Microalgal lipids biochemistry and biotechnological perspectives

Biotechnology Advances 32 (2014) 1476–1493

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Research review paper

Microalgal lipids biochemistry and biotechnological perspectives

Stamatia Bellou a, Mohammed N. Baeshen b, Ahmed M. Elazzazy b,c, Dimitra Aggeli d,Fotoon Sayegh b, George Aggelis a,b,⁎a Division of Genetics, Cell & Development Biology, Department of Biology, University of Patras, Patras 26504, Greeceb Department of Biological Sciences, King Abdulaziz University, Jeddah 21589, Saudi Arabiac Department of Chemistry of Natural and Microbial Products, National Research Centre, Dokki 12622, Giza, Egyptd Department of Genetics, Stanford University, Stanford, CA 94305, USA

⁎ Corresponding author at: Division of Genetics, Cell &E-mail address: [email protected] (G. Aggelis

http://dx.doi.org/10.1016/j.biotechadv.2014.10.0030734-9750/© 2014 Elsevier Inc. All rights reserved.

a b s t r a c t
a r t i c l e i n f o
Article history:Received 15 June 2014Received in revised form 2 October 2014Accepted 6 October 2014Available online 14 October 2014

Keywords:MicroalgaeLipid biosynthesisGenetic engineeringPolyunsaturated fatty acidsBiodieselPigmentsProteinsWastewater treatment

In the last few years, there has been an intense interest in using microalgal lipids in food, chemical and pharma-ceutical industries and cosmetology, while a noteworthy research has been performed focusing on all aspects ofmicroalgal lipid production. This includes basic research on the pathways of solar energy conversion and on lipidbiosynthesis and catabolism, and applied research dealing with the various biological and technical bottlenecksof the lipid production process. In here, we review the current knowledge in microalgal lipids with respect totheir metabolism and various biotechnological applications, and we discuss potential future perspectives.The committing step in fatty acid biosynthesis is the carboxylation of acetyl-CoA to formmalonyl-CoA that is thenintroduced in the fatty acid synthesis cycle leading to the formation of palmitic and stearic acids. Oleic acid mayalso be synthesized after stearic acid desaturation while further conversions of the fatty acids (i.e. desaturations,elongations) occur after their esterification with structural lipids of both plastids and the endoplasmic reticulum.The aliphatic chains are also used as building blocks for structuring storage acylglycerols via the Kennedy path-way. Current research, aiming to enhance lipogenesis in the microalgal cell, is focusing on over-expressingkey-enzymes involved in the earlier steps of the pathway of fatty acid synthesis. A complementary plan wouldbe the repression of lipid catabolism by down-regulating acylglycerol hydrolysis and/orβ-oxidation. The tenden-cy of oleaginousmicroalgae to synthesize, apart from lipids, significant amounts of other energy-rich compoundssuch as sugars, in processes competitive to lipogenesis, deserves attention since the lipid yield may be consider-ably increased by blocking competitive metabolic pathways.Themajority of microalgal production occurs in outdoor cultivation and for this reason biotechnological applica-tions face some difficulties. Therefore, algal production systems need to be improved and harvesting systemsneed to bemore effective in order for their industrial applications to becomemore competitive and economicallyviable. Besides, a reduction of the production cost ofmicroalgal lipids can be achieved by combining lipid produc-tion with other commercial applications. The combined production of bioactive products and lipids, whenpossible, can support the commercial viability of both processes. Hydrophobic compounds can be extractedsimultaneously with lipids and then purified, while hydrophilic compounds such as proteins and sugars maybe extracted from the defatted biomass. The microalgae also have applications in environmental biotechnologysince they can be used for bioremediation of wastewater and tomonitor environmental toxicants. Algal biomassproduced during wastewater treatment may be further valorized in the biofuel manufacture.It is anticipated that the high microalgal lipid potential will force research towards finding effective ways tomanipulate biochemical pathways involved in lipid biosynthesis and towards cost effective algal cultivationand harvesting systems, as well.

© 2014 Elsevier Inc. All rights reserved.

Contents

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1477Microalgal lipids in the forefront of lipid biotechnology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1477Lipid metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1478

Development Biology, Department of Biology, University of Patras, Patras 26504, Greece.).

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1477S. Bellou et al. / Biotechnology Advances 32 (2014) 1476–1493

Lipid biosynthesis and turnover . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1478PUFA biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1480Genetic engineering for directing metabolism towards lipid synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1480

Biotechnological perspectives of microalgal lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1481Occurrence of PUFAs in microalgal lipids and industrial applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1482Microalgal lipids as feedstock for biodiesel production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1485Combined production of microalgal lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1486

Combined production of lipids and pharmaceutical products . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1486Lipids as co-products of environmental applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1487

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1488Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1489References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1489

Introduction

The microalgae are photosynthetic microorganisms that play a keyrole in the natural ecosystems supplying organic matter and specificmolecules, such as polyunsaturated fatty acids (PUFAs), to many higherorganisms. Microalgae applications range from human and animalnutrition to cosmetics and the production of high value molecules.

The systematic development of microalgal applications started inthe 20th century. Several species are currently cultivated in largescale, in artificial or natural ponds and rarely in photobioreactors(PBRs), producing up to some tons of biomass and/or various metabo-lites per year. Algal biomass is rich in PUFAs, minerals (e.g. Na, K, Ca,Mg, Fe, Zn and traceminerals) and vitamins, such as riboflavin, thiamin,carotene and folic acid and so on (Becker, 2004; García-Garibay et al.,2003; Samarakoona and Jeona, 2012). Themicroalgal biomass, especial-ly that produced by the species Dunaliella, Arthrospira (Spirulina, cyano-bacterium) and Chlorella, is alreadymarketed in various forms designedfor human nutrition or is incorporated into foods and beverages (Lianget al., 2004; Yamaguchi, 1997), as it is considered a healthy nutritionalsupplement (Apt and Behrens, 1999; Borowitzka, 1999; Jensen et al.,2001; Priyadarshani and Rath, 2012; Soletto et al., 2005). Similarly, theconsumption of even small amounts of microalgal biomass canpositively affect the physiology of animals by improving immuneresponse, diseases' resistance, antiviral and antibacterial protection,improved gut function, probiotic colonization stimulation, aswell as en-hanced feed conversion, reproductive performance and weight control(Harel and Clayton, 2004). Although thequality of algal proteins lags be-hind that of animal proteins, it is superior to that of common plants(Barrow and Shahidi, 2008; Becker, 2004; Kay and Barton, 1991;Samarakoona and Jeona, 2012; Sydney et al., 2010; Um and Kim,2009). Particular algal peptides, such as taurine, are of great nutritionaland pharmaceutical interest (Houstan, 2005), while glycoproteins(lectins), extracted bymarine algae, are considered a type of interestingproteins for biochemical and clinical research, and can be isolated withtheir carbohydrate moiety (Silva et al., 2010).

Some species contain considerable amounts of pigments thatare used in cosmetics and as natural coloring agents. Many industrialproduction plants are established in China, Australia and USA (Brownet al., 1997; García-González et al., 2005; León et al., 2003) dealingwith beta-carotene production (e.g. from Dunaliella salina) that isused as a food coloring (Metting, 1996). Other pigments such asphycobiliproteins have been extracted from various marine algaeincluding Porphyridium cruentum and Synechococcus spp. (Viskari andColyer, 2003).

Although the majority of applications concern biomass productiondestined for animal or human consumption— in fact, 30% of the currentworld algal production is sold for animal feed applications (Becker,2004) — there has been an increased interest in the use of microalgallipids in numerous commercial applications, such as in food, chemicaland pharmaceutical industries and cosmetology. Indicative of the highinterest in microalgal lipids is the noteworthy research that has been

performed in the last decade on all aspects concerning microalgal lipidproduction. These include fundamental research on the mechanismsused for light energy conversion and on lipid biosynthesis and catabo-lism, as well as biotechnological research dealing with the various tech-nical bottlenecks of the lipid production process. In the current reviewarticle the up-to-date level of knowledge in lipid biosynthesis and turn-over inmicroalgae and the various biotechnological applications and fu-ture perspectives of microalgal lipids are comprehensively presentedand discussed. Τaking into consideration the recent techno-economicanalyses concluding that the algal lipid content is the most criticalfactor affecting the viability of large-scale applications, especiallythose related to biodiesel, the current research efforts aimed to reinforcealgal liposynthetic machinery using genetic engineering are alsodiscussed.

Microalgal lipids in the forefront of lipid biotechnology

Several microalgal species are able to accumulate appreciable lipidquantities, and therefore are characterized as oleaginous. Lipid contentin microalgae can reach up to 80% in dry biomass, but even in thesecases the lipid productivity is actually low. Inwidespread species belong-ing to the genera of Porphyridium, Dunaliella, Isochrysis, Nannochloropsis,Tetraselmis, Phaeodactylum, Chlorella and Schizochytrium, lipid contentvaries between 20 and 50%. However, higher lipid accumulation can bereached by varying the culture conditions. Factors such as temperature,irradiance and, mostly, nutrient availability have been shown to affectlipid content and composition in algal cells.

The interest for algal lipid arises mainly from the fact that theseorganisms are able to synthesize considerable quantities of PUFAs thateither reach humans via the food chain or are used as food supplements(Fig. 1). Indeed microalgae are the primary source of PUFAs havingnutritional and pharmaceutical interest (Doughman et al., 2007; Kyle,2001). Although fish are also a source of PUFAs, these organisms usuallyobtain their PUFAs via bioaccumulation through the food chain(Benemann et al., 1987). Furthermore, fish PUFA production dependson fish quality and sufficiency while that of algae does not.

Several microalgae are able to synthesize omega-3-long chainPUFAs, at levels over 20% of their total lipids. In algal cell PUFAs are es-terified with an alcohol, usually glycerol, generating triacylglycerols(TAGs) or polar lipids (i.e. phospholipids, glycolipids) of exceptionalstructure regulating membranes fluidity and function. Depending onthe strain, lipid industrial production can be combinedwith the produc-tion of other metabolic products of high value, such as beta-caroteneand astaxanthine. The main drawback in using microalgae to producePUFA rich lipid in large scale is the low lipid content in algal celland the low biomass density in the reactor, usually not exceeding400–600mg/L under industrial culture conditions, which increases con-siderably the harvesting cost.

Alternatively, microalgal lipids represent an attractive source of oil,suitable as feedstock for biodiesel production. Actually, microalgaeoffer a number of advantages from an industrial perspective. These

Fig. 1. Microalgal biomass production and transfer of polyunsaturated fatty acids (PUFAs) to man, either through food chain or microalgal supplement consumption.The individual photos are kindly provided by PLAGTON S.A. (Mytikas, Greece), Kefish (Kefalonia, Greece) and ALGAE A.C. (Nigrita Serres, Greece).

1478 S. Bellou et al. / Biotechnology Advances 32 (2014) 1476–1493

include simplicity of culture, increased photosynthetic efficiency andgrowth rates, higher biomass and oil productivities compared to terres-trial plants, and higher rates of CO2 fixation and O2 release. The finalalgal oil production per unit of surface may reach 200 times that ofcommon plant oils (Chisti, 2007). However, biodiesel produced byalgae is more expensive than that produced by conventional plants,while fossil fuel is much cheaper.

Lipid metabolism

Most microalgae accumulate lipids under specific environmentalstress conditions, such as nitrogen or phosphate limitation (Amaroet al., 2011; Bellou and Aggelis, 2012; Courchesne et al., 2009; Huet al., 2008, 2013; Msanne et al., 2012). Therefore the management ofthe environmental conditions is a common approach used for improv-ing lipid accumulation in themicroalgal cell. Strain selection is also like-ly to be of critical importance. Recently, there has been an intenseinterest in isolating new native microalgal strains when a large-scaleapplication is intended. Microalgae are exposed to a variety of changesin the environment. Seasonal cycles vary according to the climatic andgeographical location of the habitat, in which they are growing, anddifferent strains of the same speciesmay responddifferently. Strains de-rived from diverse geographical locations are physiologically different(Bouaicha et al., 2001; Delgado et al., 1991), and could vary in their re-sponse to any limiting environmental factor. Consequently indigenousstrains may have developed mechanisms for sensing and acclimatingto changes in their environment, and thus their use promises higherpotentiality (Vonshak and Torzillo, 2004).

Understanding lipid metabolism and how it is controlled duringalgal growth is of great importance for maximizing lipid production.Despite the significant biotechnological applications, microalgae have

not been fully studied in terms of their biochemistry. Therefore, fattyacid biosynthesis and modification (both elongation and desaturation)and lipid catabolism have not been clarified for microalgae as theyhave for plants and heterotrophic microorganisms (Beer et al., 2009;Boyle et al., 2012; Khozin-Goldberg and Cohen, 2011; Moelleringet al., 2010;Moseley et al., 2009). Especially, the overall lipid biosynthe-sis pathway and its regulators have not been clearly described. Efforts toimprove lipid accumulation in microalgae by modifying the expressionof key enzymes implicated in lipid synthesis often fail, suggesting a lackin our understanding of the mechanisms that govern lipid accumula-tion, if not algal cell biology.

Lipid biosynthesis and turnover

Through photosynthesis CO2 is converted to glycerate-3-phosphate(G3P). This molecule is the precursor of several storage materials,such as polysaccharides and lipids. The conversion of G3P to pyruvateand thereafter to acetyl-CoA, via a reaction catalyzed by the pyruvatedehydrogenase complex (PDC), initiates the lipid biosynthetic pathway,which occurs in the plastid. Acetyl-CoA can also be generated via a bio-chemical pathway that permits the conversion of polysaccharides intolipids (Bellou and Aggelis, 2012), pathway that is commonly utilizedby oleaginous heterotrophs during sugar assimilation (Bellou et al.,2012, 2014a,b; Chatzifragkou et al., 2010; Fakas et al., 2008; Gemaet al., 2002; Makri et al., 2010; Papanikolaou and Aggelis, 2011;Papanikolaou et al., 2004; Ratledge, 2004) (Fig. 2). Specifically,the breakdown of storage polysaccharides (occurring e.g. under lightlimitation) usually generates energy through glycolysis occurring inthe cytosol followed by the citric acid cycle occurring in the mitochon-drion. However, several environmental stresses, such as nitrogen orphosphate limitation, may disturb the citric acid cycle (i.e. by inhibiting

Fig. 2. A simplified scheme showing lipid synthesis in microalgae. For details see text (from Radakovits et al., 2010; Bellou and Aggelis, 2012; Chen and Smith, 2012; Hu et al., 2013, allmodified). Abbreviations: ACC, acetyl-CoA carboxylase; ACP, acyl-carrier protein; LACS, long-chain acyl-CoA synthetase; ATP:CL, ATP-dependent citrate lyase; CoA, coenzyme A; DGAT,diacylglycerol acyltransferase; ER, endoplasmic reticulum; FAS, fatty acid synthase; FAT, fatty acyl-ACP thioesterase; G3P, glycerate-3-phosphate; GPAT, glycerol-3-phosphate acyltransferase;KAS, 3-ketoacyl-ACP synthase; LPAAT, lyso-phosphatidic acid acyltransferase; LPAT, lyso-phosphatidylcholine acyltransferase; PDC, pyruvate dehydrogenase complex; TAG, triacylglycerol.


NAD+-isocitrate dehydrogenase) leading to citrate accumulation in themitochondrion and subsequently to its excretion in the cytosol. Cytosol-ic ATP-dependent citrate lyase converts citrate into oxaloacetate andacetyl-CoA; the latter is converted into malonyl-CoA by cytosolicacetyl-CoA carboxylase (ACC) and becomes available for fatty acidelongation in the membranes of the endoplasmic reticulum (ER) —

see below (Mühlroth et al., 2013). Although this mechanism has onlybeen shown in Nannochloropsis salina and Chlorella sp. cultivated in alab-scale open pond-simulating PBR (Bellou and Aggelis, 2012), it isprobably common in oleaginous strains that are able to growheterotrophically.

Despite the importance of acetyl-CoA biosynthesis, the committingstep in fatty acid biosynthesis is the carboxylation of acetyl-CoA toform malonyl-CoA, reaction catalyzed by the ACCs located either inthe plastid or in the cytosol (Baba and Shiraiwa, 2013; Davis et al.,2000; Khozin-Goldberg and Cohen, 2011; Kim, 1997; Lei et al.,2012) (Fig. 2). In algae, ACCs exist in two different forms, theheteromeric and homomeric. Although it is generally believed thatthe heteromeric form is the one present in algal plastids,Huerlimann and Heimann (2013) reported that this is not the casefor all algae, since the presence of heteromeric or homomeric ACCsis dependent on the origin of plastid. I.e. in Chlorophyta (except forPrasinophyceae) and Rhodophyta, heteromeric ACCs have beenfound in their plastids, whereas Heterokontophyta, Haptophytaand Apicomplexa contain homomeric ACCs in their plastids. Inseveral microalgal strains, i.e. those belonging to Galdiera sulpuraria,Cyanidioschyzon merolae, Thalassiosira pseudonana and Phaeodactylumtricornutum, two ACCs have been identified, the plastidial ACC1 andthe cytosolic ACC2 (Khozin-Goldberg and Cohen, 2011; Huerlimannand Heimann, 2013). Two genes coding for ACC have been also foundin Nannochloropsis gaditana (Radakovits et al., 2012).

In the plastid, the malonyl-CoA is transferred to the acyl-carrierprotein (ACP), which is one of the subunits of the fatty acid synthase

(FAS) complex, by the malonyl-CoA:ACP transacetylase (Blatti et al.,2012; Greenwell et al., 2010; Hu et al., 2008; Subrahmanyam andCronan, 1998). Themalonyl-ACP is introduced in the fatty acid synthesiscycle through the 3-ketoacyl-ACP synthase (KAS). The KAS catalyzes thecondensation of an acetyl groupwithmalonyl-ACP to form ketobutyryl-ACP. This compound is converted via the sequential reactions ofreduction–dehydration–reduction to butyryl-ACP and the cycle isrepeated until the formation of palmitoyl-ACP. The latter is then con-verted into stearoyl-ACP after the addition of only two carbonmoleculesoriginated from acetyl-CoA. Oleoyl-ACP is also synthesized afterdesaturation of stearoyl-ACP, reaction mediated by the plastidial Δ9desaturase (Mühlroth et al., 2013; W.L. Yu et al., 2011). Principally, thefatty acids are released from ACP by a fatty acyl-ACP thioesterase(FAT), located in the chloroplast envelope. They are activated thereafterinto acyl-CoA by the long-chain acyl-CoA synthetase (LACS), also locat-ed in the chloroplast envelope, and transferred in the cytosol, wherethey become available for lipid synthesis. Alternatively, in plants, andprobably in microalgae, the acyl chainsmay be used for structural lipids(mostly glycolipids) synthesis in the plastid. For this purpose theyare transferred from ACP either to G3P or to monoacylglycerol-3-phosphate via the action of a plastidial acyltransferase (Ohlrogge andBrowse, 1995). All three enzymes ACP, KAS and FAT, have been shownto play an essential role in fatty acid synthesis inHaematococcus pluvialis(Lei et al., 2012). Those transferred in the cytosol acyl-CoA chains are es-terified with structural phospholipids of the ER to be converted intohigher derivatives (PUFAs). The modified or not in the ER fatty acidsare used as building blocks for the formation of TAGs via the Kennedypathway. Those implicated in the Kennedy pathway acyltransferases(i.e. diacylglycerol acyltransferase — DGAT, glycerol-3-phosphateacyltransferase — GPAT, lyso-phosphatidic acid acyltransferase —

LPAAT, lyso-phosphatidylcholine acyltransferase — LPAT) are alsolocated in the ER (Chen and Smith, 2012; Liu and Benning, 2013;Merchant et al., 2012; Radakovits et al., 2010).

image of Fig.�2


As reported by Chen and Smith (2012), DGAT is found in algae intwo isoforms (DGAT1 and DGAT2) catalyzing the same reaction butwith significant variations in sequence. Although the genes encodingfor DGATs have been found in various microalgae, such as Ostreococcustauri, T. pseudonana, Nannochloropsis sp., and the model organismChlamydomonas reinhardtii, their function has not been clearly character-ized and thus it needs further studying (Khozin-Goldberg and Cohen,2011). Recently, Wang et al. (2014) reported that in Nannochloropsisspecies 1 or 2 DGAT1 and 11 DGAT2 gene doses exist, while only6 and 4 DGATs genes respectively were found in C. reinhardtiiand T. pseudonana and even fewer in some other green algae andheterokonts. N. gaditana genome analysis showed common homologuesof ACC, KAS, glycerol-3-phosphate-dehydrogenase (G3PDH), GPAT andLPAAT genes that are also found in other microalgal strains, i.e. thebrown alga Ectocarpus siliculosus, the diatom P. tricornutum, the redalga C. merolae and the green alga C. reinhardtii (Radakovits et al.,2012). LPAT gene identified in T. pseudonana is 27% homologous to theLPAT of the yeast Saccharomyces cerevisiae (Chen et al., 2010).

In oleaginous yeasts and fungi storage lipid turnover typically occursafter the depletion of the carbon source in the culture medium or, ingeneral, under carbon-limiting conditions (Aggelis and Sourdis, 1997;Fakas et al., 2007; Holdsworth and Ratledge, 1988; Papanikolaou et al.,2004). Respectively, in microalgae under light starvation conditions,storage material (both sugar and lipid) is degraded to support algalgrowth, although a conversion of sugar to lipidswas observed in the be-ginning of the degradation process in Chlorella sp. (Bellou and Aggelis,2012).

PUFA biosynthesis

Τhe synthesis of long chain unsaturated fatty acids requires the pres-ence of specific elongases and desaturases, which act primarily onpalmitic, stearic and oleic acids. Fatty acid elongation occurs in bothplastids and ER (Kunst and Samuels, 2009; Ohlrogge and Browse,1995) and requires acyl-CoA and malonyl-CoA as substrates plus 1ATP and 2 NADPH molecules per C2-unit elongation of the carbonchain. Fatty acid elongase is a complex that consisted of four subunitsnamely β-ketoacyl-CoA synthase (KCS), β-ketoacyl-CoA reductase,β-hydroxyacyl-CoA dehydratase and enoyl-CoA reductase, which aresimilar to those found in type II FAS identified in C. reinhardtii (Babaand Shiraiwa, 2013; W.L. Yu et al., 2011). KCSs are divided into the“elongase of very long-chain fatty acid” (ELOVL) contributing tosphingolipid biosynthesis and the “fatty acid elongase” (FAE) workingfor TAGs or wax biosynthesis (Khozin-Goldberg and Cohen, 2011;Venegas-Calerón et al., 2010). In the plastid, the fatty acids are used toproduce lysophosphatidic acid and phosphatidic acid (PA) via the actionof plastidial acyltransferases. The PA and its derivative product diacyl-glycerol (DAG) may act as precursors for the synthesis of plastidialmembrane structural lipids (Fan et al., 2011; Ohlrogge and Browse,1995). Instead, the fatty acids are transferred to the ER and used to acyl-ate G3P, reaction mediated by ER-localized acyltransferase isoforms. Incontrast to those produced in plastids, the PA and DAG produced inthe ER are used for the synthesis of bothmembrane and storage (mainlyTAGs) lipids (Ohlrogge and Browse, 1995).

The two types of elongases are referred to as ELOVL, commonlyfound in yeast, fungi and animal cells, and as FAE, found in plants. Forthe synthesis of the very long-chain PUFAs, such as arachidonic (ARA),eicosapentaenoic (EPA) and docosahexaenoic (DHA) acids, which arecommon in the majority of marine species including microalgae suchas P. tricornutum, N. salina, N. gaditana, Isochrysis galbana, Pavlova salina,and Tetraselmis sp., the ELOVL is required, whereas FAE activity has notbeen shown in microalgae so far (Baba and Shiraiwa, 2013). However,Ouyang et al. (2013) suggest that a FAE may exist in the microalgaMyrmecia incise, the active region of which is exposed in the cytosolicside of the ER membrane, which may explain why arachidonic acid issynthesized in the cytoplasm instead of the chloroplast as it was

previously demonstrated by Bigogno et al. (2002). Several elongase-encoding genes implicated in PUFA synthesis have been characterizedin various species, such as in Pavlova lutheri (Pereira et al., 2004) andPyramimonas cordata (Petrie et al., 2010a).

Desaturases are specialized in the location, number and stereochem-istry of double bonds in fatty acids (Heinz, 1993; Pereira et al., 2003). Theimplication of the desaturases in the biosynthesis of the very long chainPUFAs has been extensively reviewed not only in heterotrophicmicroor-ganisms but in microalgae as well (Certik and Shimizu, 1999; Guschinaand Harwood, 2006; Harwood and Guschina, 2009; Khozin-Goldbergand Cohen, 2011; Moellering et al., 2010; Pereira et al., 2003; Walliset al., 2002). The genes encoding for Δ4, Δ5 and Δ6 desaturases,implicated in DHA synthesis, have been characterized in T. pseudonana(TpDESI, TpdESO and TpDESK) (Tonon et al., 2005). Similarly, a newdesaturase-encoding gene (IgD4), responsible for the conversion ofdocosapentaenoic acid (DPA) into DHA and of adrenic acid into DPA,was found in Isochrysis strains. Genes PsD4Des, PsD5Des and PsD8Deswere also identified in P. salina and were shown to encode for Δ4, Δ5and Δ8 desaturases, respectively (Zhou et al., 2007). A Δ6 desaturase-encoding gene has been found in Ostreococcus lucimarinus (Petrie et al.,2010a), while PiELO1 was characterized in the freshwater microalgaParietochloris incise and found to be functionally similar to Δ6 PUFAelongase-encoding genes of other species (Iskandarov et al., 2009). Iden-tification of Δ6 and Δ4 desaturase-encoding genes from OstreococcusRCC809 and Δ6 elongase-encoding gene from Fragilariopsis cylindruswas also reported by Vaezi et al. (2013), while Zäuner et al. (2012)identified in Chlamydomonas a gene encoding for Δ4 desaturase.

Genetic engineering for directing metabolism towards lipid synthesis

Many techno-economic analyses suggest that themost critical factoraffecting the production cost of microalgal lipid in both open pond andPBR systems is the lipid content in algal cells followed by the specificgrowth rate (Davis et al., 2011). For instance, in the open pond case anincrease of the lipid content in the algal cell from 25 to 50% results ina cost reduction of 4$/gal, which corresponds to 46% of the total produc-tion cost. On the other hand, a decrease of the lipid content from 25 to12.5% increases the production cost 8$/gal (around 100%). The effectof the lipid content on the production cost becomes more dramaticwhen a PBR is used instead of an open pond. The specific growth ratealso considerably affects the lipid production cost but in a lesser extentthan the lipid content (Davis et al., 2011). Therefore, it is not surprisingthe amount of effort focusing on the construction of microalgal strainsable to grow fast and synthesize large amounts of lipids having a suit-able fatty acid composition.

For improving growth rate, efforts have been focused on theconstruction of strains with an enhanced photosynthetic efficiency(see review of Stephenson et al., 2011). Although photosyntheticefficiency obviously affects lipid synthesis as well, particular strategieshave been developed to improve the liposynthetic machinery (Qinet al., 2012). These strategies target the overexpression of proteinsthat are involved in the earlier steps of fatty acid synthesis, increasingin this way the availability of precursor molecules, such as acetyl-CoAand malonyl-CoA. For example, increased ACC expression may stimu-late lipid synthesis. A complementary plan would be the repression oflipid catabolism by down-regulating or inhibiting TAG hydrolysisand/or β-oxidation process. Besides, the regulation and/or insertion inmicroalgae of specific desaturases and/or elongases, along with the as-sociated FATs, is used to modify fatty acid profile but, interestingly,may also affect lipid content or the biosynthesis of particular lipidfractions.

Genetic engineering approaches in microalgae are in their infancyand, consequently, the initial efforts had relatively low successindicating that a better understanding of the algal fatty acid biosyntheticmachinery is required (Blatti et al., 2013). Actually, despite the note-worthy research, the results were not always auspicious. For example,


overexpression of the plastidial ACC gene (ACC1) in the diatomsCyclotella cryptica and Navicula saprophila did not improve fatty acidsynthesis, indicating that ACC upregulation alone may not be sufficientandmore than one enzymemay act synergistically to boost lipid biosyn-thesis (Dunahay et al., 1996; Mühlroth et al., 2013). Another obvioustarget for improving lipid biosynthesis is to manipulate FAS that isACP-dependent (Sirevag and Levine, 1972). Therefore, away to enhancelipid biosynthesis is to focus on protein–protein interactions, i.e. be-tween the ACP and FAT, as Radakovits et al. (2011) and Blatti et al.(2012) reported. However, the findings showed that FAS affected fattyacid composition but not lipid accumulation, which indicate that theknowledge on FAS manipulation for enhanced lipid production is inthe initial stage.

Considering that the overexpression of genes involved in fatty acidsynthesis had low success, research was focused on other enzymesimplicated in acylglycerols (both storage and structural) biosynthesis.Nevertheless, overexpression of DGAT genes (CrDGAT2a, CrDGAT2b,and CrDGAT2c) in the model microalga C. reinhardtii did not lead to in-creased lipid accumulation (La Russa et al., 2012). Deng et al. (2012)studying five CrDGAT2 homologous genes in C. reinhardtii, found thateach gene affects diversely lipid accumulation pattern. Specifically,RNAi silencing of CrDGAT2-1 or CrDGAT2-5 resulted in a significant de-crease in lipid content, whereas transformants harboring CrDGAT2-4exhibited increase in lipid content. No significant changes in lipidcontent were observed when CrDGAT2-2 or CrDGAT2-3 was silenced.On the contrary, overexpression of a type 2 DGAT in P. tricornutum in-creased TAG accumulation by 35% (Niu et al., 2013). Concerning the ef-fect of heterologous expression of other implicated in lipogenesisenzymes in Chlorella minutissima, such as the yeast-derived GPAT,LPAT, PAP — phosphatidic acid phosphatase, DGAT, and G3PDH, theoverexpression of single genes had limited effect on the TAG synthesis,whereas a combination of all the five genes in a unique construct result-ed in a two-fold increase of TAG content (Hsieh et al., 2012; Klok et al.,2014).

The tendency of oleaginous microalgae to synthesize, apart fromlipids, significant amounts of other energy-rich compounds such asstarch in processes competitive to lipogenesis (Bellou and Aggelis,2012), deserves attention since the lipid yield could be considerablyincreased by blocking competitive metabolic pathways. Indeed,Ramazanov and Ramazanov (2006) managed to increase lipid contentby 50% using a starchless mutant of Chlorella pyrenoidosa, while Wanget al. (2009) reported a 30-fold increase of lipid bodies by number andsize in a starchless mutant of C. reinhardtii. Similarly, Work et al.(2010) reported that the genetic blockage of starch synthesis in thesta6 and sta7-10 mutants of C. reinhardtii increased lipid contentunder nitrogen deprivation conditions. However, Siaut et al. (2011)suggested absence of negative correlation between starch reservesand lipid content when comparing mutants along with the variouswild-type strains.

Approaches to increase lipid accumulation by suppressing theβ-oxidation pathway have been successful in the case of plants andyeasts. In microalgae, this kind of gene suppression could only be ac-complished by randommutagenesis or through the use of RNA silencingas reported by Radakovits et al. (2010). Recently, research in diatomsshowed that most of the implicated lipases were downregulated duringgrowth under nitrogen deprivation, resulting in TAG accumulation(Yang et al., 2013). Similarly, targeted knockdown of a multifunctionalenzyme (lipase/phospholipase/acyltransferase) increased lipid contentwithout affecting growth in the diatom T. pseudonana (Trentacosteet al., 2013).

Both elongases and desaturases, are responsible for PUFA biosynthe-sis, and are not implicated in the lipid accumulation process. However,since PUFAs are highly desirable molecules, changes in unsaturationprofiles by introducing or regulating desaturases is a major target.Although, several elongase- and desaturase-encoding genes have beencharacterized in microalgae, engineering trials are still in the beginning.

Several efforts have been focused on the modification of the fatty acidcomposition (i.e. to increase the omega-3 or omega-6 fatty acid content)but they have been performedmostly in transgenic plants (Dehesh et al.,2001; Graham et al., 2007; Napier, 2007; Opsahl-Ferstad et al., 2003; Stollet al., 2005). Recently, Hamilton et al. (2014)managed to increase 8 foldsthe DHA content by expressing the heterologous Δ5 elongase from thepicoalga O. tauri in the diatom P. tricornutum. Surprisingly, lipid contentincreased up to 65% and EPA synthesis by 58% in P. tricornutum whenan annotatedΔ5 desaturase gene (PtD5b) was cloned and overexpressed(Peng et al., 2014). Likewise, a Chlamydomonas gene encoding for Δ4desaturase has been found to affect the biosynthesis of specific lipid frac-tions since reduced levels of this enzyme led to lower amounts ofmonogalactosyldiacylglycerol (MGDG), while its overproduction in-creased both the levels of 16:4 acyl groups in the cell extracts and thetotal amount of MGDG (Zäuner et al., 2012). Enhanced production ofPUFAs was also shown in yeast and plants when microalgal desaturaseswere introduced to either of those organisms. Specifically,Δ5 desaturasesof the microalgae O. tauri and O. lucimarinuswere functionally expressedin an engineered S. cerevisiae strain resulting in the production ofboth ARA and EPA (Tavares et al., 2010). Similarly, Petrie et al. (2010b)reported production of 26% EPA in plant leaf TAGs upon introductionof a newly-identified Δ6 desaturase-encoding gene from the marinemicroalga Micromonas pusilla.

Beside desaturases, FATs also affect fatty acid compositionof microalgal lipid. In particular, plant derived FATs (i.e. 12:0- and14:0-specific FATs from Umbellularia californica and Cinnamomumcamphora, respectively) were recently successfully engineered into thediatom P. tricornutum in order to alter the fatty acid composition andthe obtained results showed redirection of the fatty acid synthesis to-wards the biosynthesis of C12:0 and C14:0 (Radakovits et al., 2011).Contrary to plant derived FATs, microalgae derived FATs show no fattyacid specificity. As a result, when endogenous P. tricornutum FAT wasoverexpressed it did not result in an altered fatty acid composition,while overexpression of endogenous C. reinhardtii FAT resulted inshort-circuiting of fatty acids (Blatti et al., 2012; Gong et al., 2011).

Biotechnological perspectives of microalgal lipids

The production of microalgal lipids (intended for either as source ofPUFAs or as feedstock for biodiesel production) can be performedthrough specific processes (i.e. planned for this purpose) or in combina-tion with the production of other microalgal metabolic products havingpharmaceutical and/or nutritional interest, or even may arise byexploiting the algal biomass produced during wastewater treatment.

An evaluation of the various systems used for microalgal oil produc-tion is illustrated in Table 1. The majority of microalgal/cyanobacterialproduction in Australia, Israel and Japan currently occurs in openponds (Ratledge and Cohen, 2008; Spolaore et al., 2006) obviouslythanks to low capital investment and operating cost of this system.Under the best conditions, scientifically documented peak biomass pro-ductivities in open cultivation systemsdonot surpass 75–100 ton/ha·yr(Benemann et al., 1987). Therefore, algal production systems need to beimproved in order to become more competitive and economicallyviable.

Closed type PBRs (i.e. tubular, panel) are employed for the commer-cial production of pigments (i.e. beta-carotene, astaxanthin) byD. salinaandH. pluvialis (Ratledge and Cohen, 2008; Spolaore et al., 2006). Thesetypes of reactors can be used for the production of lipids containingPUFAs, since their relatively high cost can be covered by the high priceof these products.

Other than autotrophically, microalgae can be cultivated heterotro-phically or mixotrophically, as well. Mixotrophic cultivation seems tobe the most promising approach since in this case microalgae utilizeboth phototrophic and heterotrophic pathways concurrently (Lam andLee, 2012; Mata et al., 2010). Particularly, Perez-Garcia et al. (2011)reported that the specific growth rate of mixotrophically grown

Table 1Technical and economic assessment of the current systems used for cultivation of microalgae.Adapted from Davis et al. (2011).

Open pond Photobioreactor (PBR)

Algal cell density (g/L) 0.5 4Lipid in dry algal mass (%, wt/wt) 25 25Algae productivity 25 (g/m2·day) 1.25 (kg/m3·day)Capital investment Low HighEase of scale-up Good Variable (depends on PBR type)Control of growth conditions Low (practically uncontrolled) HighHarvesting cost High (low density culture) Low (higher density culture)Contamination risk High LowWater use High Low

For lipid production 10 MM gal/yrTotal capital cost (direct + indirect) ($MM) 390 990Net operating cost ($MM/yr) 37 55Total co-product credits ($MM/yr) 6 7


microalgae could be estimated as the sum of the specific growth rates ofcells grown under phototrophic and heterotrophic conditions. Further,growth under mixotrophic conditions could overcome problems relatedto light invasion. However, this kind of cultivation system alsomeets sev-eral restrictions mainly due to possible contaminations, which may becrucial for algal growth. Thus, closed type PBRs are preferred over openponds. The option of sterilizing the bioreactors in order to ensure asepticconditions might increase the whole cost of the process, which canbe only potentially overcome by the significantly higher yields obtainedin this type of culture (comparable to those obtained by heterotrophicoleaginous microorganisms) and/or the high value of the targetproduct. Several microalgal strains (i.e. Chlamydomonas globosa, Chlorellaprotothecoides, Chlorella sp., Chlorella vulgaris, Chlorella zofingiensis,C. minutissima, H. pluvialis, Nannochloropsis sp., P. tricornutum,Rhodomonas reticulate, Scenedesmus bijuga, Spirulina platensis, etc.) arecultivated under heterotrophic or mixotrophic conditions in large scalefor the production of lipids rich in PUFAs, pigments and proteins(Bhatnagar et al., 2011; Cerón García et al., 2006; Cheirsilp and Torpee,2012; Cheng et al., 2009; Ip et al., 2004; Kobayashi et al., 1992; Lee,2001; Liang et al., 2009;WenandChen, 2003;Woodet al., 2009). Biomassconcentration in the reactor can reach 5–15 g/L, 3–30 times higher thanthat produced under autotrophic growth conditions. The lipid contentof biomass grown under heterotrophic ormixotrophic growth conditionsis also much higher reaching up to 50% or more in the dry biomass.

Besides cultivation an important difficulty encountered in commer-cial microalgal applications is the harvest of biomass, which presentsseveral technological and economic complications. Biomass harvestingrequires the separation of solid from liquid and is a process coveringaround 30% of the total production cost (Brennan and Owende, 2010).Currently there are various harvesting methods, including flocculationor coagulation, flotation, filtration, sedimentation and centrifugation(see review C. Chen et al., 2011). The choice of the appropriate methodshould be considered according to the culture cell density, the size of themicroalgal cells, the target product, etc. Among the harvesting methodsmentioned above, filtration has been reported as an efficient and cost-effective one (Molina Grima et al., 2003; Zhang et al., 2010), with the vi-brating screen filter and themicrostainer to be themost popular devices(C. Chen et al., 2011). In large-scale operations, mechanical harvesters,such as continuous belts, are studied and/or already used by variouscompanies (Christenson and Sims, 2011). Alternatively, the harvestingstep may be skipped, e.g. by performing facilitated lipid extraction inthe whole culture, which reduces the process cost and enablesmicroalgal production economically feasible.

Occurrence of PUFAs in microalgal lipids and industrial applications

Microalgae belonging to different classes may have a particular fattyacid composition that is generally recognized as group specific. Diatoms

(Bucillariophyceae) are able to synthesize high amounts of palmitoleicacid (C16:1) and EPA, whereas some mutants may also produce ARA(Schneider et al., 1995; Sriharan et al., 1991). Chrysophyceae (knownas golden algae) produce a wide variety of fatty acids such as C16:1,EPA and DHA. Among them, high proportions of PUFAs have beenreported for Isochrysis strains (Molina Grima et al., 1993; Tatsuzawaand Takizawa, 1995). Gyrodinium and Crypthecodinium and othergenera belonging to Dinophyceae or Dinoflagellates have been charac-terized as C18:4, C18:5, EPA and DHA producers (Harvey et al., 1988;Kyle et al., 1992; Parrish et al., 1993; Reitan et al., 1994). Rhodophyta(red algae) are able to synthesize linoleic acid (C18:2), ARA and EPA(Dembitsky et al., 1991; Radwan, 1991). Chlorophyceae (green algae),possessing an active acyl-CoA desaturase system acting on C18 fattyacids (Regnault et al., 1995), include the most popular producers oflong chain PUFAs, such as alpha-linolenic acid (ALA), EPA and DHA(Bellou et al., 2012; Huang et al., 2013; Makri et al., 2011; Mendeset al., 2005; Patil et al., 2007; Pettitt and Harwood, 1989; Reitan et al.,1994; Yongmanitchai and Ward, 1991).

Strains with special fatty acid biosynthetic capacity, and therefore ofindustrial interest, belong to C. minutissima, having high PUFA content(Seto et al., 1984), and Schizochytrium sp., considered one of the bestsources of DHA (with a content of around 40% the total lipids), whichalso synthesizes EPA but in less percentages (i.e. 17%) (Doughmanet al., 2007; Kyle, 2001). Cryptocodinium cohnii, Amphidinium sp. andProrocentrum triestinum are also known as efficient DHA-producers(Kyle, 2001; Makri et al., 2011), while Porphyridum cruentum andN. salina are able to synthesize EPA at more than 25% of total lipids(Bellou and Aggelis, 2012; Cohen et al., 1988). Various other speciesare able to produce lipids with high content in PUFAs as summarizedin Table 2.

PUFAs, among all fatty acids and even all algal bioactive compounds,attract attention thanks to their obvious beneficial effect on humanhealth. They have been implicated in the treatment and prevention ofvarious diseases and disorders, including inflammatory disease, athero-sclerosis, thrombosis, arthritis and a variety of cancers (Dyerberg andJorgensen, 1982; Lagarde et al., 1986; Rousseau et al., 2003; Sakaiet al., 1990). EPA andDHA are known to regulate coagulation, lipoproteinmetabolism, blood pressure, and endothelial and platelet functions. Inparticular, PUFAs are important for the growth andperformance of retina,brain, reproductive tissues and for cardiovascular health (Bhakuni andRawat, 2005; Horrocks and Yeo, 1999). Moreover, they have an anti-proliferative effect on cultures of epithelial and bronchopulmonarycells (Moreau et al., 2006), caused myelo-suppression induced by lead(Queiroz et al., 2003) and improved glycogenesis in diabetic mice(Cherng and Shih, 2006).

Adarme-Vega et al. (2012) stated that microalgae are the main fattyacid producers in the marine food chain recognizing microalgal PUFAsas the essential nutrient for production of zooplankton necessary for

Table 2Fatty acid composition of selected microalgae belonging to different groups.

Microalgal strains C16:0 C16:1 n−7 C18:0 C18:1 n−9 C18:2 n−6 C18:3 n−6 C18:3 n−3 C18:4 n−3 C20:5 n− C22:6 n−3 References

ChromalveolataAmphidinium sp. 23.4 0.9 2.9 2.5 1.1 2.3 0.1 19.1 17.1 26.3 Makri et al. (2011)

ChlorophytaBotryococcus braunii 17.8 0.7 0.4 41.5 – – 28.6 – – – Tran et al. (2009)Chlamydomonas mexicana 40.0 – 2.0 3.0 37.0 2.0 16.0 – – – Salama et al. (2013)Chlamydomonas reinhardtii 36.1 1.8 4.4 13.3 17.8 – 20.5 2.1 – – Tatsuzawa et al. (1996)Chlamydomonas sp. 39.5 2.1 1.5 30.2 2.7 – 22.1 – – – Tatsuzawa et al. (1996)Chlamydomonas sp.1 16.6 3.3 0.7 1.6 10.2 – 29.0 – 19.2 – An et al. (2013)Chlorella pyrenoidosa SJTU-2 27.9 0.7 0.8 2.2 5.9 – 35.8 – – – Tang et al. (2001)Chlorella sp. 17.0 5.6 0.5 12.3 41.3 0.3 18.9 – – – Bellou and Aggelis (2012)Chlorella sp. 19.6 6.2 3.3 5.7 11.8 0.3 22.3 0.1 1.3 – Zhukova and Aizdaicher (1995)Dunaliella maritime2 11.8 2.7 0.4 2.1 4.1 3.2 42.6 1.3 – – Zhukova and Aizdaicher (1995)Dunaliella primolecta3 21.7 – 0.8 4.3 6.2 1.0 38.8 4.1 – – Viso and Marty (1993)Dunaliella salina4 17.8 0.8 1.5 2.8 6.1 2.5 36.9 0.7 0.1 – Zhukova and Aizdaicher (1995)Dunaliella tertiolecta 5 22.9 6.3 – 2.8 10.8 – 35.2 – – – Tsuzuki et al. (1990)Dunaliella tertiolecta6 10.3 4.0 0.3 1.7 5.2 3.2 38.7 1.3 0.4 – Zhukova and Aizdaicher (1995)Haematococcus pluvialisa 24.8 0.7 3.8 15.7 23.3 0.9 17.7 – 0.4 – Damiani et al. (2010)Nannochloris sp.7 15.0 16.2 1.0 3.9 0.6 – 0.8 0.3 – – Viso and Marty (1993)Parietochloris incisa8 19.8 – 18.2 10.2 14.3 14.3 – – 4.3 – Lang et al. (2011)Scenedesmus obliquus 23.2 1.3 0.7 28.3 15.7 3.6 27.3 – – – Salama et al. (2013)Scenedesmus obliquus SJTU-3 22.2 0.3 0.9 1.2 13.3 – 48.2 – 5.4 – Tang et al. (2001)Tetraselmis sp.9 16.2 3.8 0.9 4.7 7.0 0.3 15.5 12.1 5.6 – Zhukova and Aizdaicher (1995)Tetraselmis viridis10 15.9 3.3 0.8 4.6 3.1 0.2 15.0 13.3 6.7 – Zhukova and Aizdaicher (1995)

CyanobacteriaAnabaena viriabilis 35.8 21.4 – 7.4 14.3 16.0 – – – Tsuzuki et al. (1990)Anacystis (Synechococcus) nidulans 49.2 38.7 – 4.0 – – – – – – Tsuzuki et al. (1990)Anacystis (Synechococcus) sp. B-434 54.7 3.4 1.0 7.9 9.9 15.2 1.6 1.8 – – Maslova et al. (2004)Arthrospira (Spirulina) platensis 45.9 2.7 0.9 7.8 12.0 20.6 – – – – Colla et al. (2004)Arthrospira platensis 38.6 7.2 3.6 11.4 14.4 21.1 – – – – Andrich et al. (2006)Arthrospira platensis 46.8 4.4 3.1 12.2 18.8 14.3 – – – – Maslova et al. (2004)Arthrospira platensis D880 46.1 3.3 1.5 4.7 31.5 12.9 – – – – Muhling et al. (2005)Arthrospira sp. 49.9 6.1 1.2 2.7 21.2 18.5 – – – – Chaiklahan et al. (2008)Arthrospira sp. 39.8 10.2 1.8 4.2 18.4 20.9 – – – – Bellou and Aggelis (unpublished data)Arthrospira fusiformis D872/H1 46.6 3.5 1.4 6.1 18.8 23.6 – – – – Muhling et al. (2005)Arthrospira fusiformis D909 44.0 2.9 1.9 3.5 19.3 28.5 – – – – Muhling et al. (2005)Arthrospira indica D929 44.7 4.0 1.3 5.0 16.6 28.3 – – – – Muhling et al. (2005)Arthrospira maxima D867 47.0 2.8 1.4 7.5 15.2 26.1 – – – – Muhling et al. (2005)Gloeobacter violaceus 33.0 2.0 6.0 9.0 15.0 – 33.0 – – – Maslova et al. (2004)Nostoc commune 25.3 24.1 – – 12.5 38.1 – – – – Lang et al. (2011)Oscillatoria agardhii CC 1988 18.5 22.6 1.6 1.9 11.4 – 24.6 – – – Ahlgren et al. (1992)Oscillatoria agardhii NS 1988/89 17.3 15.1 2.2 3.5 4.3 0.5 23.2 1.2 2.6 1.8 Ahlgren et al. (1992)Synechocystis sp.11 18.8 30.1 – – – 14.3 – – – – Lang et al. (2011)Tolypothrix sp. 38.7 9.2 0.5 9.5 7.5 9.9 2.3 11.5 – – Maslova et al. (2004)

CryptophytaChroomonas salina 13.5 2.0 3.0 2.3 1.2 – 10.8 30.3 12.9 7.1 Zhukova and Aizdaicher (1995)Cryptomonas sp. 16.6 3.1 1.3 1.4 0.9 – 21.0 26.6 7.2 4.1 Renaud et al. (2002)Rhodomonas sp. 15.5 4.4 2.6 1.2 3.3 0.4 23.0 18.8 5.5 2.7 Renaud et al. (2002)

(continued on next page)

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3

Table 2 (continued)

Microalgal strains C16:0 C16:1 n−7 C18:0 C18:1 n−9 C18:2 n−6 C18:3 n−6 C18:3 n−3 C18:4 n−3 C20:5 n−3 C22:6 n−3 References

DinoflagellataGymnodinium kowalevskii 26.7 1.8 8.5 6.5 3.7 – 7.2 15.6 0.1 9.5 Zhukova and Aizdaicher (1995)

DinophytaProrocentrum minimum S2 39.2 3.2 4.9 3.8 2.7 2.0 2.3 12.3 3.7 20.1 Makri et al. (2011)Prorocentrum triestinum S1 38.2 0.6 4.0 6.2 7.9 0.9 0.8 11.2 1.7 22.0 Makri et al. (2011)

HaptophytaEmiliana huxleyi 10.3 – 10.8 42.2 – – – 8.7 – 9.2 Lang et al. (2011)Isochrysis galbana 11.5 3.3 – 13.1 7.0 – 3.8 12.5 0.8 15.8 Patil et al. (2007)Isochrysis sp. 12.9 6.7 0.6 8.4 5.7 0.7 8.4 13.5 0.6 6.6 Renaud et al. (2002)Isochrysis sp. 12.8 5.2 0.2 12.5 3.6 1.2 6.3 25.8 0.9 15.0 Huerlimann et al. (2010)Pavlova lutheri 11.1 26.3 – 5.2 0.6 – 0.5 9.1 18.0 9.7 Lang et al. (2011)Pavlova salina 15.1 30.4 1.0 3.1 1.5 2.2 – 4.2 19.1 1.5 Zhukova and Aizdaicher (1995)Pavlova sp. H 17.7 11.0 – 3.7 0.6 – – – 28.9 – Griffiths et al. (2012)Pavlova sp. L 19.9 16.2 – 3.8 0.6 – – – 23.4 – Griffiths et al. (2012)Pavlova viridis12 15.4 19.8 0.4 3.7 1.1 – 2.4 2.9 15.7 7.2 Huang et al. (2013)

HeterokontophytaAsterionella sp. (?) S2 22.3 13.9 2.8 3.5 2.3 3.0 1.1 7.7 26.4 8.9 Makri et al. (2011)Chaetoceros constrictus13 16.4 14.3 4.8 4.7 1.8 0.3 0.2 0.3 18.8 0.6 Zhukova and Aizdaicher (1995)Chaetoceros muelleri14 17.3 30.0 0.8 1.4 0.7 1.1 0.3 0.8 12.8 0.8 Zhukova and Aizdaicher (1995)Chaetoceros sp. (CS256)15 9.2 36.5 0.7 1.7 0.4 0.9 0.5 0.6 8.0 1.0 Renaud et al. (2002)Heterosigma akashiwo 40.0 12.7 – – 4.5 – 6.7 5.2 14.8 – Lang et al. (2011)Nannochloropsis oceanica 17.2 18.2 1.8 4.1 9.7 – 0.5 – 23.4 – Patil et al. (2007)Nannochloropsis oculata 20.5 25.2 1.8 3.6 2.2 0.7 0.2 0.1 29.7 – Zhukova and Aizdaicher (1995)Nannochloropsis oculata16 14.0 18.6 0.3 3.0 4.2 – – – 35.5 – Huang et al. (2013)Nannochloropsis salina 21.3 29.5 0.8 5.1 2.6 1.0 – – 26.3 – Bellou and Aggelis (2012)Nannochloropsis sp. 25.3 23.4 0.9 4.8 2.2 – – – 30.8 – Huerlimann et al. (2010)Nannochloropsis sp. 17.8 11.4 1.8 2.6 5.2 – 9.2 – 33.0 0.6 Andrich et al. (2005)Phaeodactylum tricornutum 13.4 29.3 0.7 5.3 – – – – 30.0 3.1 Tonon et al. (2002)Phaeodactylum tricornutum 16.6 26.0 0.6 1.8 1.5 – 0.3 3.3 28.4 0.2 Patil et al. (2007)Phaeodactylum tricornutum 11.3 21.5 0.4 2.3 1.5 0.5 0.9 0.5 28.4 0.7 Zhukova and Aizdaicher (1995)Phaeodactylum tricornutum H 23.7 45.5 – 5.7 0.9 – – – 14.0 – Griffiths et al. (2012)Schizochytrium sp.17 10.5 0.5 0.6 0.5 – – – – – 57.6 Chang et al. (2013)Thassiosira weissflogii18 17.2 24.3 2.5 5.9 – – – – 17.3 1.6 Borges et al. (2011)Thassiosira weissflogii19 27.5 0.3 1.4 1.3 – 1.0 – 0.3 3.8 0.1 Viso and Marty (1993)

MyzozoaCrypthecodinium cohnii 2.7 0.8 2.7 17.8 – – – – – 72.3 Couto et al. (2010)

OchrophytaSkeletonema costatum 9.4 19.0 2.2 2.6 1.6 – 0.2 2.9 15.4 2.3 Zhukova and Aizdaicher (1995)

RhodophytaPorphyridium cruentum20 33.5 3.0 0.8 0.7 5.7 0.2 – – 37.5 – Cohen et al. (1988)Porphyridium cruentum21 28.6 1.1 0.8 1.3 8.2 0.3 0.4 – 21.1 – Zhukova and Aizdaicher (1995)Porphyridium cruentum22 46.9 2.1 – – 8.8 – – – 20.3 – Tsuzuki et al. (1990)

Other fatty acids in significant concentrations: 1C20:3n−6, 14.6%; 2C16:4n−3, 22.6%;3C16:4n−3, 12.3%; 4C16:4n−3, 18.2%; 5C16:4n−3, 16.1%; 6C16:4n−3, 23.9%; 7C18:1n−7, 53.6%;8C20:4n−6, 14.0%; 9C16:4n−3, 18.3%; 10C16:4n−3, 19.9%;11C14:0, 42.5%; 12C22:5n−3, 7.4%; 13C14:0, 14.0; C16:3n−4, 7.9%; 14C14:0, 15.0; C16:3n−4, 7.8%; 15C14:0, 23.6%; 16C20:4n−6, 10.6%; 17C22:5n−3, 21.3%; 18C16:3n−3, 18.1%; 19C14:0, 24.8%; 20C20:4n−6, 17.3%; 21C20:4n−6, 27.6; and22C20:4n−6, 21.9.

a Calculated from data contained in Damiani et al. (2010).

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the first feeding of larvae. This can explain the fact that large amounts ofPUFAs are used infisheries,which employ an artificial food chain, for theenrichment of zooplankton (i.e. rotifer Brachionus plicatilis) that iswide-ly used in the first-feeding of marine fish larvae. Alternatively, themicroalgal cells are used as carriers for PUFA transfer when consumedby rotifers (Brown, 2002; Birkou et al., 2012; Guedes and Malcata,2012; see also supplementary material, Video 1), which are then usedas prey for fish larvae, and/or directly fed to fish larvae during a brief pe-riod (Fig. 1).

The majority of PUFAs synthesized by microalgae are of high bio-technological interest. Nevertheless, so far DHA is the only algal PUFAcommercially available. Indeed, although EPA can be produced in re-markable quantities bymicroalgal strains (i.e. Porphyridium purpureum,P. tricornutum, I. galbana, Nannochloropsis sp., etc.) it is not currentlyproduced in commercial scale because it cannot be counted as econom-ically competitive compared to other sources (Apt and Behrens, 1999;Belarbi et al., 2000; Spolaore et al., 2006). Further improvements in bio-mass and lipid yields and/or the combined production of PUFAs withother metabolites (see below) could open the market for more algalPUFAs in the near future.

Microalgal lipids as feedstock for biodiesel production

With the steady increase of world population and rapid industrialdevelopment, energy consumption has been increased significantly(http://www.eia.gov/todayinenergy/). The fossil fuels are widelyaccepted as non-renewable and unsustainable energy due to depletingresources, price fluctuating and causing increase of earth's temperature(Schenk et al., 2008). All these risks render the development of renew-able sources of energy a pressing mission.

Biodiesel arises to be premium alternative for fossil fuel, sinceseveral favorable environmental properties make it an attractive energy

Fig. 3. Concept of combined production of lipids and other high added-vThe individual photos are kindly provided by ALGAE A.C. (Nigrita Serres

source. Specifically, with biodiesel the CO2 balance is zero and there areno emissions of sulfur compounds (Antolin et al., 2002; Vicente et al.,2004), while carbon monoxide release is reduced by 30%. Additionally,its high biodegradability, lack of any aromatic compounds and 90%reduction in air toxicity may lead to up to 95% decrease in the relevantcancer cases (Sharp, 1996).

Currently biodiesel is produced from vegetable oil harvested frommany feedstock plants such as soybeans, rapeseed (Georgogianniet al., 2009), canola (Li et al., 2009; Thanh et al., 2010), sunflowerseeds (Harrington and D'Arcy-Evans, 1985), corn (Bi et al., 2010;Majewski et al., 2009) and palm (Kalam et al., 2008). About 7% of globaledible vegetable oil supplies were used for biodiesel production in 2007(Mitchell, 2008). However, extensive use of edible oils may aggravatefood supplies versus fuel issue (Anwar et al., 2010). Alternatively, non-edible vegetable oils produced by jatropha (Oliveira et al., 2009; Sahooet al., 2009), karanja (Das et al., 2009; Nabi et al., 2009; Naik et al.,2008; Sahoo et al., 2009), mahua (Godiganur et al., 2009) and polanga(Sahoo et al., 2009) can be used, as their fatty acid composition is suit-able for biodiesel production.

On the other hand, microalgae appear to be an excellent biodieselsource compared to the existing plants, since they are the fastest grow-ing photosynthetic organisms (Demirbas and Demirbas, 2011). More-over, they withstand utmost pH and temperature conditions and useCO2 in their photosynthetic process more efficiently (Shay, 1993).Microalgae do not need to be cultivated on agricultural areas but unsuit-able agricultural land can be utilized instead, and they can provide extrabiodiesel oil than oilseed crops while using minimumwater and main-land (Sheehan et al., 1998). Moreover, lipid productivity reported formany microalgae greatly exceeds the oil productivity of the best pro-ducing oil crops, demonstrating that algae give the maximum biodieselyield and thus they may be able to produce up to 200 times the amountof oil per unit of surface compared to soybeans (Chisti, 2007). These

alue metabolites, such as polysaccharides, proteins, and pigments., Greece).

http://www.eia.gov/todayinenergy/

image of Fig.�3


properties make microalgae the most promising organisms on earththat have the potential to displace the petroleum-based diesel fuelcompletely without adversely affecting supply of food and other cropproducts.

The algal biofuel technology is still in its infancy and currently thereare no industrialized systems producing algal oil on the scale requiredfor biodiesel production (Veal et al., 2013). Although the production ofbiofuels from algal biomass is technically feasible, there is a need forgreat efforts, in order tomake feasible the development of economicallyviable large-scale algal biofuel enterprises. Actually, despite the algaladvantages over all other organisms concerning biodiesel production,up to now algal biodiesel is more expensive than fossil diesel. Muchmore productive strains should be employed (i.e. having higher growthrates and able to accumulate N50% lipids), while technical restrictionssuch as the high harvesting cost and the extremely large area of lagoonsneeded for biodiesel production, which is not available (Ratledge andCohen, 2008), should be faced.

Combined production of microalgal lipids

A direct reduction of the production cost of microalgal lipids can beachieved by combining lipid production with other applications. Theconcept of combined lipid production is illustrated in Fig. 3. Actually,microalgae are used in various commercial applications (i.e. in the en-hancement of nutritional value of food and animal feed, in aquacultureand pharmaceutical industry, etc.) (Birkou et al., 2012; Spolaore et al.,2006). Besides PUFAs, compounds such as beta-carotene and polysac-charides, which are produced commercially by various species, providea strong role inmanufacturing some therapeutic supplements that com-prise an importantmarket in biotechnological industries (PriyadarshaniandRath, 2012). Such properties also enable the use ofmicroalgae in thecommercial production of cosmetics (Priyadarshani and Rath, 2012;Spolaore et al., 2006).

Microalgae also have applications in environmental biotechnologysince they can be used for bioremediation ofwastewater and tomonitorenvironmental toxicants.

Combined production of lipids and pharmaceutical productsThe microalgae have gained significant attention as natural factories

and rich sources of novel and potential bioactive molecules of interestfor the pharmaceutical and cosmetic industries (Rania and Hala,2008). The combined production of bioactive products and lipids,when possible, can obviously support the commercial viability of bothprocesses. Hydrophobic compounds can be extracted simultaneouslywith lipids and then purified, while hydrophilic compounds such asproteins and sugars can be extracted from the defatted biomass.

Natural cosmeceuticals from algae have become amajor counterpartfor superficial application to the human skin (Raja et al., 2008; Spolaoreet al., 2006). Algal proteins or derivatives are used in skin repair andhealing products (Hagino and Masanobu, 2003). Moreover, the algalcosmetics have useful features, such as anti-irritant, immuno-stimulant, antioxidant, anti-aging and anti-inflammatory properties(Morist et al., 2001). Members of Chlorella and Arthrospira, alreadyused in lipid production, are also well known in the skin care marketfor their hydrophilic extracts (Stolz andObermayer, 2005). Additionally,diatoms that produce polar lipids rich in PUFAs along with carotenoids,phytosterols, vitamins, and antioxidants, contribute to the health bene-fits of the produced oils (Li et al., 2014). The protein extract from blue-green algae, which include phycoerythrin, possesses many bioactivities,i.e. anti-inflammatory, hepetoprotective, antitumor, antiviral andneuroprotective (Bermejo et al., 2002; Kim et al., 2008; Sekar andChandramohan, 2008). Furthermore, C. vulgaris extract stimulatescollagen synthesis in the skin and is therefore used for wrinkle reduc-tion and tissue regeneration (Spolaore et al., 2006).

Besides PUFAs, several valuable products i.e. carotenoids, astaxanthin,allophycocyanin, phycocyanin, phenols, acetogenins, terpenes, indoles,

etc. of pharmacological interest can be extracted from algae. These com-pounds possess antifungal, antiprotozoal, antiviral, neuroprotective,antiplasmodial, antimicrobial, anti-inflammatory and antioxidant proper-ties (Cardozo et al., 2007; Ghasemi et al., 2004;Herrero et al., 2006; Kellanand Walker, 1989; Mayer and Hamann, 2005; Mendiola et al., 2007;Ozemir et al., 2004; Sekar and Chandramohan, 2008). Microalgal pig-ments are very interesting compounds as they are safe, eco-friendly andhave a wide range of therapeutic uses including prevention and treat-ment of acute and chronic diseases, rheumatoid arthritis, atherosclerosis,neurological disorders, cataract and muscular dystrophy (Sies and Stahl,2004). Daily consumption ofmicroalgae derived astaxanthinmay protectbody tissues from oxidative damage as this might be a practical andbeneficial strategy in health management, since it has been suggestedthat astaxanthin has a free radical fighting capacity worth 500 timesthat of vitamin E (Dufossé et al., 2005). Additionally, cytotoxic, pro-apoptotic and anti-proliferative effects were reported for a large numberofmicroalgal pigments (such as phycobilin, chlorophyll, and epoxycar de-rivatives) when applied at very low concentrations (Konishi et al., 2006;Murakami et al., 2002; Nishino et al., 1992; Pasquet et al., 2011; Shakeret al., 2010; Sugawara et al., 2007, 2009; Yoshida et al., 2007). Althoughthe natural algal carotenoids are more expensive than the syntheticones, at least natural beta-carotene has specific physical properties thatmake it superior to the synthetic one (García-González et al., 2005;Guerin et al., 2003). Among the variousmicroalgal species, the oleaginousD. salina is the preferred organism for beta-carotene production, since itcan accumulate up to 14% of this pigment in dry weight (Metting,1996). Furthermore, phycobiliproteins (including phycocyanin,phycoerythrocyanin and allophycocyanin) have been extracted fromvarious marine algae (Niu et al., 2006). Among them P. cruentum andSynechococcus spp. are currently used in large scale for phycobiliproteinproduction (Viskari and Colyer, 2003).

Recently, there is increased interest for the fucoxanthin (fuco), amicroalgal cytotoxic pigment, owing to its strong pro-apoptotic,anti-proliferative, and cytotoxic activities (Heydarizadeh et al., 2013;Ishikawa et al., 2008; Nishino et al., 1992; Yamamoto et al., 2011; R.X.Yu et al., 2011). Fuco protects against reactive oxygen species (ROS)and UV-induced DNA damage (Heo and Jeon, 2009; Shimoda et al.,2010) and easily inhibits mammalian DNA–DNA-dependent polymer-ases (Murakami et al., 2002). Odontella aurita, a microalga containingin its lipids EPA in high concentrations (28% of total lipids), has beencultivated in open ponds for commercial purposes. This organism canaccumulate fuco in high concentrations (Xia et al., 2013) and thereforeEPA production could be combinedwith the production of this valuablemolecule.

Apart from pigment compounds, taurine (an algal peptide) hasseveral functional and biological applications (Houstan, 2005). Recently,taurine has become a common component in beverages, foods andnutritional supplements (Dawczynski et al., 2007). In addition, glyco-proteins (lectins), extracted from marine algae, are considered a typeof interesting, for biochemical research, proteins as well and can be iso-lated with their carbohydrate moiety. Extracts (or purified peptides)frommacro- and microalgae are shown to have novel antihypertensiveand angiotensin converting enzyme (ACE) inhibitory activities withminimal or no side effects and can thus be used as alternatives to syn-thetic drugs (Kim and Wijesekara, 2010; Suetsuna and Chen, 2001;Suetsuna and Nakano, 2000). Enzymatic hydrolysates having ACEinhibitory properties have been extracted from various macroalgae(Athukorala and Jeon, 2005), the most potent of which were those ofEcklonia cava that is an edible marine brown algal species found inJapan and Korea. Alternatively, extracts of C. vulgaris and Arthrospiraplatensis have potent ACE inhibitory properties (Sheih et al., 2009),and the particular species are also of interest for their lipids. Other pep-tides derived from C. vulgaris have been shown to suppress matrixmetalloproteinase-1 level in human skin fibroblast cells, which is in-duced by solar ultraviolet B. The particular inhibition may occur evenat the level of gene expression (C.L. Chen et al., 2011). Furthermore,


the biliprotein C-phycocyanin, extracted from A. platensis has been suc-cessfully used against the humanhepatocarcinoma (HepG2) and chron-ic myeloid leukemia (K562) cell lines (Nishanth et al., 2010; Subhashiniet al., 2004).

Lipids as co-products of environmental applicationsMicroalgae are mainly autotrophic microorganisms that are able to

fix CO2 from different sources, such as atmosphere, industrial exhaust

Fig. 4. The facilities of municipal wastewater treatment located in the city of Patras (Westerngrown is framed in red (a). Nile red staining of lipids in micro- and macro-algal cells sampled

gases (e.g. flue gas and flaring gas) or to use fixed forms of CO2

(e.g. NaHCO3 and Na2CO3). Thanks to these characteristics, manybiotechnological applications are carried out using microalgae in envi-ronmental safety and maintenance, such as bioremediation, bioassayand bio-monitoring of toxicants (Harun et al., 2010; Hoffmann, 1988;Kirkwood et al., 2003; Phang et al., 2001).

Several oleaginous microalgae, interesting for biodiesel feedstockproduction, can be used in the treatment of municipal wastewaters.

Greece). The last-stage settling tank in which micro- and macro-algae are spontaneouslyfrom the last-stage settling tank. White arrows show the yellowish lipid bodies (b).

image of Fig.�4

Fig. 4 (continued).


Wang et al. (2010) cultivated the oleaginous microalga Chlorella sp. onvariouswastewaters coming from four different stages of the treatmentprocess in a municipal wastewater treatment plant. The microalga, inthe particular study, was able to grow in wastewaters before primarysettling, after primary settling, after activated sludge tank and thosegenerated in sludge centrifuge, and simultaneously remove nitrogen,phosphorus, chemical oxygen demand, metal ions, etc. Similar resultswere obtained for the microalgae Halochlorella rubescens, Scenedesmusacutus, and Chlorella sorokiniana when grown in wastewatersautrophically, mixotrophically and/or heterotrophically (Kim et al.,2013; Mahapatra et al., 2014; Sacristán de Alva et al., 2013; Shi et al.,2014).

The production of algal biomass spontaneously grown in biologicaltreatment plants depends on the climate. In Greece (Patras) importantquantities of biomass are produced annually in the tanks of final settling(Fig. 4a), even if the retention time is low. This biomass, consisted ofmacro- and micro-algae (Fig. 4b) contains 7–10% lipids and large quanti-ties of protein, both of interest in the biofuel (biodiesel, biogas,biohydrogen) production. Fatty acid analysis of lipids synthesized by themicroalgal community showed that the major fatty acid was palmiticacid (up to 24% w/w in total lipids), followed by ALA and oleic acids(17.6 and 15.3%, respectively). Non-negligible amounts of EPA (around7.0%) were also detected in several samples (unpublished data).

Several species can be used for the treatment of toxic waters. A pondsystem, which uses C. vulgaris as biological material, showed efficiencyin treating wastewater containing toxic contaminants (Hoffmann,1988; Phang et al., 2001). The oily biomass produced in such a reactorcould be further valorized in the production of biodiesel.

Conclusions

The most significant bottlenecks that limit the production ofmicroalgal oil in large scale are primarily the restricted lipid synthesisin the microalgal cell and secondarily the low growth rate of theseorganisms (Davis et al., 2011). Both constraints, negatively affectingoil productivity, are of biological origin and therefore their solutionsshould be sought in laboratories of molecular biology and biochemistry.Genetic engineering of strains with such enhanced performances(having improved biosynthetic capabilities) is a great challenge for theresearchers working in the field and solutions are expected in the nextfewyears. The type II CRISPR/Cas system, being developed as a powerfulgenome-editing tool, applicable to virtually any organism (Hsu et al.,2014) could also be employed for the construction of oleaginousmicroalgae with desirable properties. The versatility and tunable speci-ficity of the particular systemmake it ideal as a screening tool, targetingspecific groups of genes rather than thewhole genome, in search of cellscompetent to both grow to high densities and be sufficiently oleaginous.Further, evolution of such genetically modified cells under restrictiveconditions could potentially allow finding relatively genetically stablestrains as well. Nevertheless, it seems that blocking competitive to lipo-genesis pathways and/or inhibiting lipid turnover are more effectivestrategies than those dealingwith the improvement of the liposyntheticapparatus.

The commercial production of microalgal PUFAs is currently a muchmore attainable target than the production of biodiesel. Actually, thenatural sources of PUFAs are very limited and therefore microalgaehave a great industrial potential. It is certain that the large number of

image of Fig.�4


the current, as well as of the upcoming, applications of PUFAs will forceresearch towards seeking for effective solutions on crucial questions re-lated to the biochemical restrictions affecting microalgal biomass andlipid productivity, and harvesting.More research is needed in all aspectsof PUFA production including biochemical andmolecularwork, genomecharacterization of model organisms and the isolation and characteriza-tion of new oleaginous strains. Indeed, the introduction of new biologi-cal material holding unconventional biochemical arsenal, e.g. havinghigh light energy conversion yield, can widen the field and alternativeideas can be generated.

The perspective of algal biodiesel is currently poor due to several bi-ological and technical restrictions. The production cost of algal biodieselis currently very high when compared to that of fossil diesel. The con-struction of new strains able to accumulate large lipid quantities andthe development of new reactors for efficient cultivation of microalgaeare some urgent issues that should be faced.

Besides lipids, several valuable products, i.e. carotenoids, astaxanthin,allophycocyanin, phycocyanin, phenols, acetogenins, terpenes, indoles,etc., of pharmacological interest can be extracted from algae. The produc-tion of these valuable compounds in combination with lipid productionmay increase the financial viability of both processes. Equally, oleaginousmicroalgae able to grow onmunicipal or industrial wastewaters could befurther exploited in thebiofuelmanufacture. However, all theseprocessesneed to be individually studied for their efficiency and financial viability.

Supplementary data to this article can be found online at http://dx.doi.org/10.1016/j.biotechadv.2014.10.003.

Acknowledgments

Financial support was provided by the King Abdulaziz University(Jeddah, Saudi Arabia). Project title: Biotechnological production ofPUFA rich SCOs from microalgae.

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Microalgal lipids biochemistry and biotechnological perspectives

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