult to obtain from algae towhich bacteria are physically attached.

Can be used directly in natural samplesUseful for small and delicate taxa

Ultrasonication Useful for separating attached bacteria fromalgal cell walls or mucilage

Not a standalone method Steup and Melknonian(1981)Should be coupled subsequent to cell sorting


5. Screening of microalgae

The screening stage of bioprospecting focuses on isolation andidentification of algal species capable of substantial lipid produc-tion, targeting organisms with rapid growth rate and tolerance toenvironmental parameters. The conventional method used for lipiddetermination involves solvent extraction and gravimetric deter-mination (Bligh and Dyer, 1959). A major disadvantage of the con-ventional method is that it is time consuming, labor-intensive andit has a low throughput screening rate. Moreover, approximately10–15 mg wet weight of cells, (Akoto et al., 2005) must be culturedfor the extraction and derivatization. Consequently, there is greaterinterest on a rapid in situ measurement of the lipid content (Cook-sey et al., 1987).

Nile Red (9-diethylamino-5H-benzo[a]phenoxazine-5-one), a li-pid-soluble fluorescent dye, has been commonly used to evaluatethe lipid content of animal cells and microorganisms (Genicotet al., 2005) and especially microalgae (Cooksey et al., 1987; Elseyet al., 2007). Nile Red possesses several characteristics advanta-geous to in situ screening. It is relatively photostable, intenselyfluorescent in organic solvents and hydrophobic environments.The emission maximum of Nile Red is blue-shifted as the polarityof the medium decreases, (Cooksey et al., 1987; Greenspan andFowler, 1985; Laughton, 1986; Lee et al., 1998) which allows oneto differentiate between neutral and polar lipids at the excitationand emission wavelengths. Elsey et al. (2007), showed the techni-cal emission spectra for Nile Red in various solvents. The peakemission intensity of Nile Red in hexane occurs near 576 nm whenexcited at 486 nm. The chloroform and ethanol peak excitation wasrecorded at 600 and 632 nm, respectively (Elsey et al., 2007). Inacetone, Nile Red is excited at 488–525 nm and the fluorescentemission measured at 570–600 nm using various instruments(Cooksey et al., 1987). Measurement of neutral lipids using the NileRed application requires the instrument to be calibrated using thestain dissolved in an organic solvent and account for the nonlinearintensity emission with respect to time. Measurements of lipid per

unit cell, requires a calibration curve that correlates fluorescence tolipid content, whether determined gravimetrically or by use of li-pid standards (Elsey et al., 2007). Thick cell walls of microalgae in-hibit the permeation of Nile Red and may indicate the absence ofoil, even though gravimetric analysis shows high yields of neutrallipids. It has been noted that the permeation of Nile Red dye is alsovariable among algal species, requiring the use of high levels ofDMSO (20–30% vol./vol.) and elevated temperatures (40 �C) (Chenet al., 2009).

Alternatively, the lipophilic fluorescent dye BODIPY 505/515(4,4-difluoro-1,3,5,7-tetramethyl-4-bora-3a,4adiaza-s-indacene) hasrecently been used as a vital stain to monitor algal oil storage with-in viable cells. Lipid bodies are stained green and chloroplasts redand visualized in live oleaginous (oil-containing) algal cells (Coo-per et al., 2010) (Fig. 2). The advantage of BODIPY is that high lipidyielding cells may be identified and isolated microscopically usinga micromanipulator system, flow cytometry or a fluorescence-acti-vated cell sorter (Cooper et al., 2010). Subsequently, pure culturesmay be propagated from the isolated viable cells. BODIPY 505/515has been shown to have a narrower emission spectrum than NileRed, making it potentially more useful for confocal imaging, wherefluorescence contrast enhancement of lipid bodies is important forimage resolution (Cooper et al., 2010). Unlike Nile Red, BODIPY505/515 has the advantage that it does not bind to cytoplasmiccompartments other than lipid bodies and chloroplasts.

A recent study (Dean et al. 2010) demonstrated the use of Fou-rier transform infrared micro-spectroscopy (FTIR) to determine li-pid and carbohydrate content of freshwater microalgae. FTIR wasshown to be an efficient and rapid tool for monitoring lipid accu-mulation of microalgae. This study had reported highly significantcorrelations between the FTIR- and Nile Red-based lipid measure-ments. For the purposes of bioprospecting for high lipid yieldingmicroalgae, a rapid throughput of sample processing is required.The semi-quantification of neutral lipids using Nile Red or BODIPYand fluorescence microscopy allows for an initial rapid screeningand visualization of lipid globules. FTIR spectroscopy may be used

Fig. 2. (A) Nile Red stained Chlorella sp. viewed at 1000� using a fluorescence microscope at 490 nm excitation and 585 nm emission filters. Neutral lipid globules in thecytosol are stained yellow (unpublished data). (B) Oil-containing lipid bodies can be vitally stained and visualized in live oleaginous (oil-containing) algal cells using the greenfluorescent dye, BODIPY 505/515. In this picture, vitally stained lipid bodies (green) are easily distinguished from chloroplasts (red) in an O. maius Naegeli freshwaterfilamentous algal cell using a Zeiss Axioskop epifluorescence microscope (Cooper et al., 2010). (For interpretation of the references to colour in this figure legend, the reader isreferred to the web version of this article.)


thereafter to quantify the yield of lipids. Once the high lipid pro-ducing microalgae have been identified, isolated and purified, afurther step in the screening would be to determine the photosyn-thetic efficiency of the culture.

Subsequent to screening, understanding the physiology of thealgal isolate is imperative. Pulse Amplitude Modulated (PAM) chlo-rophyll-a fluorescence measurements are widely used as a simple,rapid, and non-invasive method to assess the physiological state ofmicroalgae (Schreiber et al., 1994). It is also a valuable tool to as-sess the optimum growth conditions required to maximize the bio-mass yield and to quantify the effect of nutrient or other extremeenvironmental stresses (salinity, temperature, PAR and pH) onthe algal culture. Neutral lipid synthesis is stimulated under nutri-ent depleted or limited conditions. Many microalgae have the abil-ity to produce up to almost 80% dry cell weight of triacylglycerols(TAG) as a storage lipid (Chisti, 2007; Spolaore et al., 2006) undernutrient or other environmental stress. The PAM fluorometerparameters (Electron transport rate [ETR], maximum quantum effi-ciency of Photosystem II [FV/Fm], and non-photochemical quench-ing [NPQ]) may be used as indicators of nutrient stress andconsequently the possibility of neutral lipid synthesis and can bea valuable instrument in the screening process. Neutral lipid syn-thesis is likely to occur during the stationary phase of growthdue to nutrient limitation (Li et al., 2008).

The screening process of microalgae bioprospecting has to becomprehensive in assessing the lipid producing potential as wellas the kinetics of growth and tolerance. The success of downstreamprocessing is dependent on reliable biochemical and physiologicalscreening tools such as the BODIPY lipid stain, FTIR spectroscopyand PAM Fluorometry.

6. Lipid analyses

6.1. Pre-treatment of algae biomass for lipid extraction

The harvesting of algae biomass can be achieved by physical,chemical or biological methods or combination of any two of thesemethods. The techniques currently employed in microalgae har-vesting cum cell recovery include centrifugation, flocculation, fil-tration, gravity sedimentation, flotation and electrophoresis.Among these methods, centrifugation is found to be the most effi-cient. Heasman et al. (2000) studied algae cell recovery at three dif-ferent centrifugation conditions for ten microalgal species andreported that efficiency of recovery decreased with decreasing

acceleration. They recorded the highest cell recovery at 13,000g(�95%) followed by 6000 g (60%) and 1300 g (40%). Highest cellrecovery obtained at 13,000g (�95%) followed by 6000 g (60%)and 1300 g (40%). Cell viability was found to depend on the micro-algal species and the method of centrifugation. Before extractingthe algae oil, high moisture present in algae biomass must be re-moved by means of drying. The temperature used in microalgaedrying is crucial factor for separating the oil from dried algae bio-mass. Higher drying temperature decreases both concentration oftriacylglycerides and lipid yield (Widjaja et al., 2009). The algaebiomass dried at 60 oC under vacuum (Widjaja et al., 2009) orfreeze drying gave the best results for extraction of algae lipid.The lipids present in the dried/freeze dried biomass must be ex-tracted and analysed for quantification of the fatty acids. The lipidsproduced by algae are often accumulated intracellularly, whichcould require extraction of the lipids from crude cell pastes (Grimaet al., 2003). These techniques are used for rupturing the algae cellsto release the lipid compounds of algae biomass. Different cell dis-ruption techniques such as autoclaving, microwaves, sonication,osmotic shock and bead beating have been evaluated in order toincrease lipid extraction efficiency (Lee et al., 1998, 2010). Theyconcluded that microwave oven method was most simple, easy,and efficient method for lipid extraction from microalgae.Although many methods for algal lipid extraction have been rec-ommended, the most popular is a slightly modified method ofBligh and Dyer (1959). The solvent extraction was still the mainextraction method used by many researchers due to its simplicityand relatively inexpensive requiring almost no investment forequipment (Letellier and Budzinski, 1999).

6.2. Sample preparation for lipid analysis

Qualitative and quantitative composition of lipids can be inves-tigated by established techniques. Thin-layer chromatography(TLC), high pressure liquid chromatography (HPLC) or gas chroma-tography (GC) or any chromatography with mass spectrometry aresome of the different techniques employed for quantification of li-pid from microalgae. Lipid profiling of biodiesel feedstock is com-monly done by GC with a Flame Ionization Detector (FID)according to ASTM D6584 and EN 14105 methods. Before the fattyacid components of algae lipids can be analysed by GC, it isnecessary to convert them to low molecular weight non-polarderivatives, such as fatty acid methyl esters (FAME). Direct transe-sterification (simultaneous extraction and transesterification) andtransesterification (transesterification after lipid extraction) were


adopted in sample preparation for algae lipid conversion to biodie-sel (Johnson and Wen, 2009). Triglycerides in oils are reacted withmethanol in presence of base/acidic catalyst in transesterificationreaction to produce FAME. Acid catalysts used in transesterificationof microalgal oil includes sulphuric acid and hydrochloric acid,while alkali catalysts like sodium hydroxide and potassiumhydroxide are suitable for vegetable oil and animal fat respectively(Huang et al., 2010). After transesterification, the microalgal fattyesters develop slight green colour due to presence of chlorophylls.Before chromatographic analysis, any traces of chlorophylls, cata-lyst or water must be removed to avoid GC column contaminations(Sheehan et al., 1998).

6.3. Characterization of algal oil for biodiesel production

The fatty acid compositions of microalgae oil may vary to indi-vidual species/strains and their environmental conditions. Thefatty acid esters compositions of microalgae sources are differentfrom plant oils. Algal oils contain a high degree of polyunsaturatedfatty acids with four or more double bonds when compared to veg-etable oils (Zittelli et al., 2006; Rodolfi et al., 2009). The identifica-tion of lipid composition in selected algae strain is essential fordetermining the suitability to biodiesel and fuel quality. The mostimportant fuel properties considered to assess the potential of bio-diesel as substitute of diesel fuel are viscosity, cetane number (CN),density, cold filter plugging point, oxidative stability, lubricity,ignition quality and combustion heat (Xiaoling and Qingyu,2006). The physical characteristics of both fatty acids and triglycer-ides can be determined by chain length, number of double bondsand amount of each fatty ester components in both fatty acidsand triglycerides (Mittelbach and Remschmidt, 2004; Ramoset al., 2009). The saponification values (SV) and iodine value (IV)represent the ignition quality of fuel and presence of unsaturatedfatty acid component in FAMEs. Higher CN values indicate betterignition properties of the fuel (Meher et al., 2006). Higher unsatu-rated fatty acids (UFA) present in the oil gave higher iodine valuesand heating of higher UFA caused the polymerization of glycerides,which leads to formation of deposits or deterioration of the lubri-cating (Mittelbach, 1996; Ramos et al., 2009). FAMEs with higherdegree of unsaturation are not suitable for biodiesel. The CN, SVand IV can be predicted from fatty acid methyl esters of oils byusing empirical equations (Krisnangkura 1986; Kalayasiri et al.,1996). According to biodiesel standard EN 14214 methods, theconcentration of linolenic acid and acid containing four doublebonds in FAMEs should not exceed the limit of 12% and 1%, respec-tively. Higher oleic acid content increases the oxidative stability forlonger storage (Knothe, 2005) and decreases the cold filter plug-ging point (CFPP) for use in cold regions (Stournas et al., 1995).The pour and cloud points of feedstock decrease with increasing

Table 6Some of the target genes and the primers used for phylogenetic studies of microalgae.

Target region Primers used

18S rDNA 16S1 N/16S2 NChloroF/ChloroREK82F/Proto5R519F/1406R

Large subunit (LSU rDNA) FD8/RB, D1/D2Plastid rbcL –Small-subunit (SSU rDNA) 18ScomF1/Dino18SR

EK82f/Proto 5rMitochondrial cytochrome c oxidase subunit

(cox2–cox1/cox2–cox1coxF/coxR

Internal Transcribed Spacer (ITS) ITS1/ITS2ITS3/ITS4

ITS1+5.8S+ITS2Microsatellite locus Kbr1–Kbr10

chain length and branching of the alcohol moiety (Foglia et al.,1997). Biodiesel feedstock with a high degree of saturation is moreresistant to oxidation and more stable in presence of light, oxygen,high temperatures, and metals (Canakci and Sanli, 2008; Knothe,2005). The determination of fatty acid composition of algae oil isessential for assessing the fuel quality of biodiesel. Different cul-ture conditions play an important role in the lipid composition, likeC:N ratio, which is the major factor (Papanikolaou et al., 2004). Inorder to improve the fuel properties of algae biodiesel, suitable cul-ture conditions may be used and the properties of biodiesel pro-duced from algae oils must meet the International Biodiesel FuelStandards.

7. Microalgal identification strategies using molecular approach

Identification and enumeration of the algae of interest is a ma-jor challenge faced by researchers worldwide. Microscope-basedmicroalgal cell identification methods are usually the standardprocedures used in laboratories for the rapid screening of algalsamples. Conventional light microscopy has been extended to theuse of phase contrast microscopy, fluorescence microscopy, scan-ning electron microscopy (SEM) and transmission electron micros-copy (TEM) for species level identification. However, conventionalmicroscopy techniques used to analyse microalgae can give mis-leading results since they lack morphological markers for preciseidentification; further microalgae can often change cell size andshape during their life cycle (Godhe et al., 2002; Bertozzini et al.,2005). Mostly algae do not survive during fixation or they shrink,lose pigmentation and flagella so that a proper identification isimpossible. In addition, some species constitute very minor frac-tion of the total planktonic community, which leads to tediousanalysis of distinct samples. The identification of microalgae infield samples using microscopy is also time consuming and re-quires both experience and significant taxonomic and technicalskills (Godhe et al., 2002).

The molecular-based techniques developed in the last two dec-ades have led to rapid and accurate monitoring, identification andquantification of microalgae species in mixed phytoplankton sam-ples. Without complete DNA sequence, the analysis of small con-served genes has proved to be very helpful in the clarification ofthe relationships between algae. The most common DNA regionsanalysed today for phylogenetic purposes are ribosomal RNA genes(rRNA), mitochondria genes, plastid genes (rbcL), ITS (InternalTranscribed Sequences) and microsatellite DNA sequences (Table6). Ribosomal RNA’s are one of the widely used and are exception-ally useful for the comparative analysis of organisms (Rasoul-Ami-ni et al., 2009; Moro et al., 2009). They are very slowly alteringmolecules and major elements in the protein synthetic machineryof all cells. Small rRNA (SSU rRNA) genes are more highly con-

References

Tinti et al. (2007)Moro et al. (2009)Auinger et al. (2008)Rasoul-Amini et al. (2009)Scholin and Anderson (1996) and Kamikawa et al. (2007)Novis et al. (2009)

1, Richlen et al. (2007) and Auinger et al. (2008)

Kamikawa et al. (2007)

Coleman (2003, 2007) and Jürgen et al. (2008)

Kamikawa et al. (2007)Henrichs et al. (2007)


served than large rRNA (LSU rRNA) genes, and are therefore moreuseful for analysis of more distantly related species. Analysis ofthe LSU rRNA gene has been very useful for sorting out closely re-lated species concepts, like species groups and geographical originof different clonal isolates (Scholin and Anderson, 1996).

When using the microalgae for Polymerase Chain reaction (PCR)studies, the genomic DNA has to be extracted and purified fromsamples, removing potential inhibitors that often cause PCR inhibi-tion and low yields of PCR products (Godhe et al., 2002; Bertozziniet al., 2005). The efficiency of the genomic DNA extraction step isvery important for the subsequent PCR assay, especially when itis to be used in quantitative investigation on cultured or environ-mental samples. To overcome this, direct PCR amplification fromfew cells without the need for DNA extractions has been reported(Godhe et al., 2002; Auinger et al., 2008). Furthermore, in the sam-pling of long-term monitoring programmes the PCR is also appliedto preserved natural samples. Fixatives, such as Lugol’s solution,formalin and glutaraldehyde, are used as preserving agents forthe long term storage of phytoplankton material without destroy-ing the DNA needed as template in morphological and molecularidentification (Godhe et al., 2002; Auinger et al., 2008). Quantita-tive analysis of planktonic protists and microalgae from preservedfield samples combining morphological and small-subunit (SSU)rRNA gene sequence using single cell PCR has been reported re-cently (Auinger et al., 2008). A further promising molecular ap-proach is the application of DNA microarrays, which are appliedgenerally for gene expression and have been used with oligonu-cleotide probes of conserved genes for species identification at alltaxonomic levels (Gescher et al., 2008).The technique is based ona minimized but, high throughput form of a dot blot through appli-cation of sequences or probes in an ordered array on the chip. Thechip is made of glass and has special properties. The microarray of-fers the potential to facilitate the analysis of multiple targets fromone sample in one experiment (Gescher et al., 2008). A combina-tion of molecular probes/primers and DNA microarrays could serveas a rapid and reliable tool for rapid screening of microalgae. Theapplication of novel molecular techniques has the potential to rev-olutionise microalgal classification and especially identifying hy-per-lipid producing microalgae.

8. Microalgal maintenance

One of the major limitations with regards to culture mainte-nance is effective technology for long term storage of cultures(Moreno-Garrido, 2008). As living microorganisms, microalgalsamples must be stored under suitable controlled laboratory con-ditions so that they do not degenerate and lose viability. Biopro-specting for microalgal strains is costly and time consuming;therefore it is crucial to store the cultures properly so that the bio-prospecting exercise is not put to waste. The best way to storepurified microalgal cells is to have the cells in an aqueous systemunder suboptimal temperature and light regimens. According toLorenz et al. (2005) standard light intensities between 10–30 mmol photons m�2 s�1 have proved appropriate in combinationwith subdued temperatures for long-term culturing of most micro-algal taxa. Over-illumination of the stored microalgal culturesshould be avoided at all costs to prevent from photo-oxidativestress in microalgae and also the problem of localised heating.Light and dark photoperiods are required for the maintenance ofmost cultures with 12:12 and 16:8 h light: dark regimens recom-mended for a wide range of microalgal strains (Lorenz et al., 2005).

For long term storage, microalgal specimens should be kept atlow temperatures of ±2 �C but however most microalgal strainscan be kept at 15–20 �C. The stored microalgal cultures should beroutinely sub cultured to get metabolically active inocula and thisshould be done at least 1–3 weeks employing conventional aseptic

microbiological techniques to avoid contaminating the purified cul-tures. Generally it is difficult to establish storage conditions fornewly isolated microalgal strains and as a rule of thumb, it is recom-mended to do trial and error of different light intensities and tem-peratures (Lorenz et al., 2005). The viability of these cultures isthen tested periodically and a suitable maintenance regime is estab-lished for future application. It is prudent to appropriately label thestored cultures using waterproof labels and permanent ink and alsoto routinely check for contamination using light microscopy.

Media for each strain must be carefully chosen so that the mic-roalgal cells are not stressed to an extent of altering their morphol-ogy and development of deleterious effects. A wide range of mediaare available such as BBM, BG-11, ASW and so on for long termmaintenance of microalgae. The different media that can be usedfor long term maintenance of microalgae are described is Section4 and Table 4. These media types can support the growth of micro-algae from specific aquatic environments such as fresh water, mar-ine and brackish. The media chosen can either be in liquid or solidagar medium. However slants can prolong the viability of microal-gal cells in storage. Microalgae can be lyophilised and stored as apowder at 2 �C. In addition, cryopreservation of microalgal cellshas been found to be effective as a culture maintenance strategy(Day and Brand, 2005). The ability to routinely cryopreserve micro-algal species reduces costs associated with maintaining large cul-ture collections and reduces the risks of losing particular strainsor species through contamination and genetic drift (Rhodes et al.,2006).Currently microalgae from diverse aquatic environmentsare maintained by serially subculturing which is labor intensiveand therefore repeatedly exposes the culture to contaminationand handling error (Cox et al., 2009). To date, standard cryogenictechniques are documented as best methods for long term micro-algal storage (Cox et al., 2009).

9. Biofuel production

9.1. Exploring possibility of algae based biofuels

The algae oil and spent biomass are considered as potentialsources for biofuels production. Microalgal biomass can be pro-duced using either open raceway ponds or photobioreactors. Thepros and cons of these two systems are described in Table 7. Thebiodiesel produced from algae oil by using transesterification pro-cess can be used for replacing the conventional diesel fuel. The al-gae oil cake produced in algae biofuel industry is ca. of 1–2 times oflipid yield. For example, 1000 kg of algae biomass having 35% oilcontent can yield 650 kg of algae deoiled cake. Biomass EnergyConversion Technologies (BECT) can be used to convert the algaebiomass into different kinds of energy fuels. BECT is divided intothermo-chemical and biochemical methods. The BECT includescombustion (for heat energy), pyrolysis (pyrolytic gas, bio-oil, bio-char), gasification (syngas), thermo-chemical liquefaction (bio-crude oil), biomethanation (biogas), photobiological hydrogenproduction (hydrogen) and alcoholic fermentation (ethanol). Algaeoil can be converted into biodiesel by the transesterification pro-cess as depicted in Fig. 3a and b. The recent developments in re-search on algae biofuels production has been extensivelyinvestigated (Harun et al., 2010).

9.2. Biodiesel production

The viscosity of the raw microalgal oil is usually higher thanthat of diesel fuel. Raw oil can be converted into biodiesel bymeans of transesterification process. The transesterification reac-tion can be catalyzed by alkalis, acids, enzymes or supercriticalmethanol. Free fatty acid (FFA) content of algae is in the range of

Table 7Generalized comparisons of two different cultivation methods of algae production.

Factors Open ponds Photobioreactors

CultivationCultivation Multi strain

cultivationWell suitable for singlestrain cultivation

Contamination High Less to noneCleaning None Required due to wall

growth and dirtControlling of growth

conditionsVery difficult Easy

Temperature Highly variable Required coolingAutomatic cooling

systemNone Built in

Automatic heatingsystem

None Built in

Microbiology safety None UV

Biomass productionBiomass quality Variable ReproducibleBiomass productivity Low HighLipid productivity Low HighLight utilization

efficiencyLow High

Operational modeAir pump Built in Built inShear Low HighCO2 transfer rate Poor ExcellentMixing efficiency Poor ExcellentWater loss Very high LowEvaporation High No evaporationO2 concentration Low due to

continuousspontaneous outgassing

Build-up occurredrequires gasexchange device

CO2 loss High, dependingon pond depth

Low

EconomicsSpace required High LowPeriodical maintenance Less MoreCapital investment Low HighOperating cost Lower HigherHarvesting cost High LowerScale up technology for

commercial levelEasy to scale up Most of photobioreactor

models are difficult to scaleup due to limitations

Modified from Pulz (2001) and Harun et al. (2010).

(b)

(a)

Deoiled cake

Algae cultivation

Drying of algae

Harvesting of algae

Dried algae

Oil extraction

Algae oil

Transesterification reactor

Algae biodiesel Glycerol

Fig. 3. Schematic representation of biodiesel production from microalgae oil (a)and Biodiesel production by transesterification reaction (b).


20–50% (Kosaric and Velikonja, 1995; Mansour et al., 2005), whichis the main factor for formation of soap in alkaline catalyst basedtransesterfication process and separation of biodiesel and glycerolis difficult. Due to these problems, the alkaline catalyst based bio-diesel production is not suitable for algae oil with high FFA content.For transesterification of high free fatty acid feedstocks, acid cata-lysts were found to be useful for pre-treating, but its reaction con-version rates are very slow (Um and Kim, 2009). Enzymes are alsoused for feedstock with higher free fatty acid, but these catalystsare expensive and unable to provide the degree of reactioncompletion required to meet the American Society for Testingand Materials (ASTM) fuel specification. The use of supercriticaltransesterification process method for algae biodiesel productionis also restricted due to process economics and safety concerns re-lated to the reaction conditions (Ehimen et al., 2010). Ehimen andco-workers (2010) studied the effect of operational parameterssuch as alcohol volume, moisture content, temperature, reactiontime, and mixing on an acid-catalyzed in situ transesterificationprocess for production of biodiesel from microalgae lipids. Theyfound that water content in the algae biomass was greater than115% w/w (based on oil weight) and this inhibited the in situtransesterification reaction. The Mcgyan process is continuoustransesterification process for biodiesel production from variousfeedstocks. In this process, the combination of alcohol and lipid

is sent into a continuous fixed-bed reactor filled with a sulfatedmetal oxide catalyst at elevated temperature and pressure to per-form the transesterification and esterification reactions simulta-neously. Added advantages of this technology are: no catalyst isused, less reaction time and uses no water or dangerous chemicals.(Um and Kim, 2009). Adapting a biorefinery approach i.e. utilisingthe oil for biodiesel, spent biomass, glycerol, etc. as value addedproducts has certainly made the technology attractive and there-fore attracted much attention in the field.

9.3. Thermo-chemical and biochemical conversion

Thermo-chemical conversion process involves thermal decom-position of organic components present in algae biomass into dif-ferent kind of energy fuels. The end products may vary from gasor liquid or solid fuel depending on temperature used in the ther-mo-chemical conversion process. The thermal decomposition oforganic components can be achieved by different processes suchas direct combustion, gasification, thermo-chemical liquefaction,and pyrolysis. Gasification is the conversion of biomass into a com-bustible gas mixture by the partial oxidation of biomass at hightemperatures, typically in the range 800–900 �C (McKendry,2002) and the produced syngas can be used for thermal applica-tions or fuel for diesel/gas turbine engines. Hirano et al. (1998)studied gasification of Spirulina at temperatures ranging from850 to 1000oC for methanol production. They estimated that algaebiomass gasification at 1000oC produced the highest theoreticalyield of 0.64 g methanol from 1 g of algae biomass.


Thermo-chemical liquefaction is a process that can be em-ployed to convert wet algal biomass material into liquid fuel.Added advantages of this technology are conversion of wet bio-mass into useful energy and no feedstock drying process involved(Clark and Deswarte, 2008). Dote et al.(1994) found that thermo-chemical liquefaction of microalgae species like Botryococcus brau-nii, Dunaliella tertiolecta and Spirulina platensis yielded 64, 42 and30–48% dry wt. basis of oil and fuel properties of biocrude oil(30–45.9 MJ/kg) which was close to that of petroleum based heavyoil (42 MJ/kg) (Jena and Das, 2009).

Pyrolysis is the thermal decomposition of materials in the ab-sence of oxygen or when significantly less oxygen is present thanrequired for complete combustion (Balat et al., 2009). Slow pyroly-sis of biomass is associated with high charcoal content (350–700 �C), but the fast pyrolysis is associated with liquid fuels, atlow temperature (675–775 K) (Bridgwater, 2003), and/or gas, athigh temperature (Encinar et al., 1998). Fast pyrolysis process forconversion of algae biomass is used to produce 60–75 wt.% of li-quid bio-oil, 15–25 wt.% of solid char, and 10–20 wt.% of non con-densable gases, depending on the feedstock used (Mohan et al.,2006). Miao et al. (2004) studied fast pyrolysis of Chlorella prototh-ecoides and Microcystis aeruginosa grown phototrophically. They re-ported that bio-oil yields of 18% (higher heating value [HHV] of30 MJ kg�1) and 24% (HHV of 29 MJ kg�1) for microalgae C. proth-othecoides and Microcystis aeruginosa, respectively. Researchershave also reported using anaerobic digestion of microalgae as anecessary step to make microalgal biodiesel sustainable (Sialveet al., 2009). The chemical composition of microalgae shows varia-tion according to species, seasons and habitats (Rupérez, 2002).Due to high protein content of microalgae, increased ammonia isproduced upon microbial protein degradation, which inhibit anaer-obic microorganisms in anaerobic digestion. The overall perfor-mance of anaerobic digestion system is affected due to low C:Nratio of the microalgae, but it can be improved by co-digestionwith a high C:N ratio material (Brennan and Owende, 2010). Phys-ico-chemical pre-treatment and co-digestion are strategies thatcan significantly and efficiently increase the conversion yield ofthe algal organic matter into methane.

10. Conclusions

Due to the increasing global appetite for renewable liquidfuels, the need for developing suitable technology to meet thisdemand is imperative. Comparative analysis of producing oilfrom algae and crop based biomass show the former to be moreadvantageous on various aspects. However, conducting an exten-sive search of environments for unique hyper-lipid producers iscrucial to the success of the technology. The application of suit-able and efficient technology for sampling, isolation and screen-ing and culture maintenance is important. Identification ofselected strains using novel molecular techniques is more accu-rate than current conventional methods. Producing the rightquality and quantity of oil and FAME product will largely deter-mine the techno-economics of the technology. In addition ther-mo-chemical and biochemical conversion process can improvethe economics. Most of the literature to date has largely focusedon downstream processing of oil to biodiesel. As shown in thecurrent communication, the upstream processing of biomass tomaximize the downstream yields is crucial for the overall successof the technology.

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