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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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References
Adarme-Vega TC, Lim DK, Timmins M, Vernen F, Li Y, Schenk PM. Microalgal
biofactories: a promising approach towards sustainable omega-3 fatty acid
production. Microb Cell Fact 2012;11:96.
Aggelis G, Sourdis J. Prediction of lipid accumulation-degradation in oleaginous
micro-organisms growing on vegetable oils. Anton Leeuw 1997;72(2):159–65.
Ahlgren G, Gustafsson IB, Boberg M. Fatty acid content and chemical composition of
freshwater microalgae. J Phycol 1992;28:37–50.
Amaro HM, Guedes A, Malcata FX. Advances and perspectives in using microalgae
to produce biodiesel. Appl Energ 2011;88(10):3402–10.
An M, Mou S, Zhang X, Zheng Z, Ye N, Wang D, et al. Expression of fatty acid
desaturase genes and fatty acid accumulation in Chlamydomonas sp. ICE-L under
salt stress. Bioresour Technol 2013;149:77–83.
Andrich G, Nesti U, Venturi F, Zinnai A, Fiorentini R. Supercritical fluid extraction
of bioactive lipids from the microalga Nannochloropsis sp. Eur J Lipid Sci
Technol 2005;107(6):381–6.
Andrich G, Zinnai A, Nesti U, Venturi F. Supercritical fluid extraction of oil from
microalga Spirulina (Arthrospira) platensis. Acta Aliment 2006;35(2):195–203.
Antolin G, Tinaut FV, Briceno Y. Optimisation of biodiesel production by sunflower
oil transesterification. Bioresour Technol 2002;83:111–4.
Anwar F, Rashid U, Ashraf M, Nadeem M. Okra (Hibiscus esculentus) seed oil for
biodiesel production. Appl Energy 2010;87:779–85.
Apt KE, Behrens PW. Commercial developments in microalgal biotechnology. J
Phycol 1999;35:215–26.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
36
Athukorala Y, Jeon YJ. Screening for angiotensin-1-converting enzyme inhibitory
activity of Ecklonia cava. J Food Sci Nutr 2005;10:134–9.
Baba M, Shiraiwa Y. Biosynthesis of Lipids and Hydrocarbons in Algae. In:
Dubinsky Z, editor. Photosynthesis. ISBN 978-953-51-1161-0; 2013. p. 331-56.
Barrow C, Shahidi F. Marine nutraceuticals and functional foods. CRC Press, Taylor
& Francis Group; 2008.
Becker EW. Handbook of microalgae culture A. In: Richmond, editor. Microalgae in
human and animal nutrition. Oxford: Blackwell Publishing; 2004. p. 312–51.
Beer LL, Boyd ES, Peters JW, Posewitz MC. Engineering algae for biohydrogen and
biofuel production. Cur Opin Biotechnol 2009;20(3):264–71.
Belarbi H, Molina E, Chisti YA. Process for high yield and scaleable recovery of high
purity eicosapentaenoic acid esters from microalgae and fish oil. Proc Biochem
2000;35:951–69.
Bellou S, Aggelis G. Biochemical activities in Chlorella sp. and Nannochloropsis
salina during lipid and sugar synthesis in a lab-scale open pond simulating
reactor. J Biotechnol 2012;164(2):318–29.
Bellou S, Makri A, Sarris D, Michos K, Rentoumi P, Celik, A, et al. The olive mill
wastewater as substrate for single cell oil production by Zygomycetes. J
Biotechnol 2014a;170:50–9.
Bellou S, Makri A, Triantaphyllidou IE, Papanikolaou S, Aggelis G. Morphological
and metabolic shifts of Yarrowia lipolytica induced by alteration of the dissolved
oxygen concentration in the growth environment. Microbiology 2014b;160:807–
17.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
37
Bellou S, Moustogianni A, Makri A, Aggelis G. Lipids containing polyunsaturated
fatty acids synthesized by Zygomycetes grown on glycerol. Appl Biochem
Biotechnol 2012;166(1):146–58.
Benemann JR, Tillett DM, Weissman JC. Microalgae biotechnology. Trends
Biotechnol 1987;5(2):47–53.
Bermejo RR, Alvarez-Pez JM, Acien Fernandez FG, Molina Grima E. Recovery of
pure B-phycoerythrin from the microalga Porphyridium cruentum. J Biotechnol
2002;93:73–85.
Bhakuni DS, Rawat DS. Bioactive Marine Natural Products. New York: Springer;
2005.
Bhatnagar A, Chinnasamy S, Singh M, Das KC. Renewable biomass production by
mixotrophic algae in the presence of various carbon sources and wastewaters.
Appl Energ 2011;88(10):3425–31.
Bi Y, Ding D, Wang D. Low-melting-point biodiesel derived from corn oil via urea
complexation. Bioresour Technol 2010;101:1220–26.
Bigogno C, Khozin-Goldberg I, Adlerstein D, Cohen Z. Biosynthesis of arachidonic
acid in the oleaginous microalga Parietochloris incisa (Chlorophyceae):
radiolabeling studies. Lipids 2002;37(2):209–16.
Birkou M, Bokas D, Aggelis G. Improving fatty acid composition of lipids
synthesized by Brachionus plicatilis in large scale experiments. J Am Oil Chem
Soc 2012;89(11):2047–55.
Blatti JL, Beld J, Behnke CA, Mendez M, Mayfield SP, Burkart MD. Manipulating
fatty acid biosynthesis in microalgae for biofuel through protein-protein
interactions. PloS one 2012;7(9):e42949.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
38
Blatti JL, Michaud J, Burkart MD. Engineering fatty acid biosynthesis in microalgae
for sustainable biodiesel. Curr Opin Chem Biol 2013;17(3):496–505.
Borges L, Morón-Villarreyes JA, ’Oca MGM, Abreu PC. Effects of flocculants on
lipid extraction and fatty acid composition of the microalgae Nannochloropsis
oculata and Thalassiosira weissflogii. Biomass Bioenerg 2011;35(10):4449–54.
Borowitzka MA. Commercial production of microalgae: ponds, tanks, tubes and
fermenters. J Biotechnol 1999;70:313–21.
Bouaicha N, Chezeau A, Turquet J, Quod JP, Puiseux Dao S. Morphological and
toxicological variability of Prorocentrum lima clones isolated from four locations
in the south–west Indian Ocean. Toxicon 2001;39:1195–1202.
Boyle NR, Page MD, Liu B, Blaby IK, Casero D, Kropat J. Three acyltransferases
and nitrogen-responsive regulator are implicated in nitrogen starvation-induced
triacylglycerol accumulation in Chlamydomonas. J Biol Chem
2012;287(19):15811–25.
Brennan L, Owende P. Biofuels from microalgae–a review of technologies for
production, processing, and extractions of biofuels and co-products. Renew Sust
Energ Rev 2010;14(2):557–77.
Brown MR, Jeffrey SW, Volkman JK, Dunstan GA. Nutritional properties of
microalgae for mariculture. Aquaculture 1997;151(1-4):315.
Brown MR. Nutritional value and use of microalgae in aquaculture. Advances en
Nutrición Acuícola VI. Memorias del VI Simposium Internacional de Nutrición
Acuícola 2002;3:281-292.
Cardozo KHM, Guaratini T, Barros MP, Falcão VR, Tonon AP, Lopes NP, Campos
S, Torres MA, Souza AO, Colepicolo P, Pinto E. Metabolites from algae with
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
39
economical impact. Comp Biochem Physiol C Toxicol Pharmacol 2007;146:60–
78.
Cerón García MC, Camacho FG, Mirón AS, Sevilla JF, Chisti Y, Molina Grima E.
Mixotrophic production of marine microalga Phaeodactylum tricornutum on
various carbon sources. J Microbiol Biotechnol 2006;16(5):689.
Certik M, Shimizu S. Biosynthesis and regulation of microbial polyunsaturated fatty
acid production. J Biosci Bioeng 1999;87(1):1–14.
Chaiklahan R, Chirasuwana N, Lohab V, Bunnagc B. Lipid and fatty acids extraction
from the cyanobacterium Spirulina. Sci Asia 2008;34:299–305.
Chang G, Luo Z, Gu S, Wu Q, Chang M, Wang X. Fatty acid shifts and metabolic
activity changes of Schizochytrium sp. S31 cultured on glycerol. Bioresour
Technol 2013;142:255–60.
Chatzifragkou A, Fakas S, Galiotou‐Panayotou M, Komaitis M, Aggelis G,
Papanikolaou S. Commercial sugars as substrates for lipid accumulation in
Cunninghamella echinulata and Mortierella isabellina fungi. Eur J Lipid Sci
Technol 2010;112(9):1048–57.
Cheirsilp B, Torpee S. Enhanced growth and lipid production of microalgae under
mixotrophic culture condition: effect of light intensity, glucose concentration and
fed-batch cultivation. Bioresour Technol 2012;110:510–6.
Chen C, Yeh K, Aisyah R, Lee D, Chang J. Cultivation, photobioreactor design and
harvesting of microalgae for biodiesel production: A critical review. Bioresour
Technol 2011a; 102(1):71–81
Chen CL, Liou SF, Chen SJ, Shih MF. Protective effects of chlorella-derived peptide
on UVB-induced production of MMP-1 and degradation of procollagen genes in
human skin fibroblasts. Regul Toxicol Pharm 2011b;60:112–9.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
40
Chen JE, Smith AG. A look at diacylglycerol acyltransferases (DGATs) in algae. J
Biotechnol 2012;162(1):28–39.
Chen Q, Zou J, Zheng Z, Xu J. Genes encoding a novel type of
lysophosphatidylcholine acyltransferases and their use to increase triacylglycerol
production and/or modify fatty acid composition. US Patent Application
2010/0016431 A1.
Cheng Y, Zhou W, Gao C, Lan K, Gao Y, Wu Q. Biodiesel production from
Jerusalem artichoke (Helianthus Tuberosus L.) tuber by heterotrophic microalgae
Chlorella protothecoides. J Chem Technol Biotechnol 2009;84:777–81.
Cherng JY, Shih MF. Improving glycogenesis in streptozocin (STZ) diabetic mice
after administration of green algae Chlorella. Life Sci 2006;78:1181–6.
Chisti Y. Biodiesel from microalgae. Biotechnol Adv 2007;25:294–306.
Christenson L, Sims R. Production and harvesting of microalgae for wastewater
treatment, biofuels, and bioproducts. Biotech Adv 2011;29(6):686–702.
Cohen Z, Vonshak A, Richmond A. Effect of environmental conditions on fatty acid
composition of the red alga Porphyridium cruentum: correlation to growth rate. J
Phycol 1988;24(3):328–32.
Colla LM, Bertolina TE, Costab JAV. Fatty acids profile of Spirulina platensis grown
under different temperatures and nitrogen concentrations. Z Naturforsch
2004;59:55–9.
Courchesne NMD, Parisien A, Wang B, Lan CQ. Enhancement of lipid production
using biochemical, genetic and transcription factor engineering approaches. J
Biotechnol 2009;141(1):31–41.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
41
Couto RM, Simões PC, Reis A, Da Silva TL, Martins VH, Sánchez‐Vicente Y.
Supercritical fluid extraction of lipids from the heterotrophic microalga
Crypthecodinium cohnii. Eng Life Sci 2010;10(2):158–64.
Damiani MC, Popovich CA, Constenla D, Leonardi PI. Lipid analysis in
Haematococcus pluvialis to assess its potential use as a biodiesel
feedstock. Bioresour Technol 2010; 101(11):3801–7.
Das LM, Bora DK, Pradhan S, Naik MK, Naik SN. Long-term storage stability of
biodiesel produced from Karanja oil. Fuel 2009;88:2315–8.
Davis MS, Solbiati J, Cronan JE. Overproduction of acetyl-CoA carboxylase activity
increases the rate of fatty acid biosynthesis in Escherichia coli. J Biol Chem
2000;275(37):28593–8.
Davis R, Aden A, Pienkos PT. Techno-economic analysis of autotrophic microalgae
for fuel production. Appl Energ 2011; 88(10):3524–31.
Dawczynski C, Schubert R, Jahireis G. Amino acids, fatty acids, and dietary fibre in
edible seaweed products. Food Chemistry 2007;103:891–9.
Dehesh K, Tai H, Edwards P, Byrne J, Jaworski JG. Overexpression of 3-ketoacyl-
acyl-carrier protein synthase IIIs in plants reduces the rate of lipid synthesis.
Plant Physiol 2001;125(2):1103–14.
Delgado M, De Jonge V, Peletier H. Experiments on the resuspension of natural
microphytobenthos populations. Mar Biol 1991;108:321–8.
Dembitsky VM, Pechenkina-Shubina EE, Rozentsvet OA. Glycolipids and fatty acids
of some seaweeds and marine grasses from the Black Sea. Phytochemistry
1991;30(7):2279–83.
Demirbas A, Demirbas FM. Importance of algae oil as a source of biodiesel. Energy
Convers Manage 2011;52:163–70.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
42
Deng XD, Gu B, Li YJ, Hu XW, Guo JC, Fei XW. The roles of acyl-CoA:
diacylglycerol acyltransferase 2 genes in the biosynthesis of triacylglycerols by
the green algae Chlamydomonas reinhardtii. Mol Plant 2012; 5(4):945–7.
Doughman D, Krupanidhi S, Sanjeeve C. Omega-3 fatty acids for nutrition and
medicine considering microalgae oil as a vegetarian source of EPA and
DHA. Curr Diabetes Rev 2007;3:198–203.
Dufossé L, Galaup P, Yaron A, Arad SM, Blanc P, Chidambara Murthy KN,
Ravishankar GA. Microorganisms and microalgae as sources of pigments for
food use: a scientific oddity or an industrial reality? Trends Food Sci Technol
2005;16(9):389–406.
Dunahay TG, Jarvis EE, Dais SS, Roessler PG. Manipulation of microalgal lipid
production using genetic engineering, Appl Biochem Biotechnol
1996;57/58:223–31.
Dyerberg J, Jorgensen KA. Marine oils and thrombogenesis. Prog Lipid Res
1982;21:255–69.
Fakas S, Galiotou-Panayotou M, Papanikolaou S, Komaitis M, Aggelis G.
Compositional shifts in lipid fractions during lipid turnover in Cunninghamella
echinulata. Enzyme Microb Technol 2007;40(5):1321–7.
Fakas S, Papanikolaou S, Galiotou‐Panayotou M, Komaitis M, Aggelis G. Organic
nitrogen of tomato waste hydrolysate enhances glucose uptake and lipid
accumulation in Cunninghamella echinulata. J Appl Microbiol
2008;105(4):1062–70.
Fan J, Andre C, Xu C. A chloroplast pathway for the de novo biosynthesis of
triacylglycerol in Chlamydomonas reinhardtii. FEBS Letters 2011;585(12):1985–
91.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
43
García-Garibay ML, Gómez-Ruiz AE, Bárzana E, Cruz-Guerrero. Single-Cell
Protein. Algae. In: Caballero B, Trugo L, Finglas P, editors. Encyclopedia of
Food Sciences and Nutrition (Second Edition). London: Elsevier Science Ltd.;
2003. p. 5269–76.
García-González M, Moreno J, Manzano JC, Florencio FJ, Guerrero MG. Production
of Dunaliella salina biomass rich in 9-cis-β-carotene and lutein in a closed
tubular photobioreactor. J Biotechnol 2005;115:81–90.
Gema H, Kavadia A, imou , Tsagou V, Komaitis M, Aggelis G. Production of γ-
linolenic acid by Cunninghamella echinulata cultivated on glucose and orange
peel. Appl Microbiol Biotechnol 2002;58(3):303–7.
Georgogianni KG, Katsoulidis AP, Pomonis PJ, Kontominas MG. Transesterification
of soybean frying oil to biodiesel using heterogeneous catalysts. Fuel Process
Technol 2009;90:671–6.
Ghasemi Y, Yazdi MT, Shafiee A, Amini M, Shokravi S, Zarrini G. Parsiguine, a
novel antimicrobial substance from Fischerella ambigua. Pharm Biol
2004;42:318–22.
Godiganur S, Murthy CHS, Reddy RP. 6BTA 5.9 G2-1 Cummins engine performance
and emission tests using methyl ester mahua (Madhuca indica) oil/diesel blends.
Renew Energy 2009;34:2172–7.
Gong Y, Guo X, Wan X, Liang Z, Jiang M. Characterization of a novel thioesterase
(PtTE) from Phaeodactylum tricornutum. J Basic Microbiol 2011;51:666–72.
Graham IA, Larson T, Napier JA. Rational metabolic engineering of transgenic plants
for biosynthesis of omega-3 polyunsaturates. Curr Opin Biotech 2007;18(2):142–
7.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
44
Greenwell HC, Laurens LML, Shields RJ, Lovitt RW, Flynn KJ. Placing microalgae
on the biofuels priority list: a review of the technological challenges. J R S
Interface 2010;7(46):703–26.
Griffiths MJ, van Hille RP, Harrison ST. Lipid productivity, settling potential and
fatty acid profile of 11 microalgal species grown under nitrogen replete and
limited conditions. J Appl Phycol 2012;24(5):989–1001.
Guedes AC, Malcata FX. Nutritional value and uses of microalgae in
aquaculture. Aquaculture 2012;10(1516):59–78.
Guerin M, Huntley ME, Olaizola M. Haematococcus astaxanthin: applications for
human health and nutrition. Trends Biotechnol 2003;21:210–6.
Guschina IA, Harwood JL. Lipids and lipid metabolism in eukaryotic algae. Prog
Lipid Res 2006;45(2):160–86.
Hagino H, Masanobu S. Use of algal proteins in cosmetics. European Patent 2003/1
433 463 B1, Dec. 18.
Hamilton ML, Haslam RP, Napier JA, Sayanova O. Metabolic engineering of
Phaeodactylum tricornutum for the enhanced accumulation of omega-3 long
chain polyunsaturated fatty acids. Metab Eng 2014;22:3-9.
Harel M, Clayton D. Feed formulation for terrestrial and aquatic animals. US Patent
20070082008 2004 (WO/2004/080196).
Harrington KJ, ’Arcy-Evans C. A comparison of conventional and in situ methods
of transesterification of seed oil from a series of sunflower cultivars. J Am Oil
Chem Soc 1985;62:1009–13.
Harun R, Singh M, Forde GM, Danquah MK. Bioprocess engineering of microalgae
to produce a variety of consumer products. Renew Sust Energ Rev
2010;14:1037–47.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
45
Harvey HR, Bradshaw SA, O'Hara S, Eglinton G, Corner ED. Lipid composition of
the marine dinoflagellate Scrippsiella trochoidea. Phytochemistry
1988;27(6):1723–9.
Harwood JL, Guschina IA. The versatility of algae and their lipid metabolism.
Biochimie 2009;91(6):679–84.
Heinz E. Biosynthesis of polyunsaturated fatty acids. In: Moore TSJ, editor. Lipid
metabolism in plants. Boca Raton: CRC Press; 1993. p. 33-89.
Heo SJ, Jeon YJ. Protective effect of fucoxanthin isolated from Sargassum
siliquastrum on UV-B induced cell damage. J Photochem Photobiol B: Biol
2009;95:101–7.
Herrero M, Ibañez E, Cifuentes A, Reglero G, Santoyo S. Dunaliella salina microalga
pressurized liquid extracts as potential antimicrobials. J Food Prot 2006;69:2471–
7.
Heydarizadeh P, Poirier I, Loizeau D, Ulmann L, Mimouni V, Schoefs B, Bertrand M.
Plastids of marine phytoplankton produce bioactive pigments and lipids. Mar
Drugs 2013; 11(9):3425–71.
Hoffmann JP. Wastewater treatment with suspended and non-suspended algae. J
Phycol 1988;34:757–63.
Holdsworth JE, Ratledge C. Lipid turnover in oleaginous yeasts. J Gen Microbiol
1988;134(2):339–46.
Horrocks L, Yeo Y. Health benefits of docosahexaenoic acid (DHA). Pharmacol Res
1999;40:211–25.
Houstan MC. Neutraceuticals, vitamins, antioxidants, and minerals in the prevention
and treatment of hypertension. Progr Cardiovasc Dis 2005;47:396–449.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
46
Hsieh, H-J, Su CH, Chien LJ. Accumulation of lipid production in Chlorella
minutissima by triacylglycerol biosynthesis-related genes cloned from
Saccharomyces cerevisiae and Yarrowia lipolytica. J Microbiol 2012;50:526–534
Hsu PD, Lander ES, Zhang F. Development and applications of CRISPR-Cas9 for
genome Engineering. Cell 2014;157(6):1262–78.
Hu G, Ji S, Yu Y, Wang SA, Zhou G, Li F. Organisms for Biofuel Production:
Natural Bioresources and Methodologies for Improving Their Biosynthetic
Potentials.Adv Biochem Eng Biotechnol 2013;DOI 10.1007/10_2013_245.
Hu Q, Sommerfeld M, Jarvis E, Ghirardi M, Posewitz M, Seibert M, et al. Microalgal
triacylglycerols as feedstocks for biofuel production: perspectives and advances.
Plant J 2008;54(4):621–39.
Huang X, Huang Z, Wen W, Yan J. Effects of nitrogen supplementation of the culture
medium on the growth, total lipid content and fatty acid profiles of three
microalgae (Tetraselmis subcordiformis, Nannochloropsis oculata and Pavlova
viridis). J Appl Phycol 2013;25(1):129–37.
Huerlimann R, de Nys R, Heimann K. Growth, lipid content, productivity, and fatty
acid composition of tropical microalgae for scale-up production. Biotechnol
Bioeng 2010; 107:245–57.
Huerlimann R, Heimann K. Comprehensive guide to acetyl-carboxylases in algae.
Crit Rev Biotechnol 2013;33(1):49–65.
Ip PF, Wong KH, Chen F. Enhanced production of astaxanthin by the green microalga
Chlorella zofingiensis in mixotrophic culture. Process Biochem
2004;39(11):1761–6.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
47
Ishikawa C, Tafuku S, Kadekaru T, Sawada S, Tomita M, Okudaira T, et al. Antiadult
T-cell leukemia effects of brown algae fucoxanthin and its deacetylated product,
fucoxanthinol. Int J Cancer 2008;123:2702–12.
Iskandarov U, Khozin-Goldberg I, Ofir R, Cohen Z. Cloning and characterization of
the∆ 6 polyunsaturated fatty acid elongase from the green microalga
Parietochloris incisa. Lipids 2009;44(6):545–54.
Jensen GS, Ginsberg DI, Drapeau C. Blue-green algae as an immuno-enhancer and
biomodulator. J Am Nutraceut Ass 2001;3:24–30.
Johnson MB, Wen Z. Development of an attached microalgal growth system for
biofuel production. Appl Microbiol Biotechnol 2010;85(3):525–34.
Kalam MA, Hassan M, Hajar R, Yusuf MS, Umar MR, Mahlia I. Palm oil diesel
production and its experimental tests on a diesel engine. In: Pandey A, editor.
Handbook of plant-based biofuels. Taylor & Francis LLC (Boca Raton, FL):
CRC Press; 2008.
Kay RA, Barton LL. Microalgae as food and supplement. Crit Rev Food Sci Nutr
1991;30:555–73.
Kellan SJ, Walker JM. Antibacterial activity from marine microalgae. Br Phycol J
1989; 23:41–4.
Khozin-Goldberg I, Cohen Z. Unraveling algal lipid metabolism: Recent advances in
gene identification. Biochimie 2011;93(1):91–100.
Kim KH. Regulation of mammalian acetyl-coenzyme A carboxylase. Annu Rev Nutr
1997;17(1):77–99.
Kim SK, Ravichandran YD, Khan SB, Kim YT. Prospective of the cosmeceuticals
derived from marine organisms. Biotechnol Bioprocess Eng 2008;13:511–23.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
48
Kim SK, Wijesekara I. Development and biological activities of marine-derived
bioactive peptides: A review. J Funct Foods 2010;2(1):1–9.
Kim TH, Lee Y, Han SH, Hwang SJ. The effects of wavelength and wavelength
mixing ratios on microalgae growth and nitrogen, phosphorus removal using
Scenedesmus sp. for wastewater treatment. Bioresour Technol 2013;130:75–80.
Kirkwood AE, Nalewajko C, Fulthorpe RR. Physiological characteristics of
cyanobacteria in pulp and paper waste-treatment systems. J Appl Phycol 2003;
15:325–35.
Klok AJ, Lamers PP, Martens DE, Draaisma RB, Wijffels RH. Edible oils from
microalgae: insights in TAG accumulation. Trends Biotechnol 2014; in press
DOI: 10.1016/j.tibtech.2014.07.004
Kobayashi M, Kakizono T, Yamaguchi K, Nishio N, Nagai S. Growth and
astaxanthin formation of Haematococcus pluvialis in heterotrophic and
mixotrophic conditions. J Ferment Bioeng 1992;74(1):17–20.
Konishi I, Hosokawa M, Sashima T, Kobayashi H, Miyashita K. Halocynthiaxanthin
and fucoxanthinol isolated from Halocynthia roretzi induce apoptosis in human
leukemia, breast and colon cancer cells. Comp Biochem Phys C 2006;142:53–9.
Kunst L, Samuels L. Plant cuticles shine: advances in wax biosynthesis and export.
Curr Opin Plant Biol 2009;12(6):721–7.
Kyle D. The large-scale production and use of a single-cell oil highly enriched in
docosahexaenoic acid. ACS Symp Ser 2001;788:92–107.
Kyle DJ, Sicotte VJ, Singer JJ, Reeb SE. Bioproduction of docosahexaenoic acid
(DHA) by microalgae. In Kyle DJ, Ratledge C, editors. Industrial Applications of
Single Cell Oils. Champaign IL.: Am. Oil Chemists' Soc; 1992. p. 287–300.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
49
La Russa M, Bogen C, Uhmeyer A, Doebbe A, Filippone E, Kruse O, et al.
Functional analysis of three type-2 DGAT homologue genes for triacylglycerol
production in the green microalga Chlamydomonas reinhardtii. J Biotechnol
2012;162(1):13–20.
Lagarde M, Croset M, Sicard B, Dechavanne M. Biological activities and metabolism
of eicosaenoic acids in relation to platelet and endothelial function. Prog Lipid
Res 1986; 25: 269–71.
Lam MK, Lee KT. Microalgae biofuels: a critical review of issues, problems and the
way forward. Biotechnol Adv 2012;30(3):673–90.
Lang I, Hodac L, Friedl T, Feussner I. Fatty acid profiles and their distribution
patterns in microalgae: a comprehensive analysis of more than 2000 strains from
the SAG culture collection. BMC Plant Biol 2011;11(1):124.
Lee YK. Microalgal mass culture systems and methods: their limitation and potential.
J Appl Phycol 2001;13(4):307–15.
Lei A, Chen H, Shen G, Hu Z, Chen L, Wang J. Expression of fatty acid synthesis
genes and fatty acid accumulation in Haematococcus pluvialis under different
stressors. Biotechnol Biofuels 2012;5(1):1–11.
León R, Martín M, Vigara J, Vilchez C, Vega JM. Microalgae mediated
photoproduction of b-carotene in aqueous–organic two phase systems. Biomol
Eng 2003;20:177–82.
Li E, Xu ZP, Rudolph V. MgCoAl–LDH derived heterogeneous catalysts for the
ethanol transesterification of canola oil to biodiesel. Appl Catalysis B: Environ
2009; 88:42–9.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
50
Li A, Cai J, Pan J, Wang Y, Yue Y, Zhang D. Multi-layer hierarchical array
fabricated with diatom frustules for highly sensitive bio-detection applications. J
Micromech Microeng 2014;24(2):025014.
Liang S, Xueming L, Chen F, Chen Z. Current microalgal health food R & D
activities in China. Hydrobiologia 2004;512:45–8.
Liang Y, Sarkany N, Cui Y. Biomass and lipid productivities of Chlorella vulgaris
under autotrophic, heterotrophic and mixotrophic growth conditions. Biotechnol
Lett 2009;31(7):1043–9.
Liu B, Benning C. Lipid metabolism in microalgae distinguishes itself. Curr Opi
Biotechnol 2013;24(2):300–9.
Mahapatra DM, Chanakya HN, Ramachandra TV. Bioremediation and lipid synthesis
through mixotrophic algal consortia in municipal wastewater. Bioresour Technol
2014, in press.
Majewski MW, Pollack SA, Curtis-Palmer VA. Diphenylammonium salt catalysts for
microwave assisted triglyceride transesterification of corn and soybean oil for
biodiesel production. Tetrahedron Lett 2009;50:5175–7.
Makri A, Bellou S, Birkou M, Papatrehas K, Dolapsakis NP, Bokas D, et al. Lipid
synthesized by micro‐algae grown in laboratory‐ and industrial‐scale bioreactors.
Eng Life Sci 2011;11(1):52–8.
Makri A, Fakas S, Aggelis G. Metabolic activities of biotechnological interest in
Yarrowia lipolytica grown on glycerol in repeated batch cultures. Bioresour
Technol 2010;101(7):2351–8.
Maslova IP, Mouradyan EA, Lapina SS, Klyachko-Gurvich GL, Los DA. Lipid fatty
acid composition and thermophilicity of Cyanobacteria. Russ J Plant Physiol
2004; 51:353–60.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
51
Mata TM, Martins AA, Caetano NS. Microalgae for biodiesel production and other
applications: A review. Renew Sust Energ Rev 2010;14:217–32.
Mayer AMS, Hamann MT. Marine pharmacology in 2001–2002: marine compounds
with antihelmintic, antibacterial, anticoagulant, antidiabetic, antifungal, anti-
inflammatory, antimalarial, antiplatelet, antiprotozoal, antituberculosis, and
antiviral activities; affecting the cardiovascular, immune and nervous systems and
other miscellaneous mechanisms of action. Comp Biochem Physiol C Toxicol
Pharmacol 2005;140:265–86.
Mendes RL, Reis AD, Pereira AP, Cardoso MT, Palavra AF, Coelho JP. Supercritical
CO2 extraction of γ-linolenic acid (GLA) from the cyanobacterium Arthrospira
(Spirulina) maxima: experiments and modeling. Chem Eng J 2005;105(3):147–
51.
Mendiola JA, Torres CF, Martín-Alvarez PJ, Santoyo S, Toré A, Arredondo BO, et al.
Use of supercritical CO2 to obtain extracts with antimicrobial activity from
Chaetoceros muelleri microalga. A correlation with their lipidic content. Eur
Food Res Technol 2007; 224:505–10.
Merchant SS, Kropat J, Liu B, Shaw J, Warakanont J. TAG, You’re it!
Chlamydomonas as a reference organism for understanding algal triacylglycerol
accumulation. Curr opin Biotechnol 2012;23(3):352–63.
Metting FB. Biodiversity and application of microalgae. J Ind Microbiol
1996;17:477–89.
Mitchell D. A note on rising food prices. World bank policy research working paper
no. 4682. Washington: World Bank – Development Economics Group (DEC)
DC; 2008.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
52
Moellering ER, Miller R, Benning C. Molecular genetics of lipid metabolism in the
model green alga Chlamydomonas reinhardtii. In: Wada H, Murata M, editors.
Netherlands: Lipids in Photosynthesis; 2010. p. 139–55.
Molina Grima E, Belarbi EH, Acién Fernández FG, Robles Medina A, Chisti Y.
Recovery of microalgal biomass and metabolites: process options and economics.
Biotechnol Adv 2003;20(7):491–515.
Molina Grima E, Pérez JS, Camacho FG, Sánchez JG, Alonso DL. n-3 PUFA
productivity in chemostat cultures of microalgae. Appl Microbiol Biotechnol
1993;38(5):599-605.
Moreau D, Tomasoni C, Jacquot C, Kaas R, Le Guedes R, Cadoret JP, et al.
Cultivated microalgae and the carotenoid fucoxanthin from Odontella aurita as
potent anti-proliferative agents in bronchopulmonary and epithelial cell lines.
Environ Toxicol Pharmacol 2006;22:97–103.
Morist A, Montesinos JL, Cusido JA, Godia F. Recovery and treatment of Spirulina
platensis cells cultured in a continuous photbioreactor to be used as food. Process
Biochem 2001;37:535–47.
Moseley JL, Gonzalez-Ballester D, Pootakham W, Bailey S, Grossman AR. Genetic
interactions between regulators of Chlamydomonas phosphorus and sulfur
deprivation responses. Genetics 2009;181(3):889–905.
Msanne J, Xu D, Konda AR, Casas-Mollano JA, Awada T, Cahoon EB, et al.
Metabolic and gene expression changes triggered by nitrogen deprivation in the
photoautotrophically grown microalgae Chlamydomonas reinhardtii and
Coccomyxa sp. C-169. Phytochemistry 2012;75:50–9.
Muhling M, Belay A, Whitton BA. Variation in fatty acid composition of Arthrospira
(Spirulina) strains. J Appl Phycol 2005;17:137–46.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
53
Mühlroth A, Li K, Røkke G, Winge P, Olsen Y, Hohmann-Marriott MF, et al.
Pathways of lipid metabolism in marine algae, co-expression network,
bottlenecks and candidate genes for enhanced production of EPA and DHA in
species of Chromista. Marine drugs 2013;11(11):4662–97.
Murakami C, Takemura M, Sugiyama Y, Kamisuki S, Asahara H, Kawasaki M,
Ishidoh T, Linn S, Yoshida S, Sugawara F, Yoshida H, Sakaguchi K, Mizushina
Y. Vitamin A-related compounds, all-trans retinal and retinoic acids, selectively
inhibit activities of mammalian replicative DNA polymerases. Biochim Biophys
Acta 2002;1574:85–92.
Nabi MD, Hoque SNN, Akhter MDS. Karanja (Pongamia Pinnata) biodiesel
production in Bangladesh, characterization of karanja biodiesel and its effect on
diesel emissions. Fuel Process Technol 2009;90:1080–6.
Naik M, Meher LC, Naik SN, Das LM. Production of biodiesel from high free fatty
acid Karanja (Pongamia pinnata) oil. Biomass Bioenergy 2008;32:354–7.
Napier JA. The production of unusual fatty acids in transgenic plants. Annu Rev Plant
Biol 2007;58:295–319.
Nishanth RP, Ramakrishna BS, Jyotsna RG, Roy KR, Reddy GV, Reddy PK, et al. C-
Phycocyanin inhibits MDR1 through reactive oxygen species and
cyclooxygenase-2. Eur J Pharmacol 2010;649:74–83.
Nishino H, Tsushima M, Matsuno T, Tanaka Y, Okuzumi J, Murakoshi M, Satomi Y,
Takayasu J, Tokuda H, Nishino A, Iwashima A. Anti-neoplastic effect of
halocynthiaxanthin, a metabolite of fucoxanthin. Anti-Cancer Drug 1992;3:493–
7.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
54
Niu JF, Wang GC, Tseng CK. Method of large-scale isolation and purification of R-
phycoerythrin from red algae Polysiphonia urceolata Grev. Protein Express Purif
2006;49:23–31.
Niu YF, Zhang MH, Li DW, Yang WD, Liu JS, Bai WB, Li HY. Improvement of
neutral lipid and polyunsaturated fatty acid biosynthesis by overexpressing a type
2 diacylglycerol acyltransferase in marine diatom Phaeodactylum
tricornutum. Mar Drugs 2013;11(11):4558–69.
Ohlrogge J, Browse J. Lipid biosynthesis. Plant Cell 1995; 7:957–70.
Oliveira J, Leite PM, de Souza LB, Mello VM, Silva EC, Rubim JC, Meneghetti
SMP, Suarez PAZ. Characteristics and composition of Jatropha gossypiifolia and
Jatropha curcas L. oils and application for biodiesel production. Biomass
Bioenergy 2009;33:449–53.
Opsahl-Ferstad H-G, Rudi H, Ruyter B, Refstie S, Biotechnological approaches to
modify rapeseed oil composition for applications in aquaculture. Plant Sci
2003;165(2):349–57.
Ouyang LL, Li H, Liu F, Tong M, Yu SY, Zhou ZG. Accumulation of arachidonic
acid in a green microalga, Myrmecia incisa H4301, enhanced by nitrogen
starvation and its molecular regulation mechanisms In: Dumancas GG, Murdianti
BS, Lucas EA, editors. Arachidonic Acid: Dietary Sources and General
Functions. New York: NOVA Science Publishers, Inc; 2013. p. 1–20.
Ozemir G, Karabay NU, Dalay MC, Pazarbasi B. Antibacterial activity of volatile
components and various extracts of Spirulina platensis. Phytother Res
2004;18:754–7.
Papanikolaou S, Aggelis G. Lipids of oleaginous yeasts. Part I: Biochemistry of single
cell oil production. Eur J Lipid Sci Technol 2011;113(8):1031–51.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
55
Papanikolaou S, Sarantou S, Komaitis M, Aggelis G. Repression of reserve lipid
turnover in Cunninghamella echinulata and Mortierella isabellina cultivated in
multiple‐limited media. J Appl Microbiol 2004;97(4):867–75.
Parrish CC, Bodennec G, Sebedio JL, Gentien P. Intra-and extracellular lipids in
cultures of the toxic dinoflagellate, Gyrodinium aureolum. Phytochemistry
1993;32(2):291–5.
Pasquet V, Morisset P, Ihammouine S, Chepied A, Aumailley L, Berard JB, et al.
Antiproliferative activity of violaxanthin isolated from bioguided fractionation of
Dunaliella tertiolecta extracts. Mar Drugs 2011;9:819–31.
Patil V, Källqvist T, Olsen E, Vogt G, Gislerød HR. Fatty acid composition of 12
microalgae for possible use in aquaculture feed. Aquacult Int 2007;15(1):1–9.
Peng KT, Zheng CN, Xue J, Chen XY, Yang WD, Liu JS, Bai W, Li HY. Delta 5
fatty acid desaturase upregulates the synthesis of polyunsaturated fatty acids in
marine diatom Phaeodactylum tricornutum. J Agric Food Chem 2014;
62(35):8773–6.
Pereira S, Leonard A, Huang Y, Chuang L, Mukerji P. Identification of two novel
microalgal enzymes involved in the conversion of the omega3-fatty acid,
eicosapentaenoic acid, into docosahexaenoic acid. Biochem J 2004;384:357–66.
Pereira SL, Leonard AE, Mukerji P. Recent advances in the study of fatty acid
desaturases from animals and lower eukaryotes. Prostag Leukotr Ess
2003;68(2):97–106.
Perez-Garcia O, Escalante FM, de-Bashan LE, Bashan Y. Heterotrophic cultures of
microalgae: metabolism and potential products. Water Res 2011;45(1):11–36.
Petrie JR, Liu Q, Mackenzie AM, Shrestha P, Mansour MP, Robert SS, et al.
Isolation and characterisation of a high-efficiency desaturase and elongases from
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
56
microalgae for transgenic LC-PUFA production. Mar Biotechnol
2010a;12(4):430–38.
Petrie JR, Shrestha P, Mansour MP, Nichols PD, Liu Q, Singh SP. Metabolic
engineering of omega-3 long-chain polyunsaturated fatty acids in plants using an
acyl-CoA Δ6-desaturase with ω3-preference from the marine microalga
Micromonas pusilla. Metab Eng 2010b;12(3):233–40.
Pettitt TR Harwood JL. Alterations in lipid metabolism caused by illumination of the
marine red algae Chondrus crispus and Polysiphonia lanosa. Phytochem
1989;28:3295–300.
Phang SM, Chui YY, Kumaran G, Jeyaratnam S, Hashim MA. High rate algal ponds
for treatment of wastewater: a case study for the rubber industry. In: Kojima H,
Lee YK, editors. Photosynthetic microorganisms in environmental biotechnology.
Hong Kong: Springer-Verlag; 2001. p. 51–76.
Priyadarshani I, Rath B. Commercial and industrial applications of micro algae – A
review. Commercial and industrial applications of micro algae – A review. J
Algal Biomass Utln 2012;3(4):89–100.
Qin S, Lin H, Jiang P. Advances in genetic engineering of marine algae. Biotechnol
Adv 2012; 30(6):1602–13.
Queiroz MLS, Rodrigues APO, Bincoletto C, Figueiredo CAV, Malacrida S.
Protective effects of Chlorella vulgaris in lead-exposed mice infected with
Listeria monocytogenes. Int Immunopharmacol 2003;3:889–900.
Radakovits R, Jinkerson RE, Darzins A, Posewitz MC. Genetic engineering of algae
for enhanced biofuel production. Eukaryot Cell 2010;9(4):486–501.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
57
Radakovits R, Jinkerson RE, Fuerstenberg SI, Tae H, Settlage RE, Boore JL, et al.
Draft genome sequence and genetic transformation of the oleaginous alga
Nannochloropsis gaditana. Nat Commun 2012;3:686.
Radwan SS. Sources of C20-polyunsaturated fatty acids for biotechnological use.
Appl Microbiol Biotechnol 1991;35(4):421–30.
Raja R, Hemaiswarya S, Kumar NA, Sridhar S, Rengasamy R. A perspective on the
biotechnological potential of microalgae. Crit Rev Microbiol 2008;34(2):77–88.
Ramazanov A, Ramazanov Z. Isolation and characterization of a starchless mutant of
Chlorella pyrenoidosa STL‐PI with a high growth rate, and high protein and
polyunsaturated fatty acid content. Phycol Res 2006;54(4):255–9.
Rania MA, Hala MT. Antibacterial and antifungal activity of Cynobacteria and green
Microalgae evaluation of medium components by Plackett-Burman design for
antimicrobial activity of Spirulina platensis. Global J Biotech Biochem
2008;3(1):22–31.
Ratledge C, Cohen Z. Microbial and algal oils: Do they have a future for biodiesel or
as commodity oils? Lipid Technology 2008;20:155–60
Ratledge C. Fatty acid biosynthesis in microorganisms being used for single cell oil
production. Biochimie 2004;86(11):807–15.
Regnault A, Chervin D, Chammai A, Piton F, Calvayrac R, Mazliak P. Lipid
composition of Euglena gracilis in relation to carbon-nitrogen balance.
Phytochemistry 1995;40(3):725–33.
Reitan KI, Rainuzzo JR, Olsen Y. Effect of nutrient limitation on fatty acid and lipid
content of marine microalgae1. J Phycol 1994;30(6):972–9.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
58
Renaud SM, Thinh LV, Lambrinidis G, Parry DL. Effect of temperature on growth,
chemical composition and fatty acid composition of tropical Australian
microalgae grown in batch cultures. Aquaculture 2002; 211:195–214.
Rousseau D, Helies-Toussaint C, Moreau D, Raederstorff D, Grynberg A. Dietary n-3
PUFAs affect the blood pressure rise and cardiac impairments in a
hyperinsulinemia rat model in vivo. Am J Physiol Heart Circ Physiol
2003;285:1294–302.
Sacristán de Alva M, Luna-Pabello VM, Cadena E, Ortíz E. Green microalga
Scenedesmus acutus grown on municipal wastewater to couple nutrient removal
with lipid accumulation for biodiesel production. Bioresour Technol
2013;146:744–8.
Sahoo PK, Das LM, Babu MKG, Arora P, Singh VP, Kumar NR, et al. Comparative
evaluation of performance and emission characteristics of jatropha, karanja and
polanga based biodiesel as fuel in a tractor engine. Fuel 2009;88:1698–707.
Sakai K, Okuzama H, Kon K, Maeda N, Sekiya M, Shiga T, et al. Effects of high γ-
linolenate and linoleate diets on erythrocyte deformability and hematological
indices in rats. Lipids 1990;25:793–7.
Salama ES, Kim HC, Abou-Shanab RA, Ji MK, Oh YK, Kim SH, et al. Biomass,
lipid content, and fatty acid composition of freshwater Chlamydomonas mexicana
and Scenedesmus obliquus grown under salt stress. Bioprocess Biosyst Eng
2013;36(6):827–33.
Samarakoona K, Jeona Y-J. Bio-functionalities of proteins derived from marine algae
-A review. Food Res Int 2012;48(2):948–60.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
59
Schenk PM, Thomas-Hall SR, Stephens E, Marx UC, Mussgnug JH, Posten C, et al.
Second generation biofuels: high-efficiency microalgae for biodiesel production.
Bioenergy Res 2008;1:20–43.
Schneider JC, Livne A, Sukenik A, Roessler PG. A mutant of Nannochloropsis
deficient in eicosapentaenoic acid production. Phytochem 1995;40:807–14
Sekar S, Chandramohan M. Phycobiliproteins as a commodity: Trends in applied
research, patents and commercialization. J Appl Phycol 2008;20:113–36.
Seto A, Wang HL, Hesseltine CW. Culture conditions affect eicosapentaenoic acid
content of Chlorella minutissima. J Am Oil Chem Soc 1984;61(5):892–4.
Shaker KH, Müller M, Ghani MA, Dahse HM, Seifert K. Terpenes from the soft
corals Litophyton arboreum and Sarcophyton ehrenbergi. Chem Biodiver
2010;7:2007–15.
Sharp CA. Emissions and lubricity evaluation of rapeseed derived biodiesel. Prepared
by Montana Department of Environmental Quality, Southwest Research Institute,
USA, 1996:1–57.
Shay EG. Diesel fuel from vegetable oils - status and opportunities. Biomass
Bioenergy 1993;4:227–42.
Sheehan J, Dunahay T, Benemann J, Roessler P. A Look Back at the U.S. Department
of Energy’s Aquatic Species Program – Biodiesel from Algae. Golden, Colorado,
USA: National Renewable Energy Laboratory; 1998.
Sheih IC, Fang TJ, Wu TK. Isolation and characterization of a novel angiotensin-I
converting enzyme (ACE) inhibitory peptide from the algae protein waste. Food
Chem 2009;115:279–84.
Shi J, Podola B, Melkonian M. Application of a prototype-scale Twin-Layer
photobioreactor for effective N and P removal from different process stages of
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
60
municipal wastewater by immobilized microalgae. Bioresour Technol
2014;154:260–66.
Shimoda H, Tanaka J, Shan SJ, Maoka T. Anti‐pigmentary activity of fucoxanthin
and its influence on skin mRNA expression of melanogenic molecules. J Pharm
Pharmacol 2010;62(9):1137–45.
Siaut M, Cuiné S, Cagnon C, Fessler B, Nguyen M, Carrier P, Beyly A, Beisson F,
Triantaphylidès C, Li-Beisson Y, Peltier G. Oil accumulation in the model green
alga Chlamydomonas reinhardtii: characterization, variability between common
laboratory strains and relationship with starch reserves. BMC
Biotechnol 2011;11(1):7.
Sies H, Stahl W. Nutritional protection against skin damage from sunlight. Annu Rev
Nutr 2004;24:173–200.
Silva LM, Lima V, Holanda ML, Pinheiro PG, Rodrigues JA, Lima ME, et al.
Antinociceptive and anti-inflammatory activities of lectin from marine red alga
Pterocladiella capillacea. Biol Pharm Bull. 2010;33(5):830–5.
Sirevag R, Levine RP. Fatty acid synthetase from Chlamydomonas reinhardtii- sites
of transcription and translation. J Biol Chem 1972; 247:2586–91.
Soletto D, Binaghi L, Lodi A, Carvalho JCM, Converti A. Batch and fedbatch
cultivations of Spirulina platensis using ammonium sulphate and urea as nitrogen
sources. Aquaculture 2005;243:217–24.
Spolaore P, Joannis-Cassan C, Duran E, Isambert A. Commercial applications of
microalgae. J Biosci Bioeng 2006;101(2):87–96.
Sriharan S, Bagga D, Nawaz M. The effects of nutrients and temperature on biomass,
growth, lipid production, and fatty acid composition of Cyclotella cryptica
Reimann, Lewin, and Guillard. Appl BiochemBiotechnol 1991;28(1):317–26.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
61
Stephenson PG, Moore CM, Terry MJ, Zubkov MV, Bibby TS. Improving
photosynthesis for algal biofuels: toward a green revolution. Trends Biotechnol
2011; 29(12):615–23.
Stoll C, Lühs W, Zarhloul MK, Friedt W. Genetic modification of saturated fatty
acids in oilseed rape (Brassica napus). Eur J Lipid Sci Technol 2005;107(4):244–
8.
Stolz P, Obermayer B. Manufacturing microalgae for skin care. Cosmetics Toiletries
2005;120:99–106.
Subhashini J, Mahipal SVK, Reddy MC, Reddy MM, Rachamallu A, Reddanna P.
Molecular mechanisms in C-Phycocyanin induce apoptosis in human chronic
myeloid leukemia cell line-K562. Biochem Pharmacol 2004;68:453–62.
Subrahmanyam S, Cronan JE. Overproduction of a functional fatty acid biosynthetic
enzyme blocks fatty acid synthesis in Escherichia coli. J Bacteriol
1998;180(17):4596–602.
Suetsuna K, Chen JR. Identification of antihypertensive peptides from peptic digests
of two microalgae, Chlorella vulgaris and Spirulina platensis. Mar Biotechnol
2001;3:305–9.
Suetsuna K, Nakano T. Identification of antihypertensive peptides from peptidic
digest of wakame (Undaria pinnatifida). J Nutr Biochem 2000;11:450–4.
Sugawara T, Yamashita K, Asai A, Nagao A, Shiraishi T, Imai I, et al. Esterification
ofxanthophylls by human intestinal Caco-2 cells. Arch Biochem Biophys
2009;483:205–12.
Sugawara T, Yamashita K, Sakai S, Asai A, Nagao A, Shiraishi T, et al. Induction of
apoptosis in DLD-1 human colon cancer cells by peridinin isolated from the
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
62
dinoflagellate, Heterocapsa triquetra . Biosci Biotech Biochem 2007;71:1069–
72.
Sydney EB, Sturm W, de Carvalho JC, Thomaz-Soccol V, Larroche C, Pandey A, et
al. Potential carbon dioxide fixation by industrially important microalgae.
Bioresour Technol 2010;101:5892–96.
Tang D, Han W, Li P, Miao X and Zhong J. CO2 biofixation and fatty acid
composition of Scenedesmus obliquus and Chlorella pyrenoidosa in response to
different CO2 levels. Bioresour Technol 2001; 102:3071–6.
Tatsuzawa H, Takizawa E. Changes in lipid and fatty acid composition of Pavlova
lutheri. Phytochemistry 1995;40(2):397–400.
Thanh LT, Okitsu K, Sadanaga Y, Takenaka N, Maeda Y, Bandow H. Ultrasound
assisted production of biodiesel fuel from vegetable oils in a small scale
circulation process. Bioresour Technol 2010;101:639–45.
Tonon T, Harvey D, Larson TR, Graham IA: Long chain polyunsaturated fatty acid
production and partitioning to triacylglycerols in four microalgae.
Phytochemistry 2002, 61:15–24.
Tonon T, Sayanova O, Michaelson LV, Qing R, Harvey D, Larson TR, et al. Fatty
acid desaturases from the microalga Thalassiosira pseudonana. FEBS Journal
2005;272(13):3401–12.
Tran HL, Hong SJ, Lee CG. Evaluation of extraction methods for recovery of fatty
acids from Botryococcus braunii LB 572 and Synechocystis sp. PCC 6803.
Biotechnol Bioprocess Eng 2009;14(2):187–92.
Trentacoste EM, Shrestha RP, Smith SR, Glé C, Hartmann AC, Hildebrand M,
Gerwick WH. Metabolic engineering of lipid catabolism increases microalgal
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
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lipid accumulation without compromising growth. Proc Nat Acad Sci USA
2013;110(49):19748–53.
Tsuzuki M, Ohnuma E, Sato N, Takaku T, Kawaguchi A. Effects of CO2
concentration during growth on fatty acid composition in microalgae. Plant
Physiol 1990; 93:851–6.
Um BH, Kim YS. A chance for Korea to advance algal-biodiesel technology: Review.
J Ind Eng Chem 2009;15:1–7.
Vaezi R, Napier JA, Sayanova O. Identification and functional characterization of
genes encoding omega-3 polyunsaturated fatty acid biosynthetic activities from
unicellular microalgae. Mar Drugs 2013;11(12):5116–29.
Veal MW, Caffrey KR, Chinn MS, Grunden AM. Algae for biofuels-economic and
environmental costs. In: SRAC Publication-Southern Regional Aquaculture
Center, (4310). Stonville: Mississippi State University; 2013. p. 8.
Venegas-Calerón M, Sayanova O, Napier JA. An alternative to fish oils: metabolic
engineering of oil-seed crops to produce omega-3 long chain polyunsaturated
fatty acids. Prog Lipid Res 2010;49(2):108–19.
Vicente G, Martinez M, Aracil J. Integrated biodiesel production: a comparison of
different homogeneous catalysts systems. Bioresour Technol 2004;92:297–305.
Viskari PJ, Colyer CL. Rapid extraction of phycobiliproteins from cultures
cyanobacteria samples. Anal Biochem 2003;319:263–71.
Viso AC, Marty JC. Fatty acids from 28 marine microalgae. Phytochemistry
1993;34:1521–33.
Vonshak A, Torzillo G. Environmental stress physiology A. In: Richmond, editor.
Handbook of Microalgal Culture. Oxford: Blackwell Publishers; 2004. p. 57–82.
ACC
EPTE
D M
ANU
SCR
IPT
ACCEPTED MANUSCRIPT
64
Wallis JG, Watts JL, Browse J. Polyunsaturated fatty acid synthesis: what will they
think of next? Trends Biochem Sci 2002;27(9):467–73.
Wang D, Ning K, Li J, Hu J, Han D, Wang H, et al. Nannochloropsis genomes reveal
evolution of microalgal oleaginous traits. PLoS Genet 2014;10(1):e1004094.
Wang L, Min M, Li Y, Chen P, Chen Y, Liu Y, et al. Cultivation of green algae
Chlorella sp. in different wastewaters from municipal wastewater treatment plant.
Appl Biochem Biotechnol 2010;162:1174–86.
Wang ZT, Ullrich N, Joo S, Waffenschmidt S, Goodenough U. Algal lipid bodies:
stress induction, purification, and biochemical characterization in wild-type and
starchless Chlamydomonas reinhardtii. Eukaryot Cell 2009;8(12):1856–68.
Wen ZY, Chen F. Heterotrophic production of eicosapentaenoic acid by microalgae.
Biotechnol Adv 2003;21(4):273–94.
Wood CC, Petrie JR, Shrestha P, Mansour MP, Nichols PD, Green AG, et al. A
leaf‐based assay using interchangeable design principles to rapidly assemble
multistep recombinant pathways. Plant Biotech J 2009;7(9):914–24.
Work VH, Radakovits R, Jinkerson RE, Meuser JE, Elliott LG, Vinyard DJ, Laurens
LML, Dismukes GC, Posewitz MC. Increased lipid accumulation in the
Chlamydomonas reinhardtii sta7-10 starchless isoamylase mutant and increased
carbohydrate synthesis in complemented strains. Eukaryot Cell 2010;9(8):1251–
61.
Xia S, Wang K, Wan L, Li A, Hu Q, Zhang C. Production, characterization, and
antioxidant activity of fucoxanthin from the marine diatom Odontella aurita, Mar
Drugs 2013;11:2667–81.
ACC
EPTE
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SCR
IPT
ACCEPTED MANUSCRIPT
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Yamaguchi K. Recent advances in microalgal bioscience in Japan, with special
reference to utilization of biomass and metabolites: a review. J Appl Phycol
1997;8:487–502.
Yamamoto K, Ishikawa C, Katano H, Yasumoto T, Mori N. Fucoxanthin and its
deacetylated product, fucoxanthinol, induce apoptosis of primary effusion
lymphomas. Cancer Lett 2011;300:225–34.
Yang ZK, Niu YF, Ma YH, Xue J, Zhang MH, Yang WD, Liu JS, Lu SH, Guan Y, Li
HY. Molecular and cellular mechanisms of neutral lipid accumulation in diatom
following nitrogen deprivation. Biotechnol Biofuels 2013;6(67):1.
Yongmanitchai W, Ward OP. Screening of algae for potential alternative sources of
eicosapentaenoic acid. Phytochem 1991;30:2963–7.
Yoshida T, Maoka T, Das SK, Kanazawa K, Horinaka M, Wakada M, et al.
Halocynthiaxanthin and peridinin sensitize colon cancer cell lines to tumor
necrosis factor-related apoptosis-inducing ligand. Mol Cancer Res 2007;5:615–
25.
Yu RX, Hu XM, Xu SQ, Jiang ZJ, Yang W. Effects of fucoxanthin on proliferation
and apoptosis in human gastric adenocarcinoma MGC-803 cells via JAK/STAT
signal pathway. Eur J Pharmacol 2011b;657:10–9.
Yu WL, Ansari W, Schoepp NG, Hannon MJ, Mayfield SP, Burkart MD.
Modifications of the metabolic pathways of lipid and triacylglycerol production
in microalgae. Microb Cell Fact 2011a;10:91.
Zäuner S, Jochum W, Bigorowski T, Benning C. A cytochrome b5-containing plastid-
located fatty acid desaturase from Chlamydomonas reinhardtii. Eukaryot Cell
2012;11(7):856–63.
ACC
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SCR
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Zhang X, Hu Q, Sommerfeld M, Puruhito E, Chen Y. Harvesting algal biomass for
biofuels using ultrafiltration membranes. Bioresour Technol 2010;101(14):5297–
304.
Zhou XR, Robert SS, Petrie JR, Frampton DM, Mansour PM, Blackburn SI, et al.
Isolation and characterization of genes from the marine microalga Pavlova salina
encoding three front-end desaturases involved in docosahexaenoic acid
biosynthesis. Phytochemistry 2007;68:785–96.
Zhukova NV, Aizdaicher NA. Fatty acid composition of 15 species of marine
microalgae. Phytochemistry 1995;39(2):351–6.
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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