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Microalgal lipids biochemistry and biotechnological perspectives

Stamatia Bellou, Mohammed N. Baeshen, Ahmed M. Elazzazy, DimitraAggeli, Fotoon Sayegh, George Aggelis

PII: S0734-9750(14)00152-9DOI: doi: 10.1016/j.biotechadv.2014.10.003Reference: JBA 6847

To appear in: Biotechnology Advances

Received date: 15 June 2014Revised date: 2 October 2014Accepted date: 6 October 2014

Please cite this article as: Bellou Stamatia, Baeshen Mohammed N., ElazzazyAhmed M., Aggeli Dimitra, Sayegh Fotoon, Aggelis George, Microalgal lipids bio-chemistry and biotechnological perspectives, Biotechnology Advances (2014), doi:10.1016/j.biotechadv.2014.10.003

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Microalgal lipids biochemistry and biotechnological perspectives

Stamatia Bellou1, Mohammed N. Baeshen

2, Ahmed M. Elazzazy

2, Dimitra Aggeli

3,

Fotoon Sayegh2 and George Aggelis

*1,2

1Division of Genetics, Cell & Development Biology, Department of Biology,

University of Patras, Patras 26504, Greece

2Department of Biological Sciences, King Abdulaziz University, Jeddah 21589,

Saudi Arabia

3Department of Genetics, Stanford University, Stanford, California 94305, USA

*Corresponding author.

Email address: George.Aggelis@upatras.gr

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Abstract

The last few years, there has been an intense interest in using microalgal lipids in

food, chemical and pharmaceutical industries and cosmetology, while a noteworthy

research has been performed focusing on all aspects of microalgal lipid production.

This includes basic research on the pathways of solar energy conversion and on lipid

biosynthesis and catabolism, and applied research dealing with the various biological

and technical bottlenecks of the lipid production process. In here, we review the

current knowledge in microalgal lipids with respect to their metabolism and various

biotechnological applications, and we discuss potential future perspecives.

The committing step in fatty acid biosynthesis is the carboxylation of acetyl-

CoA to form malonyl-CoA that is then introduced in the fatty acid synthesis cycle

leading to the formation of palmitic and stearic acids. Oleic acid may be also

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 pathway.

Current research, aiming to enhance lipogenesis in the microalgal cell, is focusing on

over-expressing key-enzymes involved in the earlier steps of the pathway of fatty acid

synthesis. A complementary plan would be the repression of lipid catabolism by

down-regulating acylglycerol hydrolysis and/or β-oxidation. The tendency of

oleaginous microalgae to synthesize, apart from lipids, significant amounts of other

energy-rich compounds such as sugars, in processes competitive to lipogenesis,

worths attention since the lipid yield may be considerably increased by blocking

competitive metabolic pathways.

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The majority of microalgal production occurs in outdoor cultivation and for

this reason biotechnological applications face some difficulties. Therefore, algal

production systems need to be improved and harvesting systems need to be more

effective in order for their industrial applications to become more competitive and

economically viable. Besides, a reduction of the production cost of microalgal lipids

can be achieved by combining lipid production with other commercial applications.

The combined production of bioactive products and lipids, when possible, can support

the commercial viability of both processes. Hydrophobic compounds can be extracted

simultaneously with lipids and then purified, while hydrophilic compounds such as

proteins and sugars may be extracted from the defatted biomass. Microalgae have also

applications in environmental biotechnology since they can be used for

bioremediation of wastewater and to monitor environmental toxicants. Algal biomass

produced 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 to manipulate biochemical pathways involved in lipid

biosynthesis and towards cost effective algal cultivation and harvesting systems, as

well.

Keywords: Microalgae; lipid biosynthesis; genetic engineering; polyunsaturated fatty

acids; biodiesel; pigments; proteins; wastewater treatment

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1. Introduction

The microalgae are photosynthetic microorganisms that play a key role in the natural

ecosystems supplying organic matter and specific molecules, such as polyunsaturated

fatty acids (PUFAs), to many higher organisms. Microalgae applications range from

human and animal nutrition to cosmetics and the production of high value molecules.

The systematic development of microalgal applications started in the 20th

century. Several species are currently cultivated in large scale, in artificial or natural

ponds and rarely in photobioreactors (PBRs), producing up to some tons of biomass

and/or various metabolites per year. Algal biomass is rich in PUFAs, minerals (e.g.

Na, K, Ca, Mg, Fe, Zn and trace minerals) and vitamins, such as riboflavin, thiamin,

carotene and folic acid and so on (Garcia-Garibay et al., 2003; Becker, 2004;

Samarakoona and Jeona, 2012). The microalgal biomass, especially that produced by

the species Dunaliella, Arthrospira (Spirulina, cyanobacterium) and Chlorella, is

already marketed in various forms designed for human nutrition or is incorporated

into foods and beverages (Yamaguchi, 1997; Liang et al., 2004), as it is considered a

healthy nutritional supplement (Apt and Behrens, 1999; Borowitzka, 1999; Jensen et

al., 2001; Soletto et al., 2005; Priyadarshani and Rath, 2012). Similarly, the

consumption of even small amounts of microalgal biomass can positively affect the

physiology of animals by improving immune response, diseases’ resistance, antiviral

and antibacterial protection, improved gut function, probiotic colonization

stimulation, as well as enhanced feed conversion, reproductive performance and

weight control (Harel and Clayton, 2004). Although the quality of algal proteins lags

behind animal proteins, is superior to that of common plants (Kay and Barton, 1991;

Becker, 2004; Barrow and Shahidi, 2008; Um and Kim, 2009; Sydney et al., 2010;

Samarakoona and Jeona, 2012). Particular algal peptides, such as taurine, are of great

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nutritional and pharmaceutical interest (Houstan, 2005), while glycoproteins (lectins),

extracted by marine algae, are considered a type of interesting proteins for

biochemical and clinical research, and can be isolated with their carbohydrate moiety

(Silva et al., 2010).

Some species contain considerable amounts of pigments that are used in

cosmetics and as natural coloring agents. Many industrial production plants are

established in China, Australia and USA (Brown et al., 1997; Leon et al, 2003;

Garcia-Gonzalez et al., 2005) dealing with beta-carotene production (e.g. from

Dunaliella salina) that is used as a food coloring (Metting, 1996). Other pigments

such as phycobiliproteins have been extracted from various marine algae including

Porphyridium cruentum and Synechococcus spp. (Viskari and Colyer, 2003).

Although the majority of applications concern biomass production destined for

animal or human consumption- in fact, 30% of the current world algal production is

sold for animal feed applications (Becker, 2004)- there has been an increased interest

in the use of microalgal lipids in numerous commercial applications, such as in food,

chemical and pharmaceutical industry and cosmetology. Indicative of the high interest

in microalgal lipids is the noteworthy research that has been performed the last decade

on all aspects concerned microalgal lipid production. These include fundamental

research on the mechanisms used for light energy conversion and on lipid

biosynthesis and catabolism, as well as biotechnological research dealing with the

various technical bottlenecks of the lipid production process. In the current review

article the up-to-date level of knowledge in lipid biosynthesis and turnover in

microalgae and the various biotechnological applications and future perspectives of

microalgal lipids are comprehensively presented and discussed. Τaking into

consideration recent techno-ecomomic analyses concluding that the algal lipid content

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is the most critical factor affecting the viability of large-scale applications, especially

those related to biodiesel, the current research efforts aimed to reinforce algal

liposynthetic machinery using genetic engineering are also discussed.

2. Microalgal lipids in the forefront of lipid biotechnology

Several microalgal species are able to accumulate appreciable lipid quantities, and

therefore are characterized as oleaginous. Lipid content in microalgae can reach up to

80% in dry biomass, but even in these cases the lipid productivity is actually low. In

widespread species belonging to the genera of Porphyridium, Dunaliella, Isochrysis,

Nannochloropsis, Tetraselmis, Phaeodactylum, Chlorella and Schizochytrium, lipid

content varies between 20 and 50%. However, higher lipid accumulation can be

reached by varying the culture conditions. Factors such as temperature, irradiance

and, mostly, nutrient availability have been shown to affect lipid content and

composition in algal cells.

The interest for algal lipid arises mainly from the fact that these organisms are

able to synthesize considerable quantities of PUFAs that either reach humans via the

food chain or are used as food supplements (Figure 1). Indeed microalgae are the

primary source of PUFAs having nutritional and pharmaceutical interest (Kyle, 2001;

Doughman et al., 2007). Although fish are also a source of PUFAs, these organisms

usually obtain their PUFAs via bioaccumulation through the food chain (Benemann et

al., 1987). Furthermore, fish PUFAs production depends on fish quality and

sufficiency while that of algae does not.

Several microalgae are able to synthesize omega-3-long chain PUFAs, at

levels over 20% of their total lipids. In algal cell PUFAs are esterified with an

alcohol, usually glycerol, generating triacylglycerols (TAGs) or polar lipids (i.e.

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phospholipids, glycolipids) of exceptional structure regulating membranes fluidity

and function. Depending on the strain, lipid industrial production can be combined

with the production of other metabolic products of high value, such as beta-carotene

and astaxanthine. The main drawback in using microalgae to produce PUFAs rich

lipid in large scale is the low lipid content in algal cell and the low biomass density in

the reactor, usually not exceeding 400-600 mg/L under industrial culture conditions,

which increases considerably the harvesting cost.

Alternatively, microalgal lipids represent an attractive source of oil, suitable as

feedstock for biodiesel production. Actually, microalgae offer a number of advantages

from an industrial perspective. These include simplicity of culture, increased

photosynthetic efficiency and growth rates, higher biomass and oil productivities

compared to terrestrial plants, and higher rates of CO2 fixation and O2 release. The

final algal oil production per unit of surface may reach 200 times that of common

plant oils (Chisti, 2007). However, biodiesel produced by algae is more expensive

than that produced by conventional plants, while fossil fuel is much cheaper.

3. Lipid metabolism

Most microalgae accumulate lipids under specific environmental stress conditions,

such as nitrogen or phosphate limitation (Hu et al., 2008; Courchesne et al., 2009;

Amaro et al., 2011; Bellou and Aggelis, 2012; Msanne et al., 2012; Hu et al., 2013).

Therefore the management of the environmental conditions is a common approach

used for improving lipid accumulation in the microalgal cell. Strain selection is also

likely to be of critical importance. Recenlty, there has been an intense interest in

isolating new native microalgal strains when a large-scale application is intended.

Microalgae are exposed to a variety of changes in the environment. Seasonal cycles

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vary according to the climatic and geographical location of the habitat, in which they

are growing, and different strains of the same species may respond differently. Strains

derived from diverse geographical locations are physiologically different (Bouaicha et

al., 2001; Delgado et al., 1997), and could vary in their response to any limiting

environmental factor. Consequently indigenous strains may have developed

mechanisms for sensing and acclimating to changes in their environment, and thus

their use promises higher potentiality (Vonshak and Torzillo, 2004).

Understanding lipid metabolism and how it is controlled during algal 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, fatty acid biosynthesis and modification (both elongation

and desaturation) and lipid catabolism have not been clarified for microalgae as they

have for plants and heterotrophic microorganisms (Beer et al., 2009; Moseley et al.,

2009; Moellering et al., 2010; Khozin-Goldberg and Cohen, 2011; Boyle et al., 2012).

Especially, the overall lipid biosynthesis pathway and its regulators have not been

clearly described. Efforts to improve lipid accumulation in microalgae by modifying

the expression of key enzymes implicated in lipid synthesis often fail, suggesting a

lack in our understanding of the mechanisms that govern lipid accumulation, if not

algal cell biology.

3.1. 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 pyruvate and thereafter to acetyl-CoA, via a reaction

catalyzed by the pyruvate dehydrogenase complex (PDC), initiates the lipid

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biosynthetic pathway, which occurs in the plastid. Acetyl-CoA can also be generated

via a biochemical pathway that permits the conversion of polysaccharides into lipids

(Bellou and Aggelis, 2012), pathway that is commonly utilized by oleaginous

heterotrophs during sugar assimilation (Gema et al., 2002; Papanikolaou et al., 2004;

Ratledge, 2004; Fakas et al., 2008; Chatzifragkou et al., 2010; Makri et al., 2010;

Papanikolaou and Aggelis, 2011; Bellou et al., 2012; 2014a; 2014b) (Figure 2).

Specifically, the breakdown of storage polysaccharides (occurring e.g. under light

limitation) usually generates energy through glycolysis occurring in the cytosol

followed by the citric acid cycle occurring in the mitochondrion. However, several

environmental stresses, such as nitrogen or phosphate limitation, may disturb the

citric acid cycle (i.e. by inhibiting NAD+-isocitrate dehydrogenase) leading to citrate

accumulation in the mitochondrion and subsequently to its excretion in the cytosol.

Cytosolic ATP-dependent citrate lyase converts citrate into oxaloacetate and acetyl-

CoA; the latter is converted into malonyl-CoA by cytosolic acetyl-CoA carboxylase

(ACC) and becomes available for fatty acid elongation in the membranes of the

endoplasmic reticulum (ER) - see below (Mühlroth et al., 2013). Although this

mechanism has only been shown in Nannochloropsis salina and Chlorella sp.

cultivated in a lab-scale open pond-simulating PBR (Bellou and Aggelis, 2012), it is

probably common in oleaginous strains that are able to grow heterotrophically.

Despite the importance of acetyl-CoA biosynthesis, the committing step in

fatty acid biosynthesis is the carboxylation of acetyl-CoA to form malonyl-CoA,

reaction catalyzed by the ACCs located either in the plastid or in the cytosol (Kim,

1997; Davis et al., 2000; Khozin-Goldberg and Cohen, 2011; Lei et al., 2012; Baba

and Shiraiwa, 2013) (Figure 2). In algae, ACCs exist in two different forms, the

heteromeric and homomeric. Although it is generally believed that the heteromeric

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form is the one present in algal plastids, Huerlimann and Heimann (2013) reported

that this is not the case for all algae, since the presence of heteromeric or homomeric

ACCs is dependent on the origin of plastid. I.e. in Chlorophyta (except for

Prasinophyceae) and Rhodophyta, heteromeric ACCs have been found in their

plastids, whereas Heterokontophyta, Haptophyta and Apicomplexa contain

homomeric ACCs in their plastids. In several microalgal strains, i.e. those belonging

to Galdiera sulpuraria, Cyanidioschyzon merolae, Thalassiosira pseudonana and

Phaeodactylum tricornutum, two ACCs have been identified, the plastidial ACC1 and

the cytosolic ACC2 (Khozin-Goldberg and Cohen, 201; Huerlimann and Heimann,

2013). Two genes coding for ACC have been also found in Nannochloropsis gaditana

(Radakovits et al., 2012).

In the plastid, the malonyl-CoA is transferred to the acyl-carrier protein

(ACP), which is one of the subunits of the fatty acid synthase (FAS) complex, by the

malonyl-CoA:ACP transacetylase (Subrahmanyam and Cronan, 1998; Hu et al., 2008;

Greenwell et al., 2010; Blatti et al., 2012). The malonyl-ACP is introduced in the fatty

acid synthesis cycle through the 3-ketoacyl-ACP synthase (KAS). The KAS catalyzes

the condensation of an acetyl group with malonyl-ACP to form ketobutyryl-ACP.

This compound is converted via the sequential reactions of reduction- dehydration-

reduction to butyryl-ACP and the cycle is repeated until the formation of palmitoyl-

ACP. The latter is then converted into stearoyl-ACP after the addition of only two

carbon molecules originated from acetyl-CoA. Oleoyl-ACP is also synthesized after

desaturation of stearoyl-ACP, reaction mediated by the plastidial Δ9 desaturase (Yu et

al., 2011a; Mühlroth et al., 2013). Principally, the fatty acids are released from ACP

by a fatty acyl-ACP thioesterase (FAT), located in the chloroplast envelope. They are

activated thereafter into acyl-CoA by the long-chain acyl-CoA synthetase (LACS),

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also located in the chloroplast envelope, and transferred in the cytosol, where they

become available for lipid synthesis. Alternatively, in plants, and probably in

microalgae, the acyl chains may be used for structural lipids (mostly glycolipids)

synthesis in the plastid. For this purpose they are transferred from ACP either to G3P

or to monoacylglycerol-3-phosphate via the action of a plastidial acyltransferase

(Ohlrogge and Browse, 1995). All three enzymes ACP, KAS and FAT, have been

shown to play an essential role in fatty acid synthesis in Haematococcus pluvialis (Lei

et al., 2012). The transferred in the cytosol acyl-CoA chains are esterified with

structural phospholipids of the ER to be converted into higher derivatives (PUFAs).

The modified or not in the ER fatty acids are used as building blocks for the formation

of TAGs via the Kennedy pathway. The implicated in the Kennedy pathway

acyltransferases (i.e. diacylglycerol acyltransferase- DGAT, glycerol-3-phosphate

acyltransferase- GPAT, lyso-phosphatidic acid acyltransferase- LPAAT, lyso-

phosphatidylcholine acyltransferase- LPAT) are also located in the ER (Radakovits et

al., 2010; Chen and Smith, 2012; Merchant et al., 2012; Liu and Benning, 2013).

As reported by Chen and Smith (2012), DGAT is found in algae in two

isoforms (DGAT1 and DGAT2) catalyzing the same reaction but with significant

variations in sequence. Although the genes encoding for DGATs have been found in

various microalgae, such as Ostreococcus tauri, Thalassiosira pseudonana,

Nannochloropsis sp., the model organism Chlamydomonas reinhardtii, etc., their

function has not been clearly characterized and thus it needs further studying (Khozin-

Goldberg and Cohen, 2011). Recently, Wang et al. (2014) reported that in

Nannochloropsis species 1 or 2 DGAT1 and 11 DGAT2 gene doses exist, while only

6 and 4 DGATs genes respectively were found in Chlamydomonas reinhardtii and

Thalassiosira pseudonana and even fewer in some other green algae and heterokonts.

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Nannochloropsis gaditana genome analysis showed common homologues of ACC,

KAS, glycerol-3-phosphate-dehydrogenase (G3PDH), GPAT and LPAAT genes that

are also found in other microalgal strains, i.e. the brown alga Ectocarpus siliculosus,

the diatom Phaeodactylum tricornutum, the red alga Cyanidioschyzon merolae and

the green alga Chlamydomonas reinhardtii (Radakovits et al., 2012). LPAT gene

identified in Thalassiosira pseudonana is 27% homologous to the LPAT of the yeast

Saccharomyces cerevisiae (Chen et al., 2010).

In oleaginous yeasts and fungi storage lipid turnover typically occurs after the

depletion of the carbon source in the culture medium or, in general, under carbon-

limiting conditions (Holdsworth and Ratledge, 1988; Aggelis and Sourdis, 1997;

Papanikolaou et al., 2004; Fakas et al., 2007). Respectively, in microalgae under light

starvation conditions, storage material (both sugar and lipid) is degraded to support

algal growth, although a conversion of sugar to lipids was observed in the beginning

of the degradation process in Chlorella sp. (Bellou and Aggelis, 2012).

3.2. PUFAs biosynthesis

Τhe synthesis of long chain unsaturated fatty acids requires the presence of specific

elongases and desaturases, which act primarily on palmitic, stearic and oleic acids.

Fatty acid elongation occurs in both plastids and ER (Ohlrogge and Browse, 1995;

Kunst and Samuels, 2009) and requires acyl-CoA and malonyl-CoA as substrates plus

1 ATP and 2 NADPH molecules per C2-unit elongation of the carbon chain. Fatty

acid elongase is a complex consisted of four subunits namely ß-ketoacyl-CoA

synthase (KCS), ß-ketoacyl-CoA reductase, ß-hydroxyacyl-CoA dehydratase and

enoyl-CoA reductase, which are similar to those found in type II FAS identified in

Chlamydomonas reinhardtii (Yu et al., 2011a; Baba and Shiraiwa, 2013). KCSs are

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divided into the “elongase of very long-chain fatty acid” (ELOVL) contributing to

sphingolipid biosynthesis and the “fatty acid elongase” (FAE) working for TAGs or

wax biosynthesis (Venegas-Calerón et al., 2010; Khozin-Goldberg and Cohen, 2011).

In the plastid, the fatty acids are used to produce lysophosphatidic acid and

phosphatidic acid (PA) via the action of plastidial acyltransferases. The PA and its

derivative product diacylglycerol (DAG) may act as precursors for the synthesis of

plastidial membrane structural lipids (Ohlrogge and Browse, 1995; Fan et al., 2011).

Instead, the fatty acids are transferred to the ER and used to acylate G3P, reaction

mediated by ER-localized acyltransferase isoforms. In contrast to those produced in

plastids, the PA and DAG produced in the ER are used for the synthesis of both

membrane and storage (mainly TAGs) lipids (Ohlrogge and Browse, 1995).

The two types of elongases are referred to as ELOVL, commonly found in

yeast, fungi and animal cells, and as FAE, found in plants. For the synthesis of the

very long-chain PUFAs, such as arachidonic (ARA), eicosapentaenoic (EPA) and

docosahexaenoic (DHA) acids, which are common in the majority of marine species

including microalgae such as Phaeodactylum tricornutum, Nannochloropsis salina,

Nannochloropsis gaditana, Isochrysis galbana, Pavlova salina, Tetraselmis sp., etc.,

the ELOVL is required, whereas FAE activity has not been shown in microalgae so

far (Baba and Shiraiwa, 2013). However, Ouyang et al. (2013) suggest that a FAE

may exist in the microalga Myrmecia incise, the active region of which is exposed in

the cytosolic side of the ER membrane, which may explain why arachidonic acid is

synthesized in the cytoplasm instead of the chloroplast as it was previously

demonstrated by Bigogno et al. (2002). Several elongase-encoding genes implicated

in PUFAs synthesis have been characterized in various species, such as in Pavlova

lutheri (Pereira et al., 2004) and Pyramimonas cordata (Petrie et al., 2010a).

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Desaturases are specialized in the location, number and stereochemistry of

double bonds in fatty acids (Heinz 1993; Pereira et al. 2003). The implication of the

desaturases in the biosynthesis of the very long chain PUFAs has been extensively

reviewed not only in heterotrophic microorganisms but in microalgae as well (Certik

and Shimizu, 1999; Wallis et al., 2002; Pereira et al., 2003; Guschina and Harwood,

2006; Harwood and Guschina, 2009; Moellering et al., 2010; Khozin-Goldberg and

Cohen, 2011). The genes encoding for Δ4, Δ5 and Δ6 desaturases, implicated in DHA

synthesis, have been characterized in Thalassiosira pseudonana (TpDESI, TpdESO

and TpDESK) (Tonon et al., 2005). Similarly, a new desaturase-encoding gene

(IgD4), responsible for the conversion of docosapentaenoic acid (DPA) into DHA and

of adrenic acid into DPA, was found in Isochrysis strains. Genes PsD4Des, PsD5Des

and PsD8Des were identified also in Pavlova salina and were shown to encode for

Δ4, Δ5 and 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 microalga Parietochloris incise

and found to be functionally similar to ∆6 PUFAs elongase-encoding genes of other

species (Iskandarov et al., 2009). Identification of Δ6 and Δ4 desaturase-encoding

genes from Ostreococcus RCC809 and Δ6 elongase-encoding gene from

Fragilariopsis cylindrus was also reported by Vaezi et al. (2013), while Zauner et al.

(2012) identified in Chlamydomonas a gene encoding for Δ4 desaturase.

3.3. Genetic engineering for directing metabolism towards lipid synthesis

Many techno-economic analyses suggest that the most critical factor affecting the

production cost of microalgal lipid in both open pond and PBR systems is the lipid

content in algal cells followed by the specific growth rate (Davis et al., 2011). For

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instance, in the open pond case an increase of the lipid content in the algal cell from

25 to 50% results in a cost reduction of 4$/gal, which corresponds to 46% of the total

production cost. On the other hand, a decrease of the lipid content from 25 to 12.5%

increases the production cost 8$/gal (around 100%). The effect of the lipid content on

the production cost becomes more dramatic when a PBR is used instead of an open

pond. The specific growth rate also considerably affects the lipid production cost but

in a lesser extent than the lipid content (Davis et al., 2011). Therefore, it is not

surprising the amout of effort focusing on the construction of microalgal strains able

to grow fast and synthesize large amounts of lipids having a suitable fatty acid

composition.

For improving growth rate, efforts have been focused on the construction of

strains with an enhanced photosynthetic efficiency (see review of Stephenson et al.,

2011). Although photosynthetic efficiency obviously affects lipid synthesis as well,

particular strategies have been developed to improve the liposynthetic machinery (Qin

et al., 2012). These strategies target the overexpression of proteins that are involved in

the earlier steps of fatty acid synthesis, increasing in this way the availability of

precursor molecules, such as acetyl-CoA and malonyl-CoA. For example, increased

ACC expression may stimulate lipid synthesis. A complementary plan would be the

repression of lipid catabolism by down-regulating or inhibiting TAGs hydrolysis

and/or β-oxidation process. Besides, the regulation and/or insertion in microalgae of

specific desaturases and/or elongases, along with the associated FATs, is used to

modify fatty acid profile but, interestingly, may also affect lipid content or the

biosynthesis of particular lipid fractions.

Genetic engineering approaches in microalgae are in their infancy and,

consequently, the initial efforts had relatively low success indicating that a better

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understanding of the algal fatty acid biosynthetic machinery is required (Blatti et al.,

2013). Actually, despite the noteworthy research, the results were not always

auspicious. For example, overexpression of the plastidial ACC gene (ACC1) in the

diatoms Cyclotella cryptica and Navicula saprophila did not improve fatty acid

synthesis, indicating that ACC upregulation alone may not be sufficient and more

than one enzyme may act synergistically to boost lipid biosynthesis (Dunahay et al.,

1996; Mühlroth et al., 2013). Another obvious targed for improving lipid biosynthesis

is to manipulate FAS that is ACP-dependent (Sirevag and Levine, 1972). Therefore, a

way to enhance lipid biosynthesis is to focus on protein-protein interactions, i.e.

between the ACP and FAT, as Radakovits et al. (2011) and Blatti et al. (2012)

reported. However, the findings showed that FAS affected fatty acid composition but

not lipid accumulation, which indicate that the knowledge on FAS manipulation for

enhanced lipid production is in initial stage.

Considering that the overexpression of genes involved in fatty acid synthesis

had low success, research was focused on other enzymes implicated in acylglycerols

(both storage and structural) biosynthesis. Nevertheless, overexpression of DGAT

genes (CrDGAT2a, CrDGAT2b, and CrDGAT2c) in the model microalga

Chlamydomonas reinhardtii did not lead to increased lipid accumulation (La Russa et

al., 2012). Deng et al. (2012) studying five CrDGAT2 homologous genes in

Chlamydomonas reinhardtii, found that each gene affects diversely lipid

accumulation pattern. Specifically, RNAi silencing of CrDGAT2-1 or CrDGAT2-5

resulted in a significant decrease in lipid content, whereas transformants harboring

CrDGAT2-4 exhibited increase in lipid content. No significant changes in lipid

content were observed when CrDGAT2-2 or CrDGAT2-3 were silenced. On the

contrary, overexpression of a type 2 DGAT in Phaeodactylum tricornutum increased

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TAG accumulation by 35% (Niu et al., 2013). Concerning the effect of heterologous

expression of other implicated in lipogenesis enzymes in Chlorella minutissima, such

as the yeast-derived GPAT, LPAT, PAP- phosphatidic acid phosphatase, DGAT,

G3PDH, the overexpression of single genes had limited effect on the TAG synthesis,

whereas a combination of all the five genes in a unique construct resulted in a two-

fold increase of TAG content (Hsieh et al., 2012; Klok et al., 2014).

The tendency of oleaginous microalgae to synthesize, apart from lipids,

significant amounts of other energy-rich compounds such as starch in processes

competitive to lipogenesis (Bellou and Aggelis, 2012), worths attention since the lipid

yield could be considerably increased by blocking competitive metabolic pathways.

Indeed, Ramazanov and Ramazanov (2006) managed to increase lipid content by 50%

using a starchless mutant of Chlorella pyrenoidosa, while Wang et al. (2009) reported

30-fold increase of lipid bodies by number and size in a starchless mutant of

Chlamydomonas reinhardtii. Similarly, Work et al. (2010) reported that the genetic

blockage of starch synthesis in the sta6 and sta7-10 mutants of C. reinhardtii

increased lipid content under nitrogen deprivation conditions. However, Siaut et al.

(2011) suggested absence of negative correlation between starch reserves and lipid

content when comparing mutants along with the various wild-type strains.

Approaches to increase lipid accumulation by suppressing the β-oxidation

pathway have been successful in the case of plants and yeasts. In microalgae, this kind

of gene suppression could be only accomplished by random mutagenesis or through

the use of RNA silencing as reported by Radakovits et al. (2010). Recently, research

in diatoms showed that most of the implicated lipases were downregulated during

growth under nitrogen deprivation, resulting in TAG accumulation (Yang et al.,

2013). Similarly, targeted knockdown of a multifunctional enzyme

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(lipase/phospholipase/acyltransferase) increased lipid content without affecting

growth in the diatom Thalassiosira pseudonana (Trentacoste et al., 2013).

Both elongases and desaturases, are responsible for PUFAs biosynthesis, and

are not implicated in the lipid accumulation process. However, since PUFAs are

highly desirable molecules, changes in unsaturation profiles by introducing or

regulating desaturases is a major target. Although, several elongase- and desaturase-

encoding genes have been characterized in microalgae, engineering trials are still in

the beginning. Several efforts have been focused on the modification of the fatty acid

composition (i.e. to increase the omega-3 or omega-6 fatty acid content) but they have

been performed mostly in transgenic plants (Dehesh et al., 2001; Opsahl-Ferstad et

al., 2003; Stoll et al., 2005; Graham et al., 2007; Napier, 2007). Recently, Hamilton et

al. (2014) managed to increase 8 folds the DHA content by expressing the

heterologous Δ5 elongase from the picoalga Ostreococcus tauri in the diatom

Phaeodactylum tricornutum. Surprisingly, lipid content increased up to 65% and EPA

synthesis by 58% in Phaeodactylum tricornutum when an annotated Δ5 desaturase

gene (PtD5b) was cloned and overexpressed (Peng et al., 2014). Likewise, a

Chlamydomonas gene encoding for Δ4 desaturase has been found to affect the

biosynthesis of specific lipid fractions since reduced levels of this enzyme led to

lower amounts of monogalactosyldiacylglycerol (MGDG), while its overproduction

increased both the levels of 16:4 acyl groups in the cell extracts and the total amount

of MGDG (Zauner et al., 2012). Enhanced production of PUFAs was also shown in

yeast and plants when microalgal desaturases were introduced to either of those

organisms. Specifically, Δ5 desaturases of the microalgae Ostreococcus tauri and

Ostreococcus lucimarinus were functionally expressed in an engineered

Saccharomyces cerevisiae strain resulting in the production of both ARA and EPA

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(Tavares et al., 2010). Similarly, Petrie et al. (2010b) reported production of 26%

EPA in plant leaf TAGs upon introduction of a newly-identified Δ6 desaturase-

encoding gene from the marine microalga Micromonas pusilla.

Beside desaturases, FATs also affect fatty acid composition of microalgal

lipid. In particular, plant derived FATs (i.e. 12:0- and 14:0-specific FATs from

Umbellularia californica and Cinnamomum camphora, respectively) were recently

successfully engineered into the diatom Phaeodactylum tricornutum in order to alter

the fatty acid composition and the obtained results showed redirection of the fatty acid

synthesis towards the biosynthesis of C12:0 and C14:0 (Radakovits et al., 2011).

Contrary to plant derived FATs, microalgae derived FATs show no fatty acid

specificity. As a result, when endogenous Phaeodactylum tricornutum FAT was

overexpressed it did not result in an altered fatty acid composition, while

overexpression of endogenous Chlamydomonas reinhardtii FAT resulted in short-

circuiting of fatty acids (Gong et al., 2011; Blatti et al., 2012).

4. Biotechnological perspectives of microalgal lipids

The production of microalgal lipids (intended for either as source of PUFAs or as

feedstock for biodiesel production) can be performed through specific processes (i.e.

planned for this purpose) or in combination with the production of other microalgal

metabolic products having pharmaceutical and/or nutritional interest, or even may

arise by exploiting the algal biomass produced during wastewater treatment.

An evaluation of the various systems used for microalgal oil production is

illustrated in Table 1. The majority of microalgal/cyanobacterial production in

Australia, Israel and Japan currently occurs in open ponds (Spolaore et al., 2006;

Ratledge and Cohen, 2008) obviously thanks to low capital investment and operating

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cost of this system. Under the best conditions, scientifically documented peak

biomass productivities in open cultivation systems do not surpass 75-100 ton/ha.yr

(Benemann et al., 1987). Therefore, algal production systems need to be improved in

order to become more competitive and economically viable.

Closed type PBRs (i.e. tubular, panel) are employed for the commercial

production of pigments (i.e. beta-carotene, astaxanthin) by Dunaliella salina and

Haematococcus pluvialis (Spolaore et al., 2006; Ratledge and Cohen, 2008). These

types of reactors can be used for the production of lipids containing PUFAs, since

their relatively high cost can be covered by the high price of these products.

Other than autotrophically, microalgae can be cultivated heterotrophically or

mixotrophically, as well. Mixotrophic cultivation seems to be the most promising

approach since in this case microalgae utilize both phototrophic and heterotrophic

pathways concurrently (Mata et al., 2010; Lam and Lee, 2012). Particularly, Perez-

Garcia et al. (2011) reported that the specific growth rate of mixotrophically grown

microalgae could be estimated as the sum of the specific growth rates of cells grown

under phototrophic and heterotrophic conditions. Further, growth under mixotrophic

conditions could overcome problems related to light invasion. However, this kind of

cultivation system meets also several restrictions mainly due to possible

contaminations, which may be crucial for algal growth. Thus, closed type PBRs are

preferred over open ponds. The option of sterilizing the bioreactors in order to ensure

aseptic conditions might increase the whole cost of the process, which can be only

potentially overcome by the significantly higher yields obtained in this type of culture

(comparable to those obtained by heterotrophic oleaginous microorganisms) and/or

the high value of the target product. Several microalgal strains (i.e. Chlamydomonas

globosa, Chlorella protothecoides, Chlorella sp., Chlorella vulgaris, Chlorella

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zofingiensis, Chlorella minutissima, Haematococcus pluvialis, Nannochloropsis sp.,

Phaeodactylum tricornutum, Rhodomonas reticulate, Scenedesmus bijuga, Spirulina

platensis, etc.) are cultivated under heterotrophic or mixotrophic conditions in large

scale for the production of lipids rich in PUFAs, pigments and proteins (Kobayashi et

al., 1992; Wen and Chen, 2003; Ceron Garcia et al., 2006; Wood et al., 2009; Lee,

2001; Ip et al., 2004; Cheng et al., 2009; Liang et al., 2009; Bhatnagar et al., 2011;

Cheirsilp and Torpee, 2012). Biomass concentration in the reactor can reach 5-15 g/L,

3-30 times higher than that produced under autotrophic growth conditions. The lipid

content of biomass grown under heterotrophic or mixotrophic growth conditions is

also much higher reaching up to 50% or more in the dry biomass.

Besides cultivation an important difficulty encountered in commercial

microalgal applications is the harvest of biomass, which presents several

technological and economic complications. Biomass harvesting requires the

separation of solid from liquid and is a process covering around 30% of the total

production cost (Brennan and Owende, 2010). Currently there are various harvesting

methods, including flocculation or coagulation, flotation, filtration, sedimentation and

centrifugation (see review Chen et al., 2011a). The choice of the appropriate method

should be considered according to the culture cell density, the size of the microalgal

cells, the target product, etc. Among the harvesting methods mentioned above,

filtration has been reported as an efficient and cost-effective one (Molina Grima et al.,

2003; Zhang et al., 2010), with the vibrating screen filter and the microstainer to be

the most popular devices (Chen et al., 2011a). In large-scale operations, mechanical

harvesters, such as continuous belts, are studied and/or already used by various

companies (Christenson and Sims, 2011). Alternatively, the harvesting step may be

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skipped, e.g. by performing facilitated lipid extraction in the whole culture, which

reduces the process cost and enables microalgal production economically feasible.

4.1. Occurrence of PUFAs in microalgal lipids and industrial applications

Microalgae belonging to different classes may have a particular fatty acid

composition that is generally recognized as group specific. Diatoms

(Bucillariophyceae) are able to synthesize high amounts of palmitoleic acid (C16:1)

and EPA, whereas some mutants may also produce ARA (Sriharan et al., 1991;

Schneider et al., 1995). Chrysophyceae (known as golden algae) produce a wide

variety of fatty acids such as C16:1, EPA and DHA. Among them, high proportions of

PUFAs have been reported for Isochrysis strains (Molina Grima et al. 1993;

Tatsuzawa and Takizawa 1995). Gyrodinium and Crypthecodinium and other genera

belonging to Dinophyceae or Dinoflagellates have been characterized 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 fatty

acids (Regnault et al., 1995), include the most popular producers of long chain

PUFAs, such as alpha-linolenic acid (ALA), EPA and DHA (Yongmanitchai and

Ward, 1991; Pettitt and Harwood, 1989; Reitan et al., 1994; Mendes et al., 2005; Patil

et al., 2007; Makri et al., 2011; Bellou et al., 2012; Huang et al., 2013).

Strains with special fatty acid biosynthetic capacity, and therefore of industrial

interest, belong to Chlorella minutissima, having high PUFAs content (Seto et al.,

1984), and Schizochytrium sp., considered one of the best sources of DHA (with a

content of around 40% the total lipids), which also synthesizes EPA but in less

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percentages (i.e. 17%) (Kyle, 2001; Doughman et al., 2007). Cryptocodinium cohnii,

Amphidinium sp. and Prorocentrum triestinum are also known as efficient DHA-

producers (Kyle, 2001; Makri et al., 2011), while Porphyridum cruentum and

Nannochloropsis salina are able to synthesize EPA at more than 25% of total lipids

(Cohen et al., 1988; Bellou and Aggelis, 2012). Various other species are able to

produce lipids with high content in PUFAs as summarized in Table 2.

PUFAs, among all fatty acids and even all algal bioactive compounds, attract

attention thanks to their obvious beneficial effect on human health. They have been

implicated in the treatment and prevention of various diseases and disorders,

including inflammatory disease, atherosclerosis, thrombosis, arthritis and a variety of

cancers (Dyerberg and Jorgensen, 1982; Lagarde et al., 1986; Sakai et al., 1990;

Rousseau et al., 2003). EPA and DHA are known to regulate coagulation, lipoprotein

metabolism, blood pressure, endothelial and platelet function. In particular, PUFAs

are important for the growth and performance of retina, brain, reproductive tissues

and for cardiovascular health (Bhakuni and Rawat, 2005; Horrocks and Yeo, 1999).

Moreover, they have an anti-proliferative effect on cultures of epithelial and

bronchopulmonary cells (Moreau et al., 2006), caused myelo-suppression induced by

lead (Queiroz et al., 2003) and improved glycogenesis in diabetic mice (Cherng and

Shih, 2006).

Adame-Vega et al. (2012) stated that microalgae are the main fatty acid

producers in the marine food chain recognizing microalgal PUFAs as the essential

nutrient for production of zooplankton necessary for the first feeding of larvae. This

can explain the fact that large amounts of PUFAs are used in fisheries, which employ

an artificial food chain, for the enrichment of zooplankton (i.e. rotifer Brachionus

plicatilis) that is widely used in the first-feeding of marine fish larvae. Alternatively,

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the microalgal cells are used as carriers for PUFAs transfer when consumed by

rotifers (Brown, 2002; Birkou et al., 2012; Guedes and Malcata, 2012; see also

supplementary material, Video 1), which are then used as prey for fish larvae, and/or

directly fed to fish larvae during a brief period (Figure 1).

The majority of PUFAs synthesized by microalgae are of high

biotechnological interest. Nevertheless, so far DHA is the only algal PUFA

commercially available. Indeed, although EPA can be produced in remarkable

quantities by microalgal strains (i.e. Porphyridium purpureum, Phaeodactylum

tricornutum, Isochrysis galbana, Nannochloropsis sp., etc.) it is not currently

produced in commercial scale because it cannot be economically competitive

compared to other sources (Apt and Behrens, 1999; Belarbi et al., 2000; Spolaore et

al., 2006). Further improvements in biomass and lipid yields and/or the combined

production of PUFAs with other metabolites (see below) could open the market for

more algal PUFAs in the near future.

4.2. Microalgal lipids as feedstock for biodiesel production

With the steady increase of world population and rapid industrial development,

energy consumption has been increased significantly

(http://www.eia.gov/todayinenergy/). The fossil fuels are widely accepted as non-

renewable and unsustainable energy due to depleting resources, price fluctuating and

causing increase of earth’s temperature (Schenk et al., 2008). All these risks render

the development of renewable sources of energy a pressing mission.

Biodiesel arises to be premium alternative for fossil fuel, since several

favorable environmental properties make it an attractive energy source. Specifically,

with biodiesel the CO2 balance is zero and there are not emissions of sulfur

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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

relevant cancer cases (Sharp, 1996).

Currently biodiesel is produced from vegetable oil harvested from many

feedstock plants such as soybeans, rapeseed (Georgogianni et al., 2009), canola (Li et

al., 2009; Thanh et al., 2010), sunflower seeds (Harrington and D’Arcy-Evans, 1985),

corn (Majewski et al., 2009; Bi et al., 2010) and palm (Kalam et al., 2008). About 7%

of global edible vegetable oil supplies were used for biodiesel production in 2007

(Mitchell, 2008). However, extensive use of edible oils may aggravate food supplies

versus fuel issue (Anwar et al., 2010). Alternatively, non-edible vegetable oils

produced by jatropha (Oliveira et al., 2009; Sahoo et al., 2009), karanja (Naik et al.,

2008; Das et al., 2009; Nabi et al., 2009; Sahoo et al., 2009), mahua (Godiganur et al.,

2009) and polanga (Sahoo et al., 2009) can be used, as their fatty acid composition is

suitable for biodiesel production.

On the other hand, microalgae appear to be an excellent biodiesel source

compared to the existing plants, since they are the fastest growing photosynthetic

organisms (Demirbas and Demirbas, 2011). Moreover, they withstand utmost pH and

temperature conditions and use CO2 in their photosynthetic process more efficiently

(Shay, 1993). Microalgae do not need to be cultivated on agricultural areas but

unsuitable agricultural land can be utilized instead, and they can provide extra

biodiesel oil than oilseed crops while using minimum water and mainland (Sheehan et

al., 1998). Moreover, lipid productivity reported for many microalgae greatly exceeds

the oil productivity of the best producing oil crops, demonstrating that algae give the

maximum biodiesel yield and thus they may be able to produce up to 200 times the

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amount of oil per unit of surface compared to soybeans (Chisti, 2007). These

properties make microalgae the most promising organisms on earth that have the

potential to displace the petroleum-based diesel fuel completely without adversely

affecting supply of food and other crop products.

The algal biofuel technology is still in its infancy and currently there are no

industrialized systems producing algal oil on the scale required for biodiesel

production (Veal et al., 2013). Although the production of biofuels from algal

biomass is technically feasible, there is a need for great efforts, in order to make

feasible the development of economically viable large-scale algal biofuels enterprises.

Actually, despite the algal advantages over all other organisms concerning biodiesel

production, up to now algal biodiesel is more expensive than fossil diesel. Much more

productive strains should be employed (i.e. having higher growth rates and able to

accumulate >50% lipids), while technical restrictions such as the high harvesting cost

and the extremely large area of lagoons needed for biodiesel production, which is not

available (Ratledge and Cohen, 2008), should be faced.

4.3. Combined production of microalgal lipids

A direct reduction of the production cost of microalgal lipids can be achieved by

combining lipid production with other applications. The concept of combined lipid

production is illustrated in Figure 3. Actually, microalgae are used in various

commercial applications (i.e. in the enhancement of nutritional value of food and

animal feed, in aquaculture and pharmaceutical industry, etc.) (Spolaore et al, 2006;

Birkou et al., 2012). Besides PUFAs, compounds such as beta-carotene and

polysaccharides, which are produced commercially by various species, provide a

strong role in manufacturing some therapeutic supplements that comprise an

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important market in biotechnological industries (Priyadarshani and Rath, 2012). Such

properties also enable the use of microalgae in the commercial production of

cosmetics (Spolaore et al, 2006; Priyadarshani and Rath, 2012).

Microalgae have also applications in environmental biotechnology since they

can be used for bioremediation of wastewater and to monitor environmental toxicants.

4.3.1. Combined production of lipids and pharmaceutical products

The microalgae have gained significant attention as natural factories and rich sources

of novel and potential bioactive molecules of interest for 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

both processes. Hydrophobic compounds can be extracted simultaneously with lipids

and then purified, while hydrophilic compounds such as proteins and sugars can be

extracted from the defatted biomass.

Natural cosmeceuticals from algae have become a major counterpart for

superficial application on to the human skin (Spolaore et al., 2006; Raja et al., 2008).

Algal proteins or derivatives are used in skin repair and healing products (Hagino and

Masanobu, 2003). Moreover, the algal cosmetics 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, already used in lipid

production, are also well known in the skin care market for their hydrophilic extracts

(Stolz and Obermayer, 2005). Additionally, diatoms that produce polar lipids rich in

PUFAs along with carotenoids, phytosterols, vitamins, antioxidants, contribute to the

health benefits of the produced oils (Li et al., 2014). The protein extract from blue-

green algae, which include phycoerythrin, possesses many bioactivities, i.e. anti-

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inflammatory, hepetoprotective, antitumor, antiviral and neuroprotective (Bermejo et

al., 2002; Kim et al., 2008; Sekar and Chandramohan, 2008). Furthermore, Chlorella

vulgaris extract stimulates collagen synthesis in the skin and is therefore used for

wrinkle reduction 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 compounds possess

antifungals, antiprotozoal, antiviral, neuroprotective, antiplasmodial, antimicrobial,

anti-inflammatory and antioxidant properties (Kellan and Walker, 1989; Ozemir et al.,

2004; Ghasemi et al., 2004; Mayer and Hamann, 2005; Herrero et al., 2006; Cardozo

et al., 2007; Mendiola et al., 2007; Sekar and Chandramohan, 2008). Microalgal

pigments are very interesting compounds as they are safe, eco-friendly and have a

wide range of therapeutic uses including prevention and treatment of acute and

chronic diseases, rheumatoid arthritis, atherosclerosis, neurological disorders, cataract

and muscular dystrophy (Sies and Stahl, 2004). Daily consumption of microalgae

derived astaxanthin may protect body tissues from oxidative damage as this might be

a practical and beneficial strategy in health management, since it has been suggested

that astaxanthin has a free radical fighting capacity worth 500 times that of vitamin E

( ufoss et al., 2005). Additionally, cytotoxic, pro-apoptotic and anti-proliferative

effects were reported for a large number of microalgal pigments (such as phycobilins,

chlorophylls, epoxycars derivatives) when applied at very low concentrations

(Nishino et al., 1992; Murakami et al., 2002; Konishi et al., 2006; Yoshida et al.,

2007; Sugawara et al., 2007; Sugawara et al., 2009; Shaker et al., 2010; Pasquet et.,

2011). Although the natural algal carotenoids are more expensive than the synthetic

ones, at least natural beta-carotene has specific physical properties that make it

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superior to the synthetic one (Guerin et al., 2003; Garcia-Gonzalez et al., 2005).

Among the various microalgal species, the oleaginous Dunaliella salina is the

preferred organism for beta-carotene production, since it can accumulate up to 14% of

this pigment in dry weight (Metting, 1996). Furthermore, phycobiliproteins (including

phycocyanin, phycoerythrocyanin and allophycocyanin) have been extracted from

various marine algae (Niu et al., 2006). Among them Porphyridium cruentum and

Synechococcus spp. are currently used in large scale for phycobiliprotein production

(Viskari and Colyer, 2003).

Recently, there is increased interest for the fucoxanthin (fuco), a microalgal

cytotoxic pigment, owing to its strong pro-apoptotic, anti-proliferative, and cytotoxic

activities (Nishino et al., 1992; Ishikawa et al., 2008; Yamamoto et al., 2011; Yu et

al., 2011b; Heydarizadeh et al., 2013). 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 polymerases (Murakami, et al.,

2002). Odontella aurita, a microalga containing in its lipids EPA in high

concentrations (28% of total lipids), has been cultivated in open ponds for commercial

purposes. This organism can accumulate fuco in high concentrations (Xia et al., 2013)

and therefore EPA production could be combined with the production of this valuable

molecule.

Apart from pigment compounds, taurine (an algal peptide) has several

functional and biological applications (Houstan, 2005). Recently, taurine has become

a common component in beverages, foods and nutritional supplements (Dawczynski

et al., 2007). In addition, glycoproteins (lectins), extracted from marine algae, are

considered a type of interesting, for biochemical research, proteins as well and can be

isolated with their carbohydrate moiety. Extracts (or purified peptides) from macro-

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and microalgae are shown to have novel antihypertensive and angiotensin converting

enzyme (ACE) inhibitory activities with minimal or no side effects and can thus be

used as alternatives to synthetic drugs (Suetsuna and Nakano, 2000; Suetsuna and

Chen, 2001; Kim and Wijesekara, 2010). Enzymatic hydrolysates having ACE

inhibitory properties have been extracted from various macroalgae (Athukorala and

Jeon, 2005), the most potent of which were those of Ecklonia cava that is an edible

marine brown algal species found in Japan and Korea. Alternatively, extracts of

Chlorella vulgaris and Arthrospira platensis have potent ACE inhibitory properties

(Sheih et al., 2009), and the particular species are also of interest for their lipids.

Other peptides derived from Chlorella vulgaris have been shown to suppress matrix

metalloproteinase-1 level in human skin fibroblast cells, which is induced by solar

ultraviolet B. The particular inhibition may occur even at the level of gene expression

(Chen et al., 2011b). Furthermore, the biliprotein C-phycocyanin, extracted from

Arthrospira platensis has been successfully used against the human hepatocarcinoma

(HepG2) and chronic myeloid leukemia (K562) cell lines (Subhashini et al., 2004;

Nishanth et al., 2010).

4.3.2. Lipids as co-products of environmental applications

Microalgae are mainly autotrophic microorganisms that are able to fix CO2 from

different sources, such as atmosphere, industrial exhaust gases (e.g. flue gas and

flaring gas) or to use fixed forms of CO2 (e.g. NaHCO3 and Na2CO3). Thanks to these

characteristics, many biotechnological applications are carried out using microalgae in

environmental safety and maintenance, such as bioremediation, bioassay and bio-

monitoring of toxicants (Hoffmann, 1988; Phang et al., 2001; Kirkwood et al., 2003;

Harun et al., 2010).

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Several oleaginous microalgae, interesting for biodiesel feedstock production,

can be used in the treatment of municipal wastewaters. Wang et al. (2010) cultivated

the oleaginous microalga Chlorella sp. on various wastewaters coming from four

different stages of the treatment process in a municipal wastewater treatment plant.

The microalga, in the particular study, was able to grow in wastewaters before

primary settling, after primary settling, after activated sludge tank and those generated

in sludge centrifuge, and simultaneously remove nitrogen, phosphorus, chemical

oxygen demand, metal ions, etc. Similar results were obtained for the microalgae

Halochlorella rubescens, Scenedesmus acutus, Chlorella sorokiniana when grown in

wastewaters autrophically, mixotrophically and/or heterotrophically (Kim et al., 2013;

Mahapatra et al., 2014; Sacristán de Alva et al., 2014; Shi et al., 2014).

The production of algal biomass spontaneously grown in biological treatment

plants depends on the climate. In Greece (Patras) important quantities of biomass are

produced annually in the tanks of final settling (Figure 4a), even if the retention time

is low. This biomass, consisted of macro- and micro-algae (Figure 4b) contains 7-10%

lipids and large quantities of protein, both of interest in the biofuel (biodiesel, biogas,

biohydrogen) production. Fatty acid analysis of lipids synthesized by the microalgal

community showed that the major fatty acid was palmitic acid (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 (around 7.0%) were also detected in several samples

(unpublished data).

Several species can be used for the treatment of toxic waters. A pond system,

which uses Chlorella vulgaris as biological material, showed efficiency in treating

wastewater containing toxic contaminants (Hoffmann, 1988; Phang et. al., 2001). The

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oily biomass produced in such a reactor could be further valorized in the production

of biodiesel.

5. Conclusions

The most significant bottlenecks that limit the production of microalgal oil in

large scale are primarily the restricted lipid synthesis in the microalgal cell and

secondarily the low growth rate of these organisms (Davis et al., 2011). Both

constraints, negatively affecting oil productivity, are of biological origin and therefore

their solutions should be seeked in laboratories of molecular biology and

biochemistry. Genetic engineering of strains with such enhanced performances

(having improved biosynthetic capabilities) is a great challenge for the researchers

working in the field and solutions are expected the next few years. The type II

CRISPR/Cas system, being developed as a powerful genome-editing tool, applicable

to virtually any organism (Hsu et al., 2014) could also be employed for the

construction of oleaginous microalgae with desirable properties. The versatility and

tunable specificity of the particular system make it ideal as a screening tool, targeting

specific groups of genes rather than the whole genome, in search of cells competent to

both grow to high densities and be sufficiently oleaginous. Further, evolution of such

genetically modified cells under restrictive conditions could potentially allow finding

relatively genetically stable strains as well. Nevertheless, it seems that blocking

competitive to lipogenesis pathways and/or inhibiting lipid turnover are more

effective strategies than those dealing with the improvement of the liposynthetic

apparatus.

The commercial production of microalgal PUFAs is currently a much more

attainable target than the production of biodiesel. Actually, the natural sources of

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PUFAs are very limited and therefore microalgae have a great industrial potential. It

is certain that the large number of the current, as well as of the upcoming, applications

of PUFAs will force research towards seeking for effective solutions on crucial

questions related to the biochemical restrictions affecting microalgal biomass and

lipid productivity, and harvesting. More research is needed in all aspects of PUFAs

production including biochemical and molecular work, genome characterization of

model organisms and the isolation and characterization of new oleaginous strains.

Indeed, the introduction of new biological material holding unconventional

biochemical arsenal, e.g. having high light energy conversion yield, can widen the

field and alternative ideas can be generated.

The perspective of algal biodiesel is currently poor due to several biological

and technical restrictions. The production cost of algal biodiesel is currently very high

when compared to that of fossil diesel. The construction of new strains able to

accumulate large lipid quantities and the development of new reactors for efficient

cultivation of microalgae are 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 production of these

valuable compounds in combination with lipid production may increase the financial

viability of both processes. Equally, oleaginous microalgae able to grow on municipal

or industrial wastewaters could be further exploited in the biofuel manufacture.

However, all these processes need to be individually studied for their efficiency and

financial viability.

Acknowledgments

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Financial support was provided by the King Abdulaziz University (Jeddah, Saudi

Arabia).

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Figure legends.

Figure 1: Microalgal biomass production and transfer of polyunsaturated fatty acids

(PUFAs) to man, either through food chain or microalgal supplements consumption.

The individual photos are kindly provided by PLAGTON S.A. (Mytikas, Greece),

Kefish (Kefalonia, Greece) and ALGAE A.C. (Nigrita Serres, Greece).

Figure 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, all modified).

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.

Figure 3: Concept of combined production of lipids and other high added-value

metabolites, such as polysaccharides, proteins, pigments, etc. The individual photos

are kindly provided by ALGAE A.C. (Nigrita Serres, Greece).

Figure 4: The facilities of municipal wastewater treatment located in the city of

Patras (Western Greece). The last-stage settling tank in which micro- and macro-algae

are spontaneously grown is framed in red (a). Nile red staining of lipids in micro- and

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macro-algal cells sampled from the last-stage settling tank. White arrows show the

yellowish lipid bodies (b).

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Table 1: Technical 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 4

Lipid in dry algal

mass (%, wt/wt)

25 25

Algae productivity 25 (g/m2.day) 1.25 (kg/m

3.day)

Capital investment Low High

Ease of scale-up Good Variable (depends on PBR type)

Control of growth

conditions

Low (practically

uncontrolled)

High

Harvesting cost High (low density culture) Low (higher density culture)

Contamination risk High Low

Water use High Low

For lipid production 10 MM gal/yr

Total capital cost

(direct + indirect)

($MM)

390 990

Net operating cost

($MM/yr)

37 55

Total co-product 6 7

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credits ($MM/yr)

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Table 2: Fatty 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-3

C22:6

n-3 References

Chromalveolata

Amphidinium 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

Chlorophyta

Botryococcus 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 and Takizawa, 1996

Chlamydomonas sp. 39.5 2.1 1.5 30.2 2.7 - 22.1 - - - Tatsuzawa and Takizawa, 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., 2011

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 maritime 2 11.8 2.7 0.4 2.1 4.1 3.2 42.6 1.3 - - Zhukova and Aizdaicher, 1995

Dunaliella primolecta 3 21.7 - 0.8 4.3 6.2 1.0 38.8 4.1 - - Viso and Marty, 1993

Dunaliella salina 4 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 tertiolecta 6 10.3 4.0 0.3 1.7 5.2 3.2 38.7 1.3 0.4 - Zhukova and Aizdaicher, 1995

Haematococcus pluvialis* 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 incisa 8 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., 2011

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 viridis 10

15.9 3.3 0.8 4.6 3.1 0.2 15.0 13.3 6.7 - Zhukova and Aizdaicher, 1995

Cyanobacteria

Anabaena 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

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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

Cryptophyta

Chroomonas 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

Dinoflagellata

Gymnodinium kowalevskii 26.7 1.8 8.5 6.5 3.7 - 7.2 15.6 0.1 9.5 Zhukova and Aizdaicher, 1995

Dinophyta

Prorocentrum 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

Haptophyta

Emiliana 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 viridis 12

15.4 19.8 0.4 3.7 1.1 - 2.4 2.9 15.7 7.2 Huang et al., 2013

Heterokontophyta

Asterionella 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 constrictus 13

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 muelleri 14

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

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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 oculata 16

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 weissflogii 19

27.5 0.3 1.4 1.3 - 1.0 - 0.3 3.8 0.1 Viso and Marty, 1993

Myzozoa

Crypthecodinium cohnii 2.7 0.8 2.7 17.8 - - - - - 72.3 Couto et al., 2010

Ochrophyta

Skeletonema costatum 9.4 19.0 2.2 2.6 1.6 - 0.2 2.9 15.4 2.3 Zhukova and Aizdaicher, 1995

Rhodophyta

Porphyridium cruentum 20

33.5 3.0 0.8 0.7 5.7 0.2 - - 37.5 - Cohen et al., 1988

Porphyridium cruentum 21

28.6 1.1 0.8 1.3 8.2 0.3 0.4 - 21.1 - Zhukova and Aizdaicher, 1995

Porphyridium cruentum 22

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%; 16

C20:4n-6, 10.6%; 17

C22:5n-3, 21.3%; 18

C16:3n-3, 18.1%; 19

C14:0, 24.8%; 20

C20:4n-6, 17.3%; 21

C20:4n-6, 27.6; 22

C20:4n-6, 21.9

* calculated from data contained in Damiani et al., 2010.

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Figure 1

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Figure 2

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Figure 3

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Figure 4a

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Figure 4b