Chapter 1 Introduction

 

CHAPTER 1 

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1.1. Introduction

Benthic marine macroalgae, commonly known as seaweeds are diverse group of

fascinating multicellular photosynthetic forms that grow mostly as attached forms on rocks

in coastal waters. They are harvested and utilized as the sources of food, feed,

phycocolloids, fertilizer, energy, medicines, cosmetics and nutraceuticals besides being used

in biotechnological, bioremediation and aquaculture applications (Holdt and Kraan, 2011;

Gupta and Abu-Ghannam, 2011; Mohamed et al. 2011). They are the first marine organisms

chemically analyzed, with more than 3,600 published articles describing 3,300 secondary

metabolites and are still remained as almost endless source of new bioactive compounds

(Davis & Vasanthi, 2011). An extensive research carried out in this area for the past one

decade has deciphered the bioactive potential of many algal extracts with anti-inflammatory,

cytotoxic, immunosuppressive, antibacterial, anti-plasmodial, antiviral, antifungal, anti-

mutagenic, free radical scavenging, anti-diabetic, anti-hypertensive and antifeedant

properties (Gupta and Abu-Ghannam, 2011; Mohamed et al. 2011).

Macroalgal research has walked miles since the discovery of agar in 1940s (Tseng,

1994). If we look at its journey, the period till 1970s was the era of macroalgal taxonomy,

eco-physiology and biochemistry followed by mutation studies, cultivation and

biotechnology in 1980s (Dring, 1982), bioactives in early 1990s (Renn, 1997). The period of

mid-1990s proved to be the milestone in macroalgal biotechnology with the development of

genetic transformation, tissue culture and the introduction of expressed sequence tags (EST)

approach in 1997 (Lluisma & Ragan, 1997). This eventually culminated in starting of a new

era of molecular phylogenetics and genomic studies (Reddy et al. 2010) which is still

continuing and has led to the development of numerous EST databases (Nikaido et al. 2000;

Weber et al. 2004; Stanley et al. 2005; Teo et al. 2007; Wong et al. 2007; Xiaolei et al.

2007; Aspilla et al. 2010) and whole genome sequencing of Ectocarpus siliculosus (Cock et

al. 2010a, b). However, a shift towards multi-disciplinary research has been made in the last

decade with new approaches for greater understanding of seaweed biology that encompasses

seaweed biodiversity, cultivation, bioactives, nutraceuticals, and developmental biology to

stress physiology, proteomics and genomics in order to realize their potentials to the fullest

extent possible (Hervé et al. 2008; Plaza et al. 2008; Dittami et al. 2009; González et al.

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2010; Gravot et al. 2010; Kumar et al. 2010; Kumari et al. 2010). In the midst of these

recent developments, the area of lipid biochemistry is still in its infancy. Much of today's

knowledge about the lipid biochemistry and metabolism is based on the advances in lipid

research made in 1960s and 70s, i.e., the time when lipid research was one of the most

intensely studied areas of biology. The bioinformatics resources for biochemical pathways

such as Kyoto Encyclopedia of Genes and Genomes (KEGG) have largely relied on the

knowledge obtained during that period (Orešič 2011). Although macroalgal lipids have been

extensively studied for their fatty acid (FA) compositions due to their nutritional

implications and for novel FA oxidation products (oxylipins), most of these studies were

mainly confined to the isolation, structural characterization and their biological properties of

different lipid/oxylipin molecules. Also, the complete lipid repertoire of any macroalgae has

yet not been resolved. Moreover, our knowledge on metabolic pathways of lipid and FA

metabolism and the genes involved, is mainly based on those of higher plants and

microalgae and is believed to be similar to them in one or more aspects. In this chapter, an

attempt was made to review the recent developments in lipid biochemistry and its status in

macroalgae including the biosynthetic pathways of lipids, fatty acids and oxylipins. The

developments in acclimatory roles of lipids and fatty acids in response to changes in

environmental factors have also been dealt. Further, the current status of lipidomics in algae

has been discussed presuming its promising implications in elucidation of novel lipids and

understanding of complex metabolic pathways.

1.2. Lipids and an update on lipid classification system

Lipids are no longer the bystanders in the drama of biological systems, assigned with

the passive role of forming structural components of cell membrane. Today, lipids are

known as diverse and ubiquitous group of compounds that plays various biological functions

besides constituting cellular membranes. Our enriched knowledge has revealed that lipids

also serve as energy reservoirs, provide hydrophobic environment for membrane protein

functions and interactions. They also play prominent roles in the regulation of cellular

bioenergetics, modulates systemic energy balance through eicosanoid and lysolipid

production (Vegiopoulos et al. 2010). Many classes of lipids such as eicosanoids, lysolipids,

diacylglycerols, phosphatidic acids and ceramides serve as secondary messengers in cellular

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signalling pathways (Haimi et al. 2006; Gross and Han, 2011; Han et al. 2012). Lipid

molecules exhibit high structural diversity due to variable chain length, a multitude of

oxidative, reductive, substitutional and ring-forming biochemical transformations,

modification with sugar residues and other functional groups of different biosynthetic origin

(Fahy et al. 2011). There are no reliable estimates of the number of discrete lipid structures

in nature, due to the technical challenges of elucidating chemical structures. It has been

hypothesized that there are approximately 200,000 lipid structures, based on acyl/alkyl chain

and glycan permutations for glycerolipids, glycerophospholipids and sphingolipids

(Yetukuri et al. 2008). Such high level of diversity makes it important to develop a

comprehensive classification, nomenclature, and chemical representation system to

accommodate the myriad lipids that exist in nature (Fahy et al. 2011).

Conventionally, lipids were defined as any group of compounds that are insoluble in

water but are soluble in organic solvents. These chemical features are present in a broad

range of molecules such as fatty acids, phospholipids, sterols, sphingolipids, terpenes and

others. According to Christie (1993), lipids are defined as amphiphilic biological substances

consisting of fatty acids and their derivatives and the substances that were biosynthetically

or functionally related to these compounds. Lipids are classified into two groups: simple

lipids and complex lipids. Simple lipids are those which yield two types of primary products

upon hydrolysis such as acylglycerols which yields fatty acids and glycerol on hydrolysis.

Complex lipids are those which yield three or more products upon hydrolysis such as

glycerophospholipids, which yield fatty acids, glycerol and head group on hydrolysis. Same

convention is followed by a number of online lipid database sources such as ‘The Lipid

Library’ and ‘Cyberlipids’ while the Japanese database ‘LipidBank’ defines an additional

third major group as “derived lipids” (alcohol and fatty acids derived from hydrolysis of

simple lipids). This database includes 26 top-level categories in their classification scheme

covering a wide variety of lipids from animal and plant sources. Considering the

heterogeneous nature of lipids and the ambiguities in lipid classification system, the

International Lipid Classification and Nomenclature Committee on the initiative of the

LIPID MAPS Consortium developed and established a comprehensive classification system

in 2005 which was later updated in 2009 (Fahy, 2005; Fahy et al. 2009, 2011). The LIPID

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MAPS classification system is based on the concept of two fundamental building blocks;

ketoacyl groups and isoprene groups. According to the modern convention, lipids are

defined as “hydrophobic or amphipathic small molecules that may originate entirely or in

part by carbanion based condensations of ketoacyl thioesters and/or by carbocation based

condensations of isoprene units”. Based on this classification system, lipids have been

divided into eight categories: fatty acyls, glycerolipids, glycerophospholipids, sphingolipids,

saccharolipids and polyketides (derived from condensation of ketoacyl subunits); and sterol

lipids and prenol lipids (derived from the condensation of isoprene subunits).

The new classification system further laid the foundation of a comprehensive object-

relational database of lipids known as LIPID MAPS Structure Database (LMSD) (Fahy et al.

2007; Sud et al. 2007). It currently contains 37,127 structures (Table 1.1) which are obtained

from various sources: LIPID MAPS Consortium's core laboratories and partners, lipids

identified by LIPID MAPS experiments, biologically relevant lipids manually curated from

LIPID BANK, LIPIDAT, Lipid Library, Cyberlipids, ChEBI and other public sources, novel

lipids submitted to peer-reviewed journals and computationally generated structures for

appropriate classes. Each lipid is assigned a unique LIPID MAPS identifier (LM _ID) of 12-

or 14-character. The format of the LM_ID contains the classification information, provides a

systematic means of assigning a unique identification to each lipid molecule and allows for

the addition of large numbers of categories, classes and subclasses in the future. The last

four characters of the LM-ID comprise a unique identifier within a particular subclass and

are randomly assigned (Fahy et al. 2005, 2009).

Table 1.1 Lipid categories of the comprehensive classification system and the number of structures in the

LIPID MAPS database.

Category Structures in database Fatty acyls 5791 Glycerolipids 7538 Glycerophospholipids 8005 Sphingolipids 3939 Sterol lipids 2617 Prenol lipids 1200 Saccharolipids 1293 Polyketides 6744 Total 37,127

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However, redefinition of lipids by LIPID MAPS has been extremely criticized by

Christie (The Lipid library), according to whom the new definition is still too broad and

appears to suggest that almost any organic compound not a carbohydrate or a protein is a

lipid. It is of no worth for physical chemists and food scientists. Also, what constitutes

'small' in molecular terms is not clear? Moreover the LIPID MAPS classification system is

biased towards animal lipids and many unique and important plant lipids have been

overlooked. Although plants glycosyldiacylglycerols and sulfoquinovosyldiacylglycerols

have now been included in their updated classification at lower hierarchy (Fahy et al. 2009),

there are many shortcomings such as regarding the classification of sphingolipids, position

of ecisosanoids/docosanoids and others, which need to be overcome in the near future.

LIPID MAPS is a private consortium and the recommendations made by them have not yet

been integrated by the international standards body IUPAC-IUB. Till date, plant researchers

follow the old convention of lipid definition and classification owing to its simplicity and

versatility as it can be applied to lipids of any source of biological origin. Christie (The

Lipid Library) has also stressed that a subdivision of glycerolipids into two broad classes

according to polarity or complexity is so convenient for analysts, biochemists and physical

chemists that it should be given greater weight in any classification system, as those defined

in the first edition of Lipid Analysis (Christie, 1973). Considering the above mentioned

ambiguities, macroalgal lipids shall be discussed according to the old lipid conventions in

this study.

1.3. Macroalgal lipids

Macroalgal lipids consist of phospholipids, glycolipids (glycosylglycerides) and non-

polar glycerolipids (neutral lipids) analogous to higher plants along with betaine and some

unusual lipids that may be characteristic of a particular genus or species. Their chain length

and degree of unsaturation are also significantly higher than those of higher plants. The

basic structure of glycerolipids consists of a glycerol backbone metabolically derived from

glycerol 3-phosphate to which hydrophobic acyl groups are esterified at sn-1 and sn-2

positions. Phospholipids are characterized by the presence of a phosphate group at sn-3

position which is further linked to a hydrophilic head group that classifies individual

phospholipid molecules. The major phospholipids found in algae are phosphatidylglycerol

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(PG), phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylserine (PS),

phosphatidylinositol (PI) and phoshatidic acid (PA) containing glycerol, choline,

ethanolamine, serine, myo-inositol, and phosphomonoester as their characteristic head

groups respectively (Fig. 1.1).

Fig. 1.1 Structure of common lipid molecules found in macroalgae.

Glycolipids contain 1, 2-diacyl-sn-glycerol moiety with a mono- or oligosaccharide groups

attached at sn-3 position of the glycerol backbone. The typical algal glycolipids include

monogalactosyldiacylglycerol (MGDG), digalactosyldiacylglycerol (DGDG) and sulfolipid,

sulfoquinovosyldiacylglycerol (SQDG) with their respective structures as 1,2-di-O-acyl-3-

O-β-D-galactopyranosyl-sn-glycerol, 1,2-di-O-acyl-3-O-(6'-O-α-D-galactopyranosyl-β-D-

galactopyranosyl)-sn-glycerol and 1,2-di-O-acyl-3-O-(6'-deoxy-6'-sulfo-α-D-

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glucopyranosyl)-sn-glycerol respectively. MGDG and DGDG contain one and two galactose

molecules respectively and are uncharged at physiological pH while, SQDG carries a

negative charge due to its sulfonic acid residue at position 6 of the monosaccharide moiety

(Fig. 1.1). In non-polar glycerolipids, either one, two or all the three positions (sn-1, sn-2

and sn-3) are esterified to the hydrophobic acyl groups that may be saturated or unsaturated,

forming monoacylglycerol, diacylglycerol and triacylglycerol respectively. Betaine lipids

contain a betaine moiety instead of phosphorus or carbohydrate as a polar group linked to

sn-3 position of glycerol by an ether bond with fatty acids esterified in sn-1 and sn-2

positions. The betaine lipids present in macroalgae are 1,2-diacylglyceryl-3-O-4'-(N,N,N-

trimethyl)-homoserine (DGTS) and 1,2-diacylglyceryl-3-O-2'-(hydroxymethyl)-(N,N,N-

trimethyl)-β-alanine (DGTA) (Fig. 1.1). These betaine lipids are all zwitterionic at neutral

pH due to their positively-charged trimethylammonium group and a negatively charged

carboxyl group.

1.3.1. Phospholipids

Phospholipids (PL) represent 10-20% of total lipids in macroalgae (Dembitsky and

Rozentsvet, 1990; Dembitsky and Rozentsvet, 1996). They are located in extra-chloroplast

membranes with the exception of PG which occurs in significant amounts in thylakoid

membranes. Cell membranes utilize the amphiphilic nature of phospholipids to maintain its

structural integrity and selective permeability while PG aids glycolipids in maintaining the

stability of photosynthetic apparatus. PG is the dominant phospholipid in Chlorophyta and

accounts for 20 - 47% of PL while PC in red, representing >60% of PL and both PC and PE

in brown algae accounting to 11.3-29.3% of PL (Dembitsky et al. 1990; Dembitsky and

Rozentsvet, 1996; Jones and Harwood, 1992; Khotimchenko et al. 1990; Kulikova and

Khotimchenko, 2000; Illijas et al. 2009; Vaśkovsky et al. 1996). However, PC is often

replaced with DGTS in green and its homologue, DGTA in brown macroalgae. PS and PI

are found in appreciable amounts while DPG and PA present as minor components. In

contrast, Rozentsvet et al. (1995) reported higher PA contents (2.5-17.1% of PL) for 12

species of fresh water algae. A large number of unidentified lipids are also found in amounts

ranging from 2.7-10.3% of PL (Dembitsky and Rozentsvet, 1990; Dembitsky et al. 1990;

Kulikova and Khotimchenko, 2000). Phospholipids are further characterized by higher

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contents of n-6 fatty acids (FAs) as compared to galactolipids except PG that has substantial

amount of n-3 FAs especially, ALA. Major FAs present are oleic, palmitic, stearic acid,

arachidonic acid (C20:4 n-6, AA) and eicosapentaenoic acid (C20:5 n-3, EPA). Further, an

unusual FA, ∆3-trans-hexadecenoic acid (16:1, 3t) is esterified to sn-2 position of PG in all

eukaryotic photosynthetic organisms (Tremolieres and Siegenthaler, 1998).

Moreover, red algae also contain small amounts of sphingolipids such as cerebrosides

and ceramides detected in Chondrus crispus, Polysiphonia lanosa, Ceratodictyon

spongiosum and Halymenia sp. (Bano et al. 1990; Lo et al. 2001; Pettitt et al. 1989).

Vaśkovsky et al. (1996) detected ceramidephosphoinositol (CPI) in 11 red algae.

Subsequently, Khotimchenko et al. (2000) quantified this lipid from 22 red algal species

belonging to Nemaliales, Cryptonemiales, Gigartinales, Rhodymeniales and Ceramiales.

They reported its range from 2.6-15.7% of PL in Nemalion vermiculare and Gracilaria

verrucosa, respectively. Further, Khotimchenko and Vaśkovsky (2004) isolated and

characterized inositol containing sphingolipid from G. verrucosa that contained palmitic

(51.7%), stearic (23.2%), myristic (9.8%), oleic (9.8%), and palmitoleic acids in its acyl

chains.

1.3.2. Glycolipids

Glycolipids are predominantly located in photosynthetic membranes with MGDG

and SQDG strictly restricted to the thylakoid membranes of the chloroplast while DGDG is

also found in extraplastidial membranes. Recently, X-ray crystallographic study of PSI and

PSII revealed the presence of 4 and 25 lipid molecules (MGDG, DGDG, SQDG and PG)

respectively in Thermosynochococcus elongatus (Guskov et al. 2009). These glycolipids are

found to be indispensible for assembly and functional regulation of PSII (Mizusawa and

Wada, 2012). Further, they invariably constitute more than half of the lipids with MGDG

representing 31 - 56% (Hofmann and Eichenberger, 1997; Khotimchenko, 2002; Muller and

Eichenberger, 1994; Sanina et al. 2004; Yan et al. 2011) with the exception of a few red

algae such as Palmaria stenogona, Ceramium kondoi, Laurencia nipponica, Ahnfeltia

tobuchiensis and Exophyllum wentii where DGDG was the characteristic glycolipid (35.7-

64% of polar lipids) (Khotimchenko, 2002; Illijas et al. 2009; Sanina et al. 2004) whereas

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the members of Fucales (brown algae) contained higher SQDG content varying between

36.8 - 48.8% (Khotimchenko, 2002; Sanina et al. 2004).

A unique feature of glycolipids is their high n-3 PUFA contents similar to higher

plants. MGDG is the most unsaturated glycolipid in green and red algae with DGDG in

brown algae while SQDG was the most saturated one. Their FA compositions revealed that

they contain a mixture of prokaryotic and eukaryotic types of FAs (FAs containing one C18

and one C16 PUFAs). Moreover, marine macroalgae also contain long chain C20 and C22

PUFAs such as AA, EPA and docosahexaenoic acid (C22:6, n3, DHA) in contrast to the

fresh water algae with α-linolenic acid (C18:3 n-3, ALA) as a major FA in galactolipids and

palmitic acid in SQDG. The chain length of these glycolipid FAs (C16 or C18) indicates

whether they are synthesized de novo within the plastid or imported from the endoplasmic

reticulum. MGDG and DGDG contain hexadecatetraenoic acid (C16:4 n-3), ALA,

stearidonic acid (C18:4 n-3, STA) and linoleic acid (C18:2 n-6, LA) in green, AA and EPA

in red and all these FAs in brown macroalgae while SQDG contains palmitic and oleic acid

as major FAs (Hofmann and Eichenberger, 1997; Illijas et al. 2009; Khotimchenko, 2002;

Sanina et al. 2004). However, higher contents of AA, EPA and ALA have been reported in

SQDG of Ahnfeltia touchiensis, Ulva fenestrata and Undaria pinnatifida (Sanina et al.

2004).

1.3.3. Betaine lipids

Betaine lipids are widely distributed in algae and extensively reviewed by Dembitsky

(1996) and Kato et al. (1996). DGTS abundantly occurs in Chlorophyta with 5.2 - 56.5% of

polar lipids and DGTA in brown algae with 7.3- 96.8% of polar lipids (Dembitsky and

Rozentsvet, 1989; Jones and Harwood, 1992; Eichenberger et al. 1993; Muller and

Eichenberger, 1994; Dembitsky and Rozentsvet, 1996; Makewicz et al. 1997; Kulikova and

Khotimchenko, 2000). However, there is no report of betaine lipids in most of the red algal

species investigated except the presence of DGTS in Lomentaria articulata, Mastocarpus

stellatus, Phyllophora pseudoceranoides, Membranoptera alata and Phycodrys rubens

(Künzler and Eichenberger, 1997). These two betaine lipids resemble PC due to their

quarternary ammonium group and hence replace PC in most of the marine algae, even to

traces such as in Ulotrichales, Scytosiphonales, Desmarestiales and others. In contrast,

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freshwater algae mainly contain PC and little DGTS. They also vary in their FA

compositions, exhibiting saturated fatty acids (SFAs), myristic and palmitic at sn-1 and C18

PUFAs, predominantly LA and ALA at sn-2 position while DGTS in marine algae are

esterified to long-chain PUFAs at both the sn-1 and sn-2 positions. DGTA contain palmitic,

myristic, oleic, LA, ALA, AA and EPA as major FAs (Hofmann and Eichenberger, 1997;

Makewicz et al. 1997). DGTA is considered to play an important role in the redistribution of

acyl chains and the biosynthesis of galactolipids and DGTS in lipid-linked desaturation of

fatty acids (Hofmann and Eichenberger, 1998).

1.3.4. Non-polar glycerolipids (Neutral lipids)

Triacylglycerol (TAG) is the most prevalent neutral lipid accumulated in macroalgae

as storage product and energy reservoirs. Its level is highly plastic in algae and ranges

between 1% and 59.3% (Dembitsky et al. 1992; Rozentsvet et al. 1995; Dembitsky and

Rozentsvet, 1996; Hofmann and Eichenberger, 1997; Khotimchenko and Kulikova, 1999;

Kulikova and Khotimchenko, 2000; Kamenarska et al. 2004; Illijas et al. 2009). Algal lipids

are mostly characterized by saturated and monounsaturated fatty acids but many oleaginous

microalgae exhibit the potential to accumulate long-chain PUFAs (AA, EPA and DHA).

Parietochloris incisa accumulates AA, Phaeodactylum tricornutum, Porphyridium

cruentum, Nitzschia laevis, Nannochloropsis sp., accumulate EPA, Pavlova lutheri

accumulates both AA and EPA and S. mangrovei, Isochrysis galbana DHA (Bigognoa et al.

2002; Chen et al. 2007; Khozin-Goldberg et al. 2000; Khozin-Goldberg and Boussiba, 2011;

Meireles et al. 2003; Patil et al. 2007).

1.3.5. Unusual lipids

In addition, a large number of unusual lipids have been reported in various algal

species and are mentioned in Table 1.2.

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Table 1.2 List of unusual lipids reported from macroalgae.

Macroalgae Novel lipids References Chondria armata Six minor new glycolipids in crude

methanolic extracts that included 1,2-di-O-acyl-3-O-(acyl-6’-galactosyl)-glycerol and sulfonoglycolipids 2-O-palmitoyl-3-O-(6’sulfoquinovopyranosyl)-glycerol and its ethyl ether derivative

Al-Fadhli et al. (2006)

Ulva fasciata

Mannose and rhamnose containing glycolipids

El-Baroty et al. (2011)

Arainvillea nigricans antimitotic ether-linked glycoglycerolipids nigricanosides A and B

Williams et al. (2007)

Sargassum thunbergii (2S)-1-O-(5Z,8Z,11Z,14Z,17Z-eicosapentaenoyl) -2-O-(9Z,12Z,15Z-octadecatrienoyl)-3-O-β-D-galactopyranosyl-sn-glycerol and (2S)-1-O-(9Z,12Z,15Z-octadecatrienoyl)-2-O-(6Z,9Z,12Z,15Z-octadecatetraenoyl)-3-O-β-D-galactopyranosyl-sn-glycerol

Kim et al. (2007)

Brown algae Phosphatidyl-O-[N-(2-hydroxyethyl) glycine] (PHEG) containing glycine head group (3% - 25% of PL) and rich in AA (80%) and EPA (10%).

Eichenberger et al. (1995), Makewicz et al. (1997), Kullikova and Khotimchenko (2000)

Brown algae Amino acid (-CH2- CH2-NH- CH (NH2) - CH2- CH2-COOH) containing PL

Khotimchenko and Titlyanova (1996)

1.3.6. Lipid biosynthesis in macroalgae

Macroalgal lipid metabolism from de novo fatty acid biosynthesis to the formation of

complex glycerolipids is similar to those of higher plants and microalgae. Lipid biosynthesis

in macroalgae occurs both by prokaryotic and eukaryotic pathway and involves the

cooperation between the plastid and the extraplastidial compartment, with the participation

of enzymes of the endoplasmic reticulum (ER) and chloroplast envelope (Guschina and

Harwood, 2006; Harwood and Guschina, 2009; Li-Beisson et al. 2010). Phospholipid

biosynthesis mainly takes place at ER. PA is the common precursor of phospholipids and is

synthesized by serial reactions catalyzed by acyl-CoA:glycerol-3-phosphate acyltransferases

(GPAT) and acyl-CoA:lysophosphatidic acid acyltransferases (LPAAT) in ER and

exclusively contains C18 FAs in the sn-2 position. PC and PE are formed from

diacylglycerol (DAG) by CDP-choline and CDP-ethanolamine pathways, which may be

obtained from PA hydrolysis by phosphatases (Ohlrogge and Browse, 1995; Li-Beisson et

al. 2010). Other phospholipids such as PI, PS and PG are synthesized by cytidine

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diphosphate diacylglycerol (CDP-DAG) pathway (Ohlrogge and Browse, 1995; Carman,

1997; Li-Beisson et al. 2010). In this pathway PA gets activated by cytidine triphosphate

(CTP) thereby, forming CDP-DAG which reacts with head groups such as myoinositol,

serine and glycerol 3-phosphate resulting in the formation of PI, PS and

phosphatidylglycerol phosphate (PGP), a precursor of PG. PGP phosphatase (PGPP) then

catalyzes dephosphorylation of PGP to produce PG Alternatively, PC is also formed by

methylation of PE and head group exchamge (Ohlrogge and Browse, 1995; Li-Beisson et al.

2010).

The glycolipid biosynthesis exclusively occurs in plastid. The diacylglycerol backbones

for chloroplast lipid (MGDG, DGDG and SQDG) synthesis are derived from both the ER-

localized eukaryotic pathway and the inner envelope-localized prokaryotic pathway

(Ohlrogge and Browse, 1995). A C16-FA on the sn-2 position is a signature for plastidial

origin of a DAG backbone while a C18-FA on the sn-2 position indicates the ER-derived

DAG backbone. MGDG and DGDG are synthesized in the envelope by two different

galactosyltransferase activities, each transferring a galactose moiety from UDP-Gal to the

head group of DAG or MGDG (Kelly and Dörmann, 2004; Andersson and Dormann, 2008).

The anomeric configuration of the resulting galactolipids is always a β-glycosidic linkage to

the first sugar and an α-glycosidic linkage to the second. Subsequently, di, tri- and

tetragalactosyl diacylglycerols are also formed by a processive galactosyl transferase

activity (Li-Beisson et al. 2010). SQDG is assembled in the chloroplast envelope and is

formed by transfer of a sulfoquinovosyl group from UDP-sulfoquinovose onto the head

group of DAG (Benning, 2008). UDP-sulfoquinovose is assembled in the plastid stroma

from sulfite and UDP-glucose, which in turn is synthesized by UDP-glucose

pyrophosphorylase 3 (Okazaki et al. 2009). Further, all the three glycolipids in the plastid

envelope are subjected to further desaturation by envelope or thylakoid-bound desaturases

(Shanklin and Cahoon, 1998; Li-Beisson et al. 2010; Shimojima, 2011). Recently, Sato and

Moriyama (2007) deciphered an alternative pathway of glycolipid biosynthesis in red

microalga Cyanidioschyzon merolae. These authors reported that this alga lacks the acyl

lipid desaturases of cyanobacterial origin as well as the stearoyl acyl-carrier protein

desaturase, which are the major desaturases in higher plants and green algae. Instead, it

synthesizes MGDG via a coupled pathway using plastidic derived 16:0 and ER derived LA.

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Moreover, cyanobacteria have a different pathway of MGDG synthesis wherein a glucose

moiety is first transferred from UDP-glucose onto DAG generating

monoglucosyldiacylgycerol. Then an epimerase activity converts the β-glucosyl polar head

into β-galactosyl, producing MGDG.

Further, TAG is synthesized in the ER from diacylglycerol by ER-specific

acyltransferases, (Kennedy pathway) and is deposited exclusively in lipid droplets in the

cytosol (Ohlrogge and Browse, 1995; Li-Beisson et al. 2010). The pathway involves

sequential acylation of glycerol 3-phosphate and subsequent dephosphorylation. The first

acyl group is added by glycerol 3-phosphate acyl transferase (GPAT), second acyl group by

lysophosphatidyl acyltransferases (LPAAT) resulting in the formation of DAG. Further

DAG is acylated on the sn-3 position using a fatty acyl-CoA molecule by diacylglycerol

acyl transferase (DGAT) to form TAG. In addition, acyl-CoA-independent reactions also

contribute significantly to the production of TAG in some plant and algal species. DAG is

directly incorporated into TAG by the action of phospholipid: diacylglycerol acyltransferase

(PDAT) (Dahlqvist et al. 2000). However, recently, Fan et al. (2011) reported a chloroplast

pathway for the de novo biosynthesis of triacylglycerol in Chlamydomonas reinhardtii,

wherein this alga uses DAG derived almost exclusively from the chloroplast to produce

TAG. This unique TAG biosynthesis pathway is largely dependent on de novo fatty acid

synthesis, and the TAG formed in this pathway is stored in lipid droplets in both the

chloroplast and the cytosol.

A little information is available on macroalgal lipid metabolism especially at the

molecular level while the information on microalgal lipid biochemistry has tremendously

increased in last few years due to their potential of being utilized as energy feedstocks for

biodiesel production. Many genes involved in lipid biosynthesis (especially TAG

biosynthesis) have been identified such as acyl transferases [acyl-CoA:glycerol-3-phosphate

acyltransferase (GPAT), acyl-CoA:diacylglycerol acyltransferases (DGAT),

phospholipid:diacylglycerol acyltransferase (PDAT), acyl-CoA:lysophosphatidic

acyltransferase (LPAAT), Lysophosphatidylcholine acyltransferase (LPCAT)] from

Chlamydomonas reinhardtii, Phaeodactylum tricornutum, Isochrysis galbana, Thalassiosira

pseudonana, Euglena gracilis, Pavlova salina, Thraustochytrium sp., Parietochloris incisa,

CHAPTER 1 

14  

Galdiera sulpharia, Ostreococcus tauri, O. lucimarinus, Micromonas pusilla and

Mantoniella squamata. (reviewed by Khozin-Goldberg and Cohen, 2011; Chen and Smith,

2012). In addition, the enzymes diacylglycerol:CDP-ethanolaminephosphotransferase (EPT)

and CTP: phosphoethanolamine cytidylyltransferase (ECT) involved in

phosphoethanolamine biosynthesis have been cloned and characterized from C. reinhardtii

(Yang et al. 2004a, b). The function of ECT was confirmed by heterologous expression in

Escherichia coli which demonstrated the production of CDP-ethanolamine from

phosphoethanolamine and CTP. The genes responsible for glycolipids; MGDG, DGDG and

SQDG synthesis have also been identified in microalgae (Riekhof et al. 2003; Sato et al.

2003; Riekhof et al. 2005). Its recently only that Chan et al. (2012a) reported on the basis of

EST analysis that lipid biosynthesis in red algae is similar to that in vascular plants, but not

all of the nuclear-encoded genes associated with lipid synthesis in plants are encoded in the

nuclear genome of Pyropia spp. For example red algae lack complete plastidial desaturase

pathway and thus the transfer of ER-derived C20 FAs into plastids becomes essential. The

gene responsible for this lipid trafficking between plastid and ER, TGD

(trigalactosyldiacylglycerol) has 3 orthologs in red alga Pyropia spp., of which TGD 1 and 2

is plastid encoded while TGD 3 is encoded on nuclear genome (Chan et al. 2012b).

1.4. Macroalgal fatty acids

Fatty acids are carboxylic acids with long aliphatic chains that may be straight or

branched, saturated or unsaturated. Most of the naturally occurring FAs contain even carbon

numbers (C4-C26) in macroalgae. On the basis of number of double bonds present, FAs are

classified as monounsaturated FAs (MUFAs, with 1 double bond), and polyunsaturated FAs

(PUFAs, with ≥ 2 double bonds). Further, PUFAs are classified as n-3 or n-6 FAs depending

on the position of the first double bond from the methyl end. n-3 PUFAs are of nutritional

importance as these cannot be synthesized by humans and thus obtained through diet. Often

FAs also contain other groups such as, hydroxyl, halogens, keto, epoxy groups and others

thereby forming hydroxyl-, halogenated-, oxo- and epoxy FAs. Macroalgae are extensively

explored for their fatty acids, especially PUFAs (representing 10-60% of total fatty acids;

TFAs) due to their chemotaxonomic and nutritional importance, with their compositions

varying even within the same phyla. It has been demonstrated that green macroalgae are rich

CHAPTER 1 

15  

in C18 PUFAs (ALA, STA and LA), reds in C20 PUFAs (AA and EPA) while brown

macroalgae exhibit both C18 and C20 PUFAs in appreciable amounts (Khotimchenko et al.

2002; Li et al. 2002; Colombo et al. 2006; Yazici et al. 2007; Chakraborty and Santra, 2008;

Galloway et al. 2012; Pereira et al. 2012). These long chain PUFAs are indispensible for

proper growth and development of organisms with n-3 PUFAs (ALA, STA and EPA) being

beneficial for the prevention of cardiovascular and other chronic diseases such as diabetes,

hypertension and autoimmune diseases, DHA for visual and neurological health while AA

and EPA are precursors of bioregulators prostaglandins, thromboxanes and other

eicosanoids, which influence inflammation processes and immune reactions (Calder and

Grimble, 2002).

The primary FA biosynthesis in macroalgae takes place in plastids analogous to higher

plants and microalgae catalyzed by fatty acid synthase (FAS). The initial substrate malonyl-

CoA is formed in a two step reaction by acetyl CoA carboxylase (ACCase). Two types of

ACCase have been identified in algae: a prokaryotic-type multisubunit enzyme in the plastid

and a multifunctional homomeric enzyme in the cytosol, similar to higher plants (Sato and

Moriyama, 2007). However, recently Huerlimann and Heimann (2012) reported that

heteromeric form of ACCase is found in green and red algae of primary endosymbiosis and

homomeric ACCase in brown algae and heterokonts of secondary endosymbiosis. Malonyl-

CoA enters into a series of condensation reactions with acetyl-CoA, then acyl-ACP

acceptors. (Somerville et al. 2000; Li-Beisson et al. 2010). These reactions are catalyzed by

3-ketoacyl-ACP synthases (KAS) resulting in the formation of a carbon-carbon bond and

decarboxylation of malonyl-ACP. Three KAS isoforms have been identified that are

required to produce an 18-carbon FA. The initial condensation reaction of acetyl-CoA and

malonyl-ACP is catalyzed by KAS isoform III (KASIII), yielding a four-carbon product (3-

ketobutyrl-ACP). Subsequent condensations (up to 16:0-ACP) require a second enzyme,

namely KASI, whereas the final elongation of the 16-carbon palmitoyl-ACP to the 18-

carbon stearoyl-ACP is catalyzed by a third condensing enzyme, KASII (Pidkowich et al.

2007; Li Beisson et al. 2010). In addition to the condensing reaction, the successive addition

of two-carbon units to the growing fatty acyl chain requires the participation of two

reductases and a dehydrase. 3-ketoacyl-ACP is first reduced by a 3-ketoacyl-ACP reductase

CHAPTER 1 

16  

(KAR), which uses NADPH as the electron donor; 3-hydroxyacyl-ACP is then subjected to

dehydration by hydroxyacyl-ACP dehydratase (HAD), and enoyl-ACP thus obtained is

finally reduced by enoyl-ACP reductase (ENR), which uses NADH or NADPH to form a

saturated fatty acid (Mou et al. 2000). Further, 16:0-ACP is released from the FAS

machinery, molecules elongated to 18:0-ACP are efficiently desaturated to 18:1-ACP by

stromal Δ9 stearoyl-ACP desaturases (SAD). Long-chain acyl groups are then hydrolyzed

by acyl-ACP thioesterases that release fatty acids (Somerville et al. 2000; Li Beisson et al.

2010). The initial substrate for PUFA biosynthesis is 18:1 which after incorporation into PC,

is further desaturated to 18:2 (n-6) (LA) by Δ12 destaurases. LA is desaturated to either α-

linolenic acid (18:3, n-3; ALA) or γ-linolenic acid (18:3, n-6; GLA) by Δ15 and Δ6

desaturases respectively (Fig. 1.2).

Fig. 1.2 Fatty acid biosynthetic pathway in algae. Modified from Guschina and Harwood (2006), Harwood and

Guschina (2009) and Chan et al. (2011a). The blue dashed line reaction pathways are not found in red

algae (Chan et al. 2011a). The green dashed line reactions (Spreacher pathway) are found in mammals

and algae of group dinophyceae (Guschina and Harwood, 2006). Note: ALA-α-linolenic acid (C18:3,

n-3), GLA-γ-linolenic acid (C18:3, n-8), STA-stearidonic acid (C18:4, n-3), AA-arachidonic acid

(C20:4, n-6), EPA-eicosapentaenoic acid (C20:5, n-3), DHA-docosahexaenoic acid (C22:5, n-3).

CHAPTER 1 

17  

These PUFAs are then exchanged by other 18:1 acyl residues and may be released into

cytosol as acyl-CoA derivatives, where they could be extended to about C-22 or longer acyl

chains by specific elongase complexes. In addition very long chain PUFAs are also found in

algae such as AA, EPA and DHA and their proposed biosynthetic pathways (reviewed by

Guschina and Harwood, 2006; Harwood and Guschina, 2009; Khozin-Goldberg and Cohen,

2011) have been summarized in Fig. 1.2. In addition to the standard FA biosynthetic

pathway consisting of oxygen dependent desaturation and elongation steps, long chain

PUFAs such as EPA and DHA are also synthesized by polyketide synthases (PKS)

especially in thraustochytrids (Metz et al. 2001). Further, numerous desaturases and

elongases have been cloned and characterized from microalgae extensively reviewed by

Guschina and Harwood (2006), Harwood and Guschina (2009) and Khozin-Goldberg and

Cohen (2011). However, there are only a few such studies in macroalgae. Recently, Chan et

al (2011a) identified the enzymes involved in FA biosynthesis such as acetyl CoA

carboxylase, FAS I/II, desaturases and elongases and studied the FA desaturation patterns in

transcriptomes of Pyropia spp. obtained from available EST databases. These authors

identified all the four genes encoding the subunits of acetyl-CoA carboxylase complex

(accA through accD) on the plastid genome of Pyropia sp., except for the biotin carboxylase

gene (accC) which was located on the nuclear transcriptome data of Pyropia sp. Moreover,

no KAS II gene was identified in Pyropia sp., suggesting that 16:0-ACP (rather than 18:0-

ACP) is the final product of fatty acid synthesis, or this last elongation step can alternatively

be accomplished by KAS I. This indicated that 16:0-ACP is the main fatty acid conjugate

exported from the plastid and/or the elongation rate of C18-fatty acids is high. Besides this

“plant-type” FAS complex, orthologs of the fungal enzymes were also identified in Pyropia

spp. (Chan et al. 2012a). Furthermore, these authors reported that Pyropia spp. lack plastid

desaturation pathway including the soluble acyl-ACP-desaturase FAB2, as earlier reported

in C. merolae (Sato and Moriyama 2007). Therefore, they hypothesized that possibly,

saturated FAs (16:0 and possibly 18:0) are exported from the plastid to the ER for

desaturation in contrast to higher plants, where, oleic acid (18:1) is the major fatty acid that

is synthesized in chloroplasts and exported to the ER (Li- Beisson et al. 2010). With the

increasing availability of ESTs in macroalgae including the species of Gracilaria, Chondrus,

Griffithsia, Eucheuma, Pyropia, Ulva and Sargassum (Llusima and Ragan, 1997; Nikaido et

CHAPTER 1 

18  

al. 2000; Asamizu et al. 2003; Stanley et al. 2005; Jianfeng et al. 2010; Collén et al. 2006;

Teo et al. 2007; Aspilla et al. 2010) and the whole genome sequence of brown macroalga

Ectocarpus siliculosus (Cock et al. 2010a, b), soon the complete FA biosynthetic genes will

be identified in other macroalgae as well.

1.5. Macroalgal oxylipins

Oxylipins are oxygenated derivatives of PUFAs formed enzymatically either by

lipoxygenases (LOX) and α-dioxygenases (α-DOX) or by chemical (auto) oxidation. The

occurrence and distribution of these molecules are widespread within the lineage with

considerable species-specific differences due to the variability of both FAs and enzymatic

transformations. As macroalgae contain both the C18 and C20 PUFAs, they possess both the

plant- and animal-type oxylipins, i.e. octadecanoid as well eicosanoid pathways emanating

from C18 and C20 PUFAs respectively. In higher plants, PUFAs; roughanic acid (C16:3),

LA and ALA are the major substrates of LOX/α-DOX, resulting in the formation of

respective hydroperoxides. These hydroperoxides form the central branch point of the LOX

pathway and are metabolized in six different reaction pathways. The allene oxide synthase

(AOS) pathway leads to the formation of unstable allene oxides which are further

hydrolyzed to α-, γ-ketols and racemic OPDA (Mosblech et al. 2009). Moreover, the allene

oxides of 13-hydroperoxyoctadecatrienoic acids (13-HpOTrEs) are converted to chiral

OPDA or dinor-OPDA by allene oxide cyclase (AOC) to phytohormones jasmonic acid. The

epoxy alcohol synthase (EAS) pathway leads to the formation of epoxy hydroxy FAs,

peroxidase activity of LOX leads to the formation of ketodienes, hydroperoxide lyase (HPL)

to the formation of short chain aldehydes and the corresponding ω-oxo-FAs. Divinyl ether

synthase (DES) pathway leads to the production of divinyl ethers such as colneleic and

colnelenic acid while peroxygenase pathway leads to the formation of epoxy- or dihydroxy

FAs (Bleé, 1998; Mosblech et al. 2009).

Similarly, in macroalgae, C18 PUFAs are metabolized either at C-9 and C-13 via 9-

and 13-LOX respectively while C20 PUFAs are transformed at C-5, C-8, C-9, C-11, C-12

and C-15 via 5-, 8-, 9-, 11-, 12- and 15-LOX, forming their respective hydroperoxides,

reviewed by Guschina and Harwood (2006) and Andreou et al. (2009). Further, these

hydroperoxides are transformed into hydroxy-, oxo-, epoxy- fatty acids, polyunsaturated

CHAPTER 1 

19  

aldehydes (PUAs) by the action of peroxidases, oxygenases, epoxygenases and

hydroperoxide lyases (HPL) respectively (Gerwick et al. 1993; Kuo et al. 1997; Bouarab et

al. 2004; Guschina and Harwood, 2006; Ritter et al. 2008; Andreou et al. 2009). The various

oxylipins found in algae are presented in Table 1.3 and the biosynthetic pathway of common

oxylipins is exemplified in Fig. 1.3 and 1.4. Moreover, some red algae also form

prostaglandins and leukotrienes either non-enzymatically or by the enzymatic action of

allele oxide synthase/cyclase (AOS/AOC) or cycloxygenase (COX) analogous to animals

(Guschina and Harwood, 2006; Andreou et al. 2009). Recently, Kanamoto et al. (2011)

identified COX gene in Gracilaria vermiculophylla and cloned it in Escherichia coli for the

production of PGF2α. Apart from these simple oxylipins, macroalgae also contain various

complex oxylipins such as polycyclic oxylipins, cyclopropyl hydroxyeicosanoids,

egregialactones, ecklonialactones, hybridialactones, bicyclic cymathere ethers,

cymatherelactones and cymatherols, most of which are formed from intra-molecular

rearrangements of hydroperoxides of either ALA (C18:3, n-3) or stearidonic acid (C18:4, n-

3) (Gerwick et al. 1990; Nagle and Gerwick, 1990; Proteau and Gerwick, 1993; Kousaka et

al. 2003; Lion et al. 2006; Weinberger et al. 2011; Choi et al. 2012; Rempt et al. 2012).

Similar to phyto-oxylipins, algal oxylipins also play various important role in

defense and confer innate immunity in response to biotic and abiotic stress such as

pathogenic bacteria, herbivores, wounding and metal toxicity (Bouarab et al. 2004; Lion et

al. 2006; Gaquerel et al. 2007; Ritter et al. 2008; Küpper et al. 2009; Nylund et al. 2011;

Weinberger et al. 2011; Rempt et al. 2012). However, most of the information available

regarding macroalgal oxylipins has come from metabolic studies rather than genomic studies

due to limited number of available macroalgal genome sequences as compared to higher

plants and microalgae. Consequently, only four putative LOX sequences are available in

NCBI database isolated from the gametophyte Pyropia purpurea (Liu and Reith, 1994), P.

haitensis (accession number-JX188386), Gracilaria chilensis (accession number- JF896804)

and Ectocarpus siliculosus (Cock et al. 2010) and one AOC sequence in E. siliculosus (Cock

et al. 2010).

CHAPTER 1 

20  

Table 1.3 Different types of oxylipins reported from algae.

Macroalgae Biosynthetic enzymes Oxylipins References Acrosiphonia coalita Linoleate and linolenate

9- LOX, 16-LOX Hydroxy and hydroperoxy FAs Coalital (C10-oxylipin), Epoxy alcohol

Bernart et al. (1993)

Cladophora columbiana

Linoleate 9- LOX Hydroxy and hydroperoxy FAs Gerwick et al. (1993)

Ulva intestinalis 12-, 8- , 15- LOX 12-, 8- and 15-HETE Kuo et al. (1997)

Ulva conglobata Linoleate and linolenate 9- LOX, arachidonate 11-LOX

9(R)-HPODE, 9(R)-HPOTrE, 11-HPETE, aldehydes (2,4-decadienal)

Akakabe et al. (2002, 2003)

Ulva lactuca n-9 and n-6 LOX 9- and 13-HODE, 9-HOTrE, 12- and 15-HETE, 12-HEPE, 14-HDHE

Kuo et al. (1997)

Chondrus crispus

Arachidonate (5R)-, (8R)-, (9S)- and (15S)- LOX, linoleate (9S)- and (13S)- LOX, (n-7) Bisallylic hydroxylase (BAH)

Hydroperoxy FAs, hydroxy FAs, diols, epoxy FAs, prostaglandins (PGB1, PGB2, PGA2, 15-keto-PGE2and leukotrienes.

Bouarab et al. (2004), Gaquerel et al. (2007)

Constantinea simplex Arachidonate (12S)- LOX

Cyclopropyl hydroxyeicosanoids Nagle and Gerwick (1990)

Gracilariopsis lemaneiformis

Arachidonate (12S)- LOX , Hydroperoxide isomerase

12 (S)-HpETE, hydroxy FAs2, vicinal dihydroxy FAs (12R, 13S-diHETE)1, 3, Eicosanoids2

1Gerwick et al. (1991), 2Jiang and Gerwick (1991), 3Hamberg and Gerwick (1993)

Murrayella periclados Arachidonate (12S)-LOX Eicosanoids Bernart and Gerwick (1994) Gracilaria asiatica, G. verrucosa and G. lichenoides

Arachidonate 8-LOX, AOS/AOC

Prostaglandins (PGE2 , 15 keto-PGE2,

PGA2, LTB4 ), 8-HETE Sajiki and Kakimi (1997), Imbs et al. (2001)

Gracilaria chilensis Arachidonate LOX , peroxidase

(8R)- HETE, 7S,8R-di-HETE Lion et al. (2006)

Lithothaamnion coralloides

Arachidonate and linoleate LOX, BAH

5-, 11-, 12-, 15-HETE, 11-, 13- and 9-HODE, 11-keto-9Z-12Z-octadecadienoic acid

Gerwick et al. (1993)

Rhodymenia pertusa Arachidonate (12R)- and (5S)-LOX

Eicosanoids (5R,6S-diHETE, 5R,6S-diHEPE, 5-HETE, 5-HEPE)

Jiang et al. (2000)

Polyneura latissima (9S)- LOX, DES, peroxidase

Hepoxilin like metabolite, Polyneuric acid, 9(S)-HETE, 9,15-diHETE

Jiang and Gerwick (1997)

Cymathere triplicata LOX [Bicyclic cymathere ethers] 1, [polycyclic oxylipins cymatherelactone and cymatherols]

2

1Proteau and Gerwick (1992, 1993), Choi et al. (2012)

2 Ecklonia stolonifera LOX Ecklonialactones Todd et al. (1994) Egregia menziesii LOX Egregialactones Todd et al. (1993)

Eisenia spp. 13-LOX Carbocyclic eiseniachlorides , eiseniaiodides and bicyclic cymathere ethers

Kousaka et al. (2003)

Laminaria angustata Arachidonate 12S- and 15S- LOX , linoleate 13-LOX, HPL

13-HPODE, 13-HPOTrE, 12S-, 15-, 11-, 9-, 8-HpETE, C-9 aldehydes from C20 PUFAs and C-6 from C18/C20-PUFAs

Boonprab et al. (2003, 2004)

Laminaria digitata LOX, epoxygenase2

Hydroxy- , hydroperoxy FAs derived from LA, ALA, AA, prostaglandins (PGE1, PGD1, 15 keto-PGF2),12,13- epoxy- octadecaenoic acid, 18-hydroxy-17-oxo-eicosatetraenoic acid

Küpper et al. (2006), Ritter et al. (2008,) Küpper et al. (2009)

CHAPTER 1 

21  

Fig. 1.3 Octadecanoid pathway in macroalgae. Modified from Andreou et al. (2009).

Fig. 1.4 Eicosanoid pathway in macroalgae. Dashed line in red shows putative reactions. (LOX-Lipoxygenase, AOS-Allene oxide synthase, AOC-Allene oxide cyclase, COX-Cyclooxygenase, DES-Divinyl ether synthase). Modified from Andreou et al. (2009).

CHAPTER 1 

22  

1.6. Jasmonates in higher plants and its current status in macroalgae

Jasmonic acid and its derivatives, collectively referred to as jasmonates, comprise a

group of oxylipin signaling molecules that share a high a degree of structural and functional

similarity to prostaglandins found in animals. They are derived from the AOS/AOC branch

of octadecanoid pathway. In higher plants they are involved in regulation of various abiotic

and biotic stress responses including, wounding, UV radiation, ozone treatment, desiccation,

salinity, herbivore attack and infection by microbial pathogens (Wasternack, 2007; Browse

and Howe, 2008; Browse, 2009; Wasternack et al. 2012). In addition in healthy plants, they

mediate developmental processes such as root growth, seed germination, tendril coiling,

trichome initiation, flower development and senescence (Mandaokar et al. 2006;

Wasternack, 2007; Browse, 2009). Jasmonates exert their function by large scale

reprogramming of gene expression, which in part is mediated by the transcription factor

MYC2 (Lorenzo et al. 2004; Kombrink, 2012). Methyl jasmonate (MeJA) is one of the most

active forms of jasmonic acid in plants that is formed by methylation of C1 of jasmonic acid

by jasmonic acid-specific methyl transferase (JMT) (Seo et al. 2001). It was identified as the

odor of Jasminum grandiflorum in 1962 (Demole et al. 1962). First JA-specific

physiological responses were observed by recording root growth inhibition and promotion of

senescence in the early 80s (Ueda and Kato, 1980). After elucidation of the pathway of

jasmonate biosynthesis by Vick and Hamberg in the middle of 80s, first JA-induced

alterations of protein pattern was observed for barley, while the altered gene expression was

found for tomato and both have become the origin for functional analysis of mode of action

of jasmonates. In the early 90s a link between environmental cues, endogenous rise in

jasmonates and altered gene expression was observed. Subsequently, mutants affected in

jasmonate biosynthesis and signaling were isolated. In the last 10 years an exponential

increase of data appeared on jasmonates covering biosynthesis, metabolism, signal

transduction, mutant analyses, gene expression and cross-talk to other phytohormones.

(Wasternack, 2007; Bnac et al. 2009; Kombrink, 2012). Most of our understanding of

jasmonate metabolism has been derived from the mutational studies accomplished with

mutants defective in JA biosynthesis or the physiological response of JA/MeJA treatment

(Wasternack, 2006; Browse, 2009; Kombrink, 2012). In addition to such biological

approaches, chemical research has also been an integral part of jasmonate research.

CHAPTER 1 

23  

Extensive studies on structure-activity relationships have been carried out, including the

synthesis of numerous JA-derivatives and determining their impact on plant responses

(Wasternack, 2007; Wasternack and Kombrink, 2010; Kombrink, 2012). Such approach

provided the first insight into the structural requirement for jasmonates bioactivity and later

culminated in the synthesis of highly active jasmonate analog coronalon (Schüler et al.

2004; Kombrink, 2012). Most recently the jasmonate receptor was identified, and its mode

of action including the proteasomal degradation of repressors of JA-induced gene expression

was a breakthrough in our understanding JA-dependent processes by identification of JAZ

(Jasmonic acid ZIM domain)-proteins (Thines et al. 2007; Wasternack, 2007; Staswick,

2008; Kombrink, 2012). JAZ proteins together with the adaptor protein NINJA (novel

interactor of JAZ) and co-repressor TOPLESS form a transcriptional repressor complex. The

current model of JA perception and signaling implies the SCFCOI1 complex that operates as

E3 ubiquitin ligase and upon binding of JA-Ile targets JAZ proteins for degradation by the

26S proteasome pathway, thereby allowing MYC2 and other transcription factors to activate

gene expression (Pauwels et al. 2010; Wasternack and Kombrink 2010; Kombrink, 2012).

Now, JA biosynthetic pathway (Fig. 1.5) is well established in plants and most of the

participating enzymes have been characterized by biochemical, molecular genetics and

structural approaches (Wastenack, 2007; Wasternack and Kombrink, 2010). JA biosynthesis

initiates by the release of α-linolenic acid (ALA) or hexadecatrienoic acid (HTrA) from

plastid membranes by the action of lipases such as DEFECTIVE IN ANTHER

DEHISCENCE1 (DAD1) and DONGLE (DGL) (Hyun et al. 2008). The released ALA is

oxidized by 13-lipoxygenase (13-LOX) to 13(S)-hydroperoxyoctadecatrienoic acid [13(S)-

HPOT]. The conversion of 13(S)-HPOT into to 12,13(S)-epoxy-octadecatrienoic acid

[12,13(S)-EOT] by AOS is the first committed step of JA biosynthesis. 12,13(S)-EOT is

converted by AOC to optically pure 9(S),13(S)-12-oxo-phytodienoic acid (OPDA), which is

the end product of the plastid localized part of the JA biosynthesis (Stenzel et al. 2003;

Wasternack, 2007; Kombrink, 2012). Similarly, HTrA is converted to dinor-OPDA by the

same set of enzymes (Weber et al 1997). Further, OPDA is transloacted to peroxisome,

mediated by the ABC transporter COMATOSE (CTS1) or by ion trapping mechanism

(Theodoulou et al. 2005). OPDA reductase 3 (OPR3) catalyzes the reduction of OPDA to 3-

oxo-2-(20[Z]-pentenyl)-cyclopentan-1-octanoic acid (OPC-8) (OPC-8). Shortening of the

CHAPTER 1 

24  

octanoic acid side chain of OPC-8 occurs by three subsequent steps of β-oxidation, which is

initiated by the activation of the carboxylic acid moiety to the corresponding CoA ester by

OPC-8:CoA ligase 1 (OPCL1) (Wasternack and Kombrink, 2010; Kombrink, 2012).The

endproduct of β-oxidation, jasmonoyl CoA, is cleaved by an unknown thioesterase (TE) to

(+)-7-isoJA that equilibrates to more stable (-)- JA.

Fig. 1.5 Jasmonic acid biosynthetic pathway in plants. (Note: DAD1-DEFECTIVE IN ANTHER

DEHISCENCE 1, DGL-DONGLE, LOX-Lipoxygenase, AOS-Allene oxide synthase, AOC-Allene

oxide cyclase, OPR 3-OPDA reductase 3, OPCL1- OPC-8 CoA: ligase 1, ACX- acyl-CoA oxidase,

MFP- Multifunctional protein, KAT- L-3-ketoacyl-CoA thiolase, TE-Thioesterase).

CHAPTER 1 

25  

However, jasmonate study in macroalgae is still in its infancy. Till date, JA/MeJA

has only been identified in the red macroalga Gelidium latifolium (Krupina and Dathe,

1991). Bouarab et al. (2004) detected JA in the cell free extracts of Chondrus crispus after

the addition of linolenic acid but their attempts to identify JA in C. crispus cell homogenates

remained unsuccessful. Furthermore, Wiesemeier et al. (2008) were also unable to detect

JA/MeJA and even their biosynthetic precursor 12-oxophytodienoic acid (12-OPDA) in

seven brown algal species of Dictyota, Colpomenia, Ectocarpus, Fucus, Himanthalia,

Saccharina and Sargassum. Moreover, treatment with ecologically relevant concentrations

of JA and MeJA did not lead to a significant change in the profile of medium- and non-polar

metabolites of the tested algae. Only after the application of higher concentrations of ≥ 500

μg ml-1 medium of the phytohormones a metabolic response of unspecific stress was

observed (Wiesemeier et al. 2008). In contrast, Ritter et al. (2008) detected 12-OPDA in

response to copper stress in Laminaria digitata, indicating that Laminaria sp. do employ the

plant-like octadecanoid metabolites to regulate protective mechanisms at least towards

copper stress. Even the entire set of enzymes necessary for the biosynthesis of JA from

linolenic acid have also been identified in the marine red algae Gracilariopsis sp. (Hamberg

and Gerwick, 1993) and Lithothamnion corallioides (Hamberg, 1992). But their presence in

other red, green or brown macroalgae is still an enigma. However, researchers in the last

decade have succeeded a step in establishing their roles in few algae Chondrus crispus

(Bouarab et al. 2004; Collén et al. 2006; Gaquerel et al. 2007), Fucus vesiculosus (Arnold et

al. 2011) and Laminaria digitata (Küpper et al. 2009) in oxidative stress and defense against

endophytes and pathogenic attack.

1.7. Environmental variations

Algae in their natural habitats experience severe environmental stresses including

salinity variations, intense radiation, temperature, desiccation, and chemical pollution that

limit their distribution, production and fecundity (Aguilera and Rautenberger, 2011). Such

fluctuating and dynamic environmental conditions are often associated with cellular increase

in the formation of reactive oxygen species (ROS) as a consequence of photosynthetic

inhibition with excess energy resulting in the production of singlet oxygen (Dring, 2006)

causing “oxidative” stress. Macroalgae acclimate to such abiotic stresses by altering the

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fluidity of cell membranes. The most commonly observed change in membrane lipids

following adverse environmental conditions in algae is the alteration in fatty acid

unsaturation (Guschina and Harwood, 2006).

Some seasonal patterns in lipid and FA contents are reported in macroalgae due to

fluctuations in environmental factors and are summarized here. High lipid content in winter

and autumn as compared to summer season has been observed in Undaria pinnatifida,

Laminaria japonica, Fucus serratus, Egregia menziensii, Condrocanthus canaliculatus, and

Ulva lobata (Kim et al. 1996; Nelson et al. 2002; Gerasimenko et al. 2011). However, high

TAG contents are observed in summer while polar lipids (PL and GL) depended on the algal

development stages throughout the year (Gerasimenko et al. 2011; Kim et al. 1996).

Gerasimenko et al. (2010) reported higher TAG contents in May at the time of sporulation in

brown alga Costaria costata and different classes of GL were in the following order of

MGDG>SQDG>DGDG in April (growth period) and May (sporogenesis period) and

MGDG>DGDG>SQDG in July (beginning of senescence). These lipid changes are often

accompanied by high PUFAs, high unsaturation index (UI) and n-3 > n-6 PUFAs in winter

over summer season as observed in A. touchiensis, L. japonica, U. fenestrata, S. pallidum,

U. pinnatifida, C. taxifolia (Kim et al. 1996; Nelson et al. 2002; Iveša et al. 2004; Sanina et

al. 2008; Gerasimenko et al. 2011). The higher percentage of PUFAs in winter aids in low-

temperature acclimatization and protect photosynthetic machinery from low temperature

photoinhibition (Blankenship, 2002). The substitution of n-6 by n-3 PUFAs, with the change

in season from summer to winter, was also accompanied by the partial substitution of C20

by C18 PUFAs in GLs and PG in contrast to PC and PE in A. touchiensis, L. japonica, U.

fenestrata, S. pallidum, U. pinnatifida, C. costata and F. serratus (Kim et al. 1996; Sanina et

al. 2008; Gerasimenko et al. 2010; Gerasimenko et al. 2011). The sampling season also

affects the concentration of CPI in red algae but the effect is species-specific with

Tichocarpus crinitus and Rhodoglossum japonicum exhibiting higher CPI levels in summer

(12.9% and 13.9% of PL) than fall (8.9% and 6.5% respectively) while Rhodomela larix

reported 1% of PL as CPI in summer that increased to 3.7% in fall (Khotimchenko et al.

2000).

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Nutrient limitation which generally causes a reduced cell division rate in algae,

surprisingly activates the biosynthesis of storage lipids primarily TAGs (Floreto et al. 1996;

Cakmak et al. 2012; Feng et al. 2012; Ördög et al. 2012). Nitrogen-deprivation leads to the

accumulation of neutral lipids (triacylglycerol; TAG) while increase in N leads to increased

biosynthesis of glycolipids especially, monogalactosyldiacylglycerol (MGDG) (Regnault et

al. 1995; Guschina and Harwood, 2006). Similarly, under phosphorus limitation, the content

of phospholipids decreases with a concomitant increase in non-phosphorus glycolipids

(DGDG and SQDG) (Khozin-Goldberg and Cohen, 2006; Bellinger and Van Mooy, 2012).

Also, an increase in SFA accompanied by decrease in UFAs under N-starvation while an

increase in PUFAs at the expense of SFAs and MUFAs under P-starvation has been

demonstrated in Ulva pertusa (Floreto et al. 1996). Although only a few number of species

have been assessed for lipid alterations undergoing in macroalgae subjected to nutritional

stress. This strategy is now-a-days employed to obtain higher biodiesel yields from various

microalgae such as Chlamydomonas, Chlorella, Dunaliella, Scendesmus, Parietochloris,

Isochrysis and Botryococcus (Wang et al. 2009; Li et al. 2010; Chen et al. 2011; Feng et al.

2011; Ördög et al., 2011; Yeesang and Benjamas, 2011; Cakmak et al. 2012; Krasikov et al.

2012).

Salinity is an important environmental factor that affects growth and productivity of

algae by altering membrane permeability and fluidity. The restructuring of membrane lipid

composition is one of the adaptations to survive in high concentration of salt, which is

mainly achieved by increasing the unsaturation of its phospholipid FAs as PUFAs play an

important role in protecting the photosynthetic machinery (Lu et al. 2009). Recently, an

enhanced relative proportion of oleic, LA and ALA by 1.3-1.6 fold with a parallel decrease

in palmitoleic acid at hypersalinities have been observed in Gracilaria corticata while

shifting the cultures from 30‰ to 45‰ (Kumar et al., 2010a). Similarly, Dittami et al.

(2012) also reported an increase in n-3 PUFAs and upregulation of ∆12 and ∆15 desaturases

in Ectocarpus sp. that helped the alga to adapt to low salinities.

Light has been reported to produce profound effects on algal lipid metabolism. The

qualitative changes in lipids as a result of various light conditions have been shown to be

associated with alterations in chloroplast development (Harwood 1998). The effect of light

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on FA composition in macroalgae has been examined only for a few algal species and these

studies yielded contradictory results. In Gracilaria sp., the content of the main PUFA, EPA

increased with increasing photon flux density (Levy et al., 1992), whereas in Gracilaria

verrucosa, the proportion of main PUFA, AA decreased under high light (Floreto and

Teshmina, 1998; Levy et al., 1992). In Gracilaria tikvahiae and Grateloupia sparsa, FA

composition was not affected by light (Dawes et al., 1993). In the green alga Ulva

fenestrata, grown under different solar irradiances in field experiments, MGDG, SQDG and

PG increased 2-3.5 fold when grown at 24% of photosynthetically active radiations (PAR)

compared with algae cultured at 80% of PAR (Khotimchenko and Yakovleva, 2004).

Exposure of algae to low light resulted in increase in the content of EPA in MGDG while

decrease in PG in Tichocarpus crinitus. Light conditions influenced the total lipid content,

wherein algae exposed to 8-10% PAR accumulated lipid 4.2 mg g-1 FW and at 70–80%

PAR, 3.4 mg g-1 FW lipid (Khotimchenko and Yakovleva, 2005). However, Gracilaria

tenuistipitata cultures exposed to low (100 μmol. photons. m-2. s-1) and high light intensity

(1000 μmol. photons. m-2. s-1) for five days showed no statistically significant differences in

the FA contents (Pinto et al., 2011).

Macroalgae adapt to temperature fluctuations by adjusting the lipid composition of their

cell membranes by a process referred to as ‘‘homeoviscous adaptation’’ (Guschina and

Harwood, 2006). The detrimental effects of low temperature on the rigidification of the

membrane lipid bilayers such as loss of ion permeability have been clearly demonstrated in

many psychrophilic and psychrotrophic organisms (Morgan-Kiss et al., 2006). These

organisms utilize a combination of changes in FA composition to regulate the fluidity of the

membrane at low temperatures, including the incorporation of polyunsaturated, short-chain,

branched, or cyclic FAs. Macroalgae inhabiting at low temperature waters are usually richer

in PUFA, with a higher n-3/n-6 fatty acid ratio, and possess a higher degree of total

unsaturation such as those observed in Gracilaria sp. acclimated to low temperature

(Gressler et al., 2010) and Palmaria decipiens endemic to Antarctic region (Becker et al.

2010).

In the recent years, the release of toxic pollutants by growing number of diverse

anthropogenic sources (industrial effluents and wastes, urban runoff, sewage treatment

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29  

plants, agricultural fungicide runoff, domestic garbage dumps, and mining operations) have

become a major threat for aquatic ecosystem negatively affecting the benthic flora and fauna

assemblages (Pinto et al., 2011). These pollutants alter the unsaturation degree of membrane

lipids in algae to combat the oxidative stress conditions. For example, a significant

accumulation of di- and tri-unsaturated FAs, LA and ALA at the expense of palmitic,

palmitoleic and oleic acid has been demonstrated during cadmium stress in Ulva lactuca

(Kumar et al., 2010b). On contrary, a considerable reduction in LA, γ-linolenic acid (18:3 n-

6, GLA), octadecapentaenoic acid (18:5 n-4), dihomogammalinolenic acid (20:3 n-6,

DGLA), AA, EPA and DHA have been observed in red algae G. tenuistipitata and G. dura

exposed to higher concentrations of Cd and Cu (Pinto et al., 2011; Kumar et al., 2012).

Similarly, the exposure to imidazolium ionic liquids in U. lactuca resulted in significant

decrease in n-3 and n-6 PUFAs with a concomitant increase in SFAs (Kumar et al., 2011b).

1.8. Advances in lipid analysis methods and lipidomics

The detection, identification and precise quantification of lipid compounds are

prerequisite for their potential utilization and exploration. Historically, the progress in lipid

research has been hindered mainly due to the analytical constraints in the identification of

lipid structures and quantification of individual molecular species. Multiple approaches have

been employed for the separation, quantification and characterization of lipids including thin

layer chromatography (TLC), gas chromatography (GC), gas chromatography mass

spectrometry (GC-MS), nuclear magnetic resonance (NMR) and high pressure liquid

chromatography (HPLC) in conjunction with a variety of complementary procedures such as

chemical hydrolysis, regiospecific enzymatic cleavage and spetrophotometric assays to

unravel the lipid diversity in biological systems and to quantitate their abundance (Milne et

al. 2006; Gross and Han, 2011). However, most of these strategies required multiple

sequential steps each of which possesses limited sensitivity and accuracy that collectively

resulted in the propagation of errors. Although the utility of GC-MS has provided a robust

platform for the analysis of volatile lipids, the overwhelming majority of cellular membrane

constituents are nonvolatile charged moieties that are not accessible to GC-MS. Thus, the

advances in lipid research has been largely dependent on the development of mass

spectrometric approach and gained impetus with the introduction of soft-ionization

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30  

techniques such as electrospray mass spectrometry and matrix assisted laser desorption

ionization by Fenn (1989) and Karas and Hillenkamp (1988). ESI-MS has greatly simplified

the procedure of lipid analysis and provides better reproducibility and lower detection limits.

In ESI-MS, lipid samples are injected through a capillary tube to which an electric field is

applied. The field generates additional charges to the liquid at the end of the capillary and

produces a fine spray of highly charged droplets that are electrostatically attracted to the

mass spectrometer inlet. The evaporation of the solvent from the surface of a droplet as it

travels through the desolvation chamber substantially increases its charge density. When this

exceeds the Rayleigh stability limit, ions are ejected and ready for MS analysis. The

ionization efficiency of lipids depends on the charge density and the magnitude of dipole

present in the lipid molecule. The advent of ESI-MS analysis heralded the beginning of a

new era of lipidomics in lipid research that enabled researchers to understand the pleiotropic

roles of lipids in biological systems (Welti and Wang, 2004; Forrester et al. 2004; Han and

Gross, 2005; Gross and Han, 2011; Harkewiz and Dennis, 2011; Jung et al. 2011; Murphy

and Gaskell, 2011). ‘Lipidomics’ is a branch of omics science that aims at quantifying a full

complement of lipid molecules in cells, tissues or organisms (Schuhmann et al. 2011). There

are two basic approaches for ESI-MS based lipidomics analysis, each with context

dependent strengths and limitations (Gross and Han, 2011). In traditional approach, lipids

are separated by HPLC and directly sprayed into the ESI ion source for MS analysis by

molecular ion monitoring in conjunction with product ion analysis, selected reaction

monitoring (SRM), or other fragmentation strategies. In the second approach, which is

popularly known as shotgun lipidomics, lipid extracts are directly infused into the MS for

analysis. The direct infusion facilitates the utilization of a wide variety of informative

fragmentation strategies that are not limited by transient elution of individual lipid molecular

species during column chromatography (Jung et al. 2011; Gross and Han, 2011). High

throughput lipidomics generates an enormous amount of data that need to be translated into

knowledge and understanding of biological phenomena (Orešič et al. 2011; Herzog et al.

2011). At present a large number of dedicated online databases (LIPID MAPS, METLIN,

The Human Metabolome Database, LipidBank, MassBank, LIPIDAG, LIPIDAT,

SphingoMap, KEGG, Lipid library, CyberLipids, LMSAD) and softwares (AMDMS-SL,

HMDS, LipidInspector, LipidQA, LipidSearch, LipidView, LipidXplorer, Profiler-Merger-

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31  

Viewer, Lipid MS prediction tool and others) are available that facilitate the lipidomics data

processing including data merging, quality control steps and statistical analysis.

The lipidomics analysis in conjunction with sophisticated software analysis has

identified more than 400 lipid molecular species in RAW 267.2 cells (Dennis et al. 2010),

250 lipids from yeast (Ejsing et al. 2009), 167 galactolipid molecular species in Arabidopsis

thaliana (Ibrahim et al. 2011) and 223 phospholipid molecules in Candida albicans (Singh

et al. 2012). Recently, Vu et al. (2012) identified about 86 oxylipin containing membrane

lipids from Arabidopsis thaliana using shotgun approach. However, this omics approach

remains largely unexplored in algae with a few exceptions for microalgae that utilized the

potential of LC-Q-TOF-MS and ESI-MS for the elucidation of different lipid molecules.

Leblond and co-workers extensively studied the glycolipid profiles of dinoflagellate

Pyrocystis spp., glaucocystophytes Cyanophora paradoxa and Glaucocystis

nostochinearum, raphidophytes Chattonella, Fibrocapsa and Heterosigma spp. and

chlororachinophytes Bigelowiella natans, Gymnochlora stellata and Lotharella spp., with

the latter chlororachinophytic algae exhibiting a novel lauric acid containing MGDG

(C20:5/C12:0, sn-1/sn-2) (Leblond and Roche, 2009; Leblond et al., 2010a, b; Roche and

Leblond, 2011). Recently, He et al. (2011) characterized the polar lipid profile of

Nannochloropsis occulata and identified 200 unique lipid species by online nanoscale high-

performance liquid chromatography followed by electrospray ionization and mass analysis

with a linear ion trap (LTQ) coupled with 14.5 T Fourier transform ion cyclotron resonance

mass spectrometry (FT-ICR MS). Among macroalgae, there are a few reports where

researchers have used ESI-MS for the analysis of desired lipid molecules, where lipids were

pre-separated on HPLC/LC and then analyzed by MS. Kim et al. (2007) used the ESI-MS to

elucidate two new monogalactosyldiacylglycerols from Sargassum thunbergii,

Khotimchenko and Vaśkovsky (2004) identified an inositol-containing sphingolipid from

Gracilaria verrucosa and Al-Fadhli et al. (2006) studied glycolipids in Chondria armata.

However, the shotgun approach has still not been employed to characterize the macroalgal

lipidome.

Macroalgae are untapped sources of various pharmacologically and nutritionally

important lipid molecules. Although they contain low lipid contents (~5% on dry weight

basis), recent studies have shown higher lipid contents of 8-12% DW in various marine

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32  

macroalgal species such as Dictyota, Spatoglossum, Sargassum, Derbesia, Ulva and

Caulerpa (Yaich et al. 2011; Gosch et al. 2012) pointing at a need to re-look at the

macroalgal lipids. The nutritional constraint has been suggested as one of the viable

approach to increase lipid content in biofuels. Thus, it is speculative to study whether similar

lipid accumulation is also profound in macroalgal species. Moreover, macroalgal species are

also rich in nutritionally important PUFAs, and such higher lipid values may facilitate their

use in bio-oil production and PUFA rich food supplements and nutraceuticals. In addition,

the fatty acid oxidation products (oxylipins and jasmonates) have also gained interest in

macroalgal physiology due to their roles in wounding and chemical defense. Nevertheless,

these studies are limited to the few macroalgal species and only a few reports are present for

tropical species that need to be investigated. Therefore, the present study was proposed to

undertake a comprehensive investigation of lipids, fatty acids and their derivatives with the

following objectives:

Optimization of lipid and fatty acid extraction methods in macroalgae.

Fatty acid profiling of different macroalgae from their nutritional and chemotaxonomic

perspectives.

Polar lipid profiling of different macroalgae using shotgun ESI-MS approach.

Quantitative profiling of fatty acid derivatives (hydroxy-oxylipins) in macroalgae.

Study of nutritional constraint imposed by nitrogen and phosphorus in selected

macroalga (Ulva lactuca) with an emphasis on the role of lipids, fatty acids and

oxylipins.

Study of the effect of methyl jasmonate on lipidomics, fatty acid and oxylipin profiling

and oxidative stress in selected macroalga (Gracilaria dura).