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Research review paper

Omega-3 biotechnology: Thraustochytrids as a novel source of omega-3 oils

Adarsha Gupta, Colin J. Barrow, Munish Puri ⁎Industrial Biotechnology Laboratory, Centre for Biotechnology and Interdisciplinary Sciences (BioDeakin), Geelong Technology Precinct, Waurn Ponds, Deakin University,Victoria 3216, Australia

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

Available online 3 March 2012

Keywords:ThraustochytridsMicrobial productionω-3Docosahexaenoic acidEicosapentaenoic acidOleaginousCarotenoidsBiofuel

Thraustochytrids are large-celled marine heterokonts and classified as oleaginous microorganisms due totheir production of docosahexaenoic (DHA) and eicosapentaenoic (EPA) ω-3-fatty acids. The applicationsof microbial DHA and EPA for human health are rapidly expanding, and a large number of clinical trialshave been carried out to verify their efficacy. The development of refined isolation and identification tech-niques is important for the cultivation of thraustochytrids. With a high proportion of lipid biomass, thrausto-chytrids are also amenable to various production strategies which increase omega-3 oil output. Modificationsto the existing lipid extraction methods and utilisation of sophisticated analytical instruments have increasedextraction yields of DHA and EPA. Other metabolites such as enzymes, carotenoids and extracellular polysaccha-rides can also be obtained from these marine protists. Approaches such as the exploration for more diverse iso-lates having fast growth rates, metabolic engineering including gene cloning, and growing thraustochytrids onalternate low cost carbon source, will further enhance the biotechnological potential of thraustochytrids.

© 2012 Elsevier Inc. All rights reserved.

Contents

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17341.1. Omega-3 fatty acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1734

1.1.1. Omega-3 fatty acids as nutraceuticals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17341.1.2. Biosynthesis of omega-3 fatty acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1735

2. Microorganisms producing fatty acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17362.1. Other PUFAs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17362.2. Omega-3 fatty acids: DHA and EPA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1736

3. Thraustochytrids: occurrence and methods for isolation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17363.1. Pollen baiting and direct plating . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17363.2. Antibiotics and antifungal supplemented medium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17373.3. Identification techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1737

3.3.1. Biochemical identification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17383.3.2. Molecular identification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1738

3.4. Ultra structure of thraustochytrids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17384. Classification of thraustochytrids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17385. Production of omega-3 fatty acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1738

5.1. Cultivation medium and bioprocess engineering . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17385.2. Factors affecting omega-3 fatty acid production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1740

6. Lipid extraction methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17407. Quantification of fatty acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17418. Other metabolites produced by thraustochytrids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1741

8.1. Carotenoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17418.2. Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17418.3. Extracellular polysaccharides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1741

Biotechnology Advances 30 (2012) 1733–1745

⁎ Corresponding author at: BioDeakin, Deakin University, Australia. Fax: +61 3 5227 2170.E-mail address: munish.puri@deakin.edu.au (M. Puri).

0734-9750/$ – see front matter © 2012 Elsevier Inc. All rights reserved.doi:10.1016/j.biotechadv.2012.02.014

Contents lists available at SciVerse ScienceDirect

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j ourna l homepage: www.e lsev ie r .com/ locate /b iotechadv

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9. Thraustochytrids and their potential . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 174210. Limitations of marine exploration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 174211. Conclusions and future prospects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1742

. Note added in Proof . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1742Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1743References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1743

1. Introduction

The potential for identifying and extracting novel bio-actives valu-able to themedical and food industriesmakesmarine environments at-tractive to researchers. Exploring these environments might appearsimple but it is a challenging task given the magnitude of marine life,complicating the investigation of oceans and other aquatic ecosystems(Karl, 2007; Lee et al., 2010). Further adding to the difficulties is thecontinuing anthropogenic contamination of marine environmentswith heavy metals and other contaminants. Pollution of marine envi-ronments is directly responsible for their deterioration, resulting in de-creased phytoplankton growth and reduced synthesis of omega-3 fattyacids (Kang, 2011). A variety of fish species such as herring, mackerel,sardine and salmon are regarded as good sources of omega-3 fattyacids (Gunstone, 1996). Due to the many shortcomings of fish-derivedoil including undesirable taste and odour, diminishing supplies, objec-tions by vegetarians, its chemical processing methods, and the presenceof contaminants such asmercury, dioxins and polychlorinated biphenyls(Certik and Shimizu, 1999; Hooper et al., 2006), research has beendiverted towards the exploitation of other marine species for the devel-opment of a suitable alternatives. It is hoped that novel oleaginous or-ganisms (microorganisms which produce more than 25% lipid on a drycell weight basis) can be tailored to produce high amounts of omega-3fatty acids, particularly docosahexaenoic acid (DHA) and eicosapentae-noic acid (EPA) (Barrow, 2010; Dyal and Narine, 2005).

The microbial production of omega-3 fatty acids with special refer-ence to thraustochytrids, has been gaining interest in recent years(Raghukumar, 2008; Sijtsma and Swaaf, 2004). Thraustochytrids belongto a marine group of eukaryotic protists called Labyrinthulomycetes;single celled organisms abundantly distributed in marine ecosystems.These mono-centric protists are receiving attention on the basis oftheir ultrastructure andbiochemical characterisation. Based on phyloge-netic analysis, thraustochytrids are related to the heterokont groups ofmicroorganisms and also some other phyla of marine microbes such asChromophytes and Prymnesiophytes, although they are considered pri-marily as a group of their own (Chamberlain andMoss, 1988). Thrausto-chytrid cells are of uniform spherical shape, ranging between 30 and100 μm in diameter (Bremer, 2000). In contrast to low lipid accumulat-ing bacteria, they can produce oils with a high percentage of polyunsat-urated fatty acids (PUFAs) and a potentially better yield of DHA and EPAthrough fermentation (Barrow, 2010). As well as applications in mari-culture, DHA and EPA are prescribed in diets due to their clinical impor-tance to human health (Lewis et al., 1999). Thraustochytrids couldtherefore emerge as an alternative to fish oil with the potential for com-mercialisation andmay grow and accumulate lipids by utilising low costsubstrates such as bio-diesel derived glycerol (Chi et al., 2007; Ethier etal., 2011; Pyle et al., 2008). This review highlights some of the significantprogress made in last two decades and discusses the potential applica-tion of thraustochytrids for omega-3 fatty acids in biotechnology byemploying aspects of metabolic engineering and media optimisationstrategies with a brief discussion of the isolation and identification ofthraustochytrids.

1.1. Omega-3 fatty acids

PUFAs constitute a large group of fatty acids containing long chaincarbonic molecules that include ω-3-fatty acids (Lozac'h, 1986). Omega,

‘ω’ is the position of thefirst double bondwhen counted from themethylend and the number ‘3’ refers to the number of carbon atoms at that po-sition from themethyl end (Fig. 1).When counting thefirst occurrence ofdouble bonds from the carboxyl end, delta, ‘Δ’ is used to indicate thatposition. The molecular structure of the fatty acids consists of an evennumber of carbon atoms (4 to 24)with diverse saturations (0 to 6 doublebonds). Docosahexaenoic acid (DHA) and eicosapentaenoic acid (EPA)are two members of the ω-3 family. The prefixes ‘docosa’ and ‘eicosa’are of Greek descent, meaning the 22 and 20 C-atoms present in DHAand EPA, which contain 6 and 5 cis-double bonds respectively (Lozac'h,1986).

1.1.1. Omega-3 fatty acids as nutraceuticalsOngoing debate about appropriate levels of long-chain fatty acid

consumption has led to some confusion among consumers. The foodstandards codes for Australia do not prescribe specific quantities of ω-6 and ω-3 fatty acid content in the food products (Food standardsCode, Australia, 2008). The Food standards Code specifies only theamount of PUFAs and total saturated fat content, which is inadequatefor consumers wishing to make informed decisions about their dietaryintake. PUFAs originate from plants are consumed in higher abundancethan those from fish, so that plant ω-6 PUFA are consumed in higherquantities than ω-3 PUFA (Newton, 1998). This over consumptionof ω-6 relative to ω-3 oil has been linked to increased risk of cancer,diabetes, cardiovascular and neurodegenerative diseases (Simopoulos,2006). To restore a balance, consumption of ω-3 fatty acids should beincreased compared to ω-6 fatty acids. ω-6 to ω-3 PUFA ratios of be-tween 5:1 and 3:1 have been suggested as optimum for human con-sumption (Simopoulos, 2008). A recent report by the National Healthand Medical Research Council included references to the amountand types of fat consumed. The consumption of PUFAs at an average

1 2 3 4

H3C — CH2 – C = C – CH2 – C = C – CH2 – C = C – (CH2)7 – COOH

H H H H H H

16 15 14 13 12 11 10 9

Omega

Carbon

Double bond is present between 3rd

carbon (from -carbon) and 4th carbon

18: 3; 9, 12, 15

The other -3 fatty acids are

(a) Eicosapentaenoic acid 20:5; 5, 8, 11, 14, 17

(b) Docosapentaenoic acid 22:5; 7, 10, 13, 16, 19

(c) Docosahexaenoic acid 22:6; 4, 7, 10, 13, 16, 19

Fig. 1. Chemical structure of ω-3-fatty acids.

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of 6% of total energy and not exceeding 10% was recommended, with areduction in the consumption of saturated fatty acids tomaintain levelsat 8–10% of total energy intake (National Health and Medical ResearchCouncil, Australia, 2003). In the revised versions of the same report,the recommended intake of long chain fatty acids (DHA/EPA/DPA) inorder to lower the risk of chronic diseases was 610 mg/day for menand 430 mg/day for women (National Health and Medical ResearchCouncil, Australia, 2005).

The human health benefits of DHA and EPA have been extensivelyevaluated through a wide range of clinical studies (Barrow and Shahidi,2007; Kang and Weylandt, 2008; Takahata et al., 1998). Omega-3 fattyacids have been shown to help prevent heart attack as well as decreasethe overall risk of cardiovascular disease in general (Masson et al.,2007; Russo, 2009). DHA is ‘physiologically essential’ for brain andeye development, particularly for infants, which has led to the additionof DHA tomost infant formulas andmanyother infant related food prod-ucts (Birch et al., 2007; Innis, 2008). Clinical trials with DHA-enricheddiets have shown improvement in the learning capacity of school-agedchildren (Richardson and Montgomery, 2005).

The biochemical and biological functions of DHA were studied tobetter understand why it is required by the central nervous system.The accumulation of DHA in the neuronal membranes is a significantindicator of brain development and endurance (Kim, 2008). It wasobserved that omega oils play a crucial role in visual development,particularly in infants supplemented with diets rich in DHA and Ara-chidonic acids (AA) (Birch et al., 2007). A major contribution of lipidperoxidation products was reported in the DHA-stimulated apoptosisof cancerous cells (Siddiqui et al., 2008). Positive effects on visualfunctions and motor skills, such as coin sorting and dynamicbalance were observed in children with phenylketonuria when theirdiet was supplemented with DHA (Ryan et al., 2010). DHA and EPAinhibit the proliferation of vascular smoothmuscle cells, thus contribut-ing to the prevention of atherosclerosis disease (Horrocks and Yeo,1999). A diet with inadequate DHA during infancy and childhoodmight lead to inhibited development of the brain (Innis, 2008). The ef-fect of dietary ω-3-fatty acids on human colon cancer cells is also posi-tive acting to hinder the escalation of the deadly carcinomas (Kato et al.,2002), while the type of fat supplement chosen had a significant effect,the benefits appeared insensitive to the quantity administered. Thissuppression of tumour-genesis can be accredited to the diminished ac-tion of the genes involved in the growth of the tumour. Aside frombeingconsidered an inhibitor of carcinomas, DHA was found to exhibitchemo-protective properties towards normal tissues, facilitated by pro-tectins, a downstreamproduct of DHA synthesis whichmay be of specif-ic importance to sufferers of neuroblastoma and medulloblastoma(Gleissman et al., 2010).

DHA also plays a vital role in thraustochytrid cell life (Lewis et al.,1999). A study to investigate the biological significance of DHA inthraustochytrid cells concluded that DHA is conserved in the formof fatty acid energy reserves and utilised during starvation (Jain etal., 2007). A decline in total lipids was observed with the extensionin the starvation phase.

1.1.2. Biosynthesis of omega-3 fatty acidsAnimals, including humans, lack the Δ-12 and Δ-15 desaturase en-

zymes, which are essential for the synthesis of ω-3 and ω-6 fattyacids. As such they are not capable of building long chain omega fattyacids, and must acquire them from their diets. Humans and other ani-mals obtain some DHA and intermediate products such as EPA throughbioconversion of α-linolenic acid (18:3, ω-3), and some from directconsumption of DHA itself (Innis, 2003). ω-3 fatty acids are typicallysynthesized fromα-linolenic acids acquired from dietary plant sources,and are hence regarded as biological sources of EPA and DHA. Furthermetabolisation of α-linolenic acids is characterised by the action ofthe Δ-6 desaturase enzyme for the saturation of the fatty acid, followedby the action of the elongase enzyme for the addition of a carbon atom

to themolecular chain, leading to action byΔ-5 desaturase to form EPA.Earlier studies revealed the occurrence of the desaturation process ofthe long chain fatty acids in endoplasmic reticulum (Innis, 2003).

b

a

Fig. 2. a— PKS pathway for the synthesis of DHA in Schizochytrium sp. (figure reproducedand modified from Ratledge, 2004) b — Biosynthesis of ω-3 fatty acids (Innis, 2003).

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Interestingly, the contribution of Δ-4 desaturase in the final steps ofDHA synthesis (22:5ω–3 to 22:6ω–3) was established in the microbialpathway. This part of the synthesis was later interpreted in a differentmanner after the discovery of 24:5ω–3 with the help of the elongationof 22-C-atoms in the mammalian pathway (Sprecher et al., 1999). This24:5ω–3 position is desaturated to yield 24:6ω–3 in the peroxisomeswhere DHA is formed by the partial oxidation process (Ferdinandusseet al., 2001). Discovery of the mechanism for DHA synthesis may aidin the treatment of peroxisomal disorders such as Zellweger syndromein infants, using DHA-supplemented diets. The unsaturation (additionof a double bond) and elongation (addition of a 2-C unit) mechanismin the synthesis of these omega oils are shown in Fig. 2b.

2. Microorganisms producing fatty acids

Numerous organisms including bacteria, yeast, filamentous fungi, andmicroalgae have been reported to produce different types of fatty acids(see Table 1). These fatty acids include long-chainω-6 andω-3polyunsat-urated fatty acids such as Gamma linolenic acid (GLA), AA, DHA and EPA.

2.1. Other PUFAs

Plants were initially considered to be the main source of PUFAs forhuman health, particularly GLA (Gamma linolenic acid), which isavailable commercially. However, the main drawback of utilizingplant-based PUFAs is the fact that more genes must be engineeredto produce PUFAs in commercial quantities. In the absence of thistechnology, there is reduced likelihood of DHA production using engi-neered oil seed crops (Alonso and Maroto, 2000). Microbial produc-tion of other long chain PUFAs such as arachidonic acid (AA, 20:4;ω-6), dihomo-γ-linolenic acid (DGLA, 20:3; ω-6), and γ-linolenic acid(GLA, 18:3; ω-6) has also been reported (Ratledge, 2001). The firstcommercial oil produced by microbes was of GLA. GLA was extractedpredominantly from lower fungi (Phycomycetes) as other sourcessuch as Pythium debaryanam (fungus) had lower quantities of lipids(Shaw, 1965). Mortierella alpina produced total lipid content as 26% ofbiomass with 80% of total fatty acids as AA. This fungus required longcultivation periods of more than two weeks, hindering its commerciali-sation (Totani and Oba, 1987). However, AA (11% of TFA) was reportedto be produced in the fungus, Mortierella alpinapeyron (Kendrick and

Ratledge, 1992). Other fungi such as Conidiobolus nanodes and Ento-morphthora exitalis produced both ω-3 and ω-6 PUFAs but the outputof ω-6 fatty acids was significantly higher than that of ω-3 fatty acids(Kendrick and Ratledge, 1992). Mortierella alliacea was reported to bethe industrial producer of AA and with the supplementation of plantoils EPA productivity was also documented (Jermsuntiea et al., 2011).Mortierella sp. was found to be resistant to the high glucose concentra-tions (4–11%) that elicited an improvement in AA production (Suzukuet al., 2010) and other PUFAs such as GLA and DGLA. These organismscan be classified as oleaginous as they have more than 20% lipids accu-mulation in the cell.

2.2. Omega-3 fatty acids: DHA and EPA

Many bacterial strains, including Shwenella sp. and Colwellia sp.,have been reported to produce DHA and EPA (Bowman et al., 1997,1998; Nichols and McMeekin, 2002). Some fungal strains belongingto Mortierella sp. were also reported to be producers of EPA (Jacobset al., 2009). These species were also found to produce enzymes andcarotenoids other than PUFAs. Higher EPA production was achievedwith the fungus Pythium irregulare in a 14 L fermentor using lactose,rather than glucose, as a carbon source with an EPA output of 25%of total lipids content. This represented the highest reported EPA out-put using fungi (O'Brien et al., 1993). It has been demonstrated thatPichia methanolica, can be developed as a DHA enriched organism ifit is cultured in DHA-supplemented medium and also observed thatvarious nutrients promoted DHA accumulation (Aoki et al., 2002).This DHA enriched yeast was promoted over other microorganismscontaining DHA as triglycerides or phospholipids. Temperature cul-turing (12–25 °C) using Mortierella sp. under different temperatureregime assisted in EPA production (Suzuku et al., 2010). Arthrospiramaxima, a mesophilic strain of cyanobacteria, produced DHA (28% ofTFA) at 20 °C (Hu et al., 2011). However, bacteria are not consideredviable for the production of PUFAs as they have lower lipid accumula-tion (2–5%). In addition, the presence and characteristics of some unde-sirable lipids hinder the possibility of promoting bacteria as a suitablePUFA producer. Moreover, there is some scepticism regarding the qual-ity of bacterial EPA and DHA (Ratledge, 2001). Although bacteria arenot toxic to animals, there is no record of the use of bacterial lipids foranimal consumption. Marine microalgae have been found to be thebest alternative to fish oil in terms of ω-3 fatty acids production with alow likelihood of exhibiting the taste, odour and instability linked tofish oils (Barclay et al., 1994). In fact, microalgae produced 1–2 timesmore ω-3 fatty acids than bacterial and fungal cultures.

3. Thraustochytrids: occurrence and methods for isolation

Mangrove leaves were found to be a plentiful thraustochytrid habi-tat, as thraustochytrids feed on the leaves resulting in their degradation(Bremer, 1995; Wong et al., 2008). Invertebrates such as nudibranchsand squid are also reported to be associated with thraustochytrids interms of their presence in these invertebrates (Jones and O'Dor, 1983;McLean and Porter, 1982). However, the type of biological associationsthat exist between thraustochytrids and the invertebrates is yet to beexplored (Rabinowitz et al., 2006). Researchers have isolated thrausto-chytrid strains from marine and coastal seawater environments (Burjaet al., 2006; Kimura et al., 1999; Yang et al., 2010), as well as low tem-perature environments (Zhou et al., 2010) and colonial tunicate Botryl-lus schlosseri (Rabinowitz et al., 2006). Salinity is a vital environmentalvariable which may limit the occurrence of thraustochytrids as theycan tolerate salinity to a limited extent (Bremer, 2000).

3.1. Pollen baiting and direct plating

Baiting techniques and direct plating methods were recommendedfor the isolation of thraustochytrid strains (Fig. 3) (Bremer, 2000).

Table 1PUFAs (% Total fatty acids, TFA) producing microorganisms, mainly, oleic acid, linoleicacid, ALA, GLA, DGLA, AA, DHA and EPA.

Name of microorganism Fatty acids (% TFA) Reference

Pythium debaryanam γ-Linolenic acid (10.4) Shaw (1965)Mortierella alpina Arachidonic acid

(68.5–78.8)Totani and Oba (1987)

Mortierella alpinapeyron Arachidonic acid (11) Kendrick and Ratledge(1992)Saprolegnia parasitica AA (19) / EPA (18)

Vibrio sp. DHA and EPA Ando et al. (1992)Pythium irregulare EPA (25.2) O'Brien et al. (1993)Candida guilliermondii DHA (6.7%) and EPA

(2.8)Guo and Ota (2000)

Colwellia sp. and DHA (0.7–8) Bowman et al. (1997, 1998)Shwenella sp. EPA (2–22) Nichols and McMeekin

(2002)Pichia methanolica DHA (32) Aoki et al. (2002)Mortierella sp. EPA Jacobs et al. (2009)Mortierella sp. AA (25.9–53.8) Suzuku et al. (2010)

GLA (4.3–4.7)DGLA (4.7–4.9)EPA (6.1)

Mortierella alliacea YN-15

ALA (33–41) Jermsuntiea et al. (2011)EPA (1.3–13)

Rhodotorula mucilaginosaAMCQ8A

ALA (4.1–5.4) Gupta et al. (2011)Oleic acid (45.4)Linoleic acid (24.7)

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The pollen baiting method is a commonly used procedure for isolationof thraustochytrids (Burja et al., 2006; Jakobsen et al., 2007). Sterilizedpollen grains are interspersed on the surface of the water sample con-taining antibiotics and antifungal agents and incubated for 1–2 weeks.Once thraustochytrid cells are observed to be growing on the pollensurface, they are spread plated on agar plates and selectively grown inliquid medium to obtain pure cultures (Wilkens and Maas, 2012). Thedirect plating method includes the distribution of the samples on agarplates under aseptic conditions (Bowles et al., 1999; Bremer, 2000).

3.2. Antibiotics and antifungal supplemented medium

Bacterial and fungal contaminations should be avoided during theisolation of thraustochytrids, as the isolation temperatures (18–25 °C)

may favour the growth of contaminant microbes. Antibiotics such aspenicillin and streptomycin (Burja et al., 2006; Quilodran et al., 2010),and rifampicin and antifungal drugs such as nystatin (Jakobsen et al.,2007; Wilkens and Maas, 2012) and amphotericin B (Taoka et al.,2010) have been used in the agar plates and liquidmedium tominimisethe growth of bacterial and mycelial colonies.

3.3. Identification techniques

Biochemical and molecular techniques have been used recentlyto identify newly isolatedmarine organisms (Gupta et al., 2011). Stainingprocedures usingfluorochrome for direct detectionbased onmorpholog-ical characteristics of the organism have been used (Raghukumar andSchaumann, 1993). Species-level identification is based on 18s rDNA

c d

e f

a b

Fig. 3. Light microscopy of thraustochytrid cells; a — ATCC Thraustochytrium PRA 296 (scale bar: 20 μm), b — cluster of fresh zoospores on the surface of pine pollen (our isolates)(scale bar: 20 μm), c — single thraustochytrid cell (sporangium) attached to pine pollen (scale bar: 20 μm), d — several zoospores released from a mature sporangium (scale bar:20 μm), e — mature sporangium (scale bar: 10 μm), f - exponential phase of growth of our isolate (scale bar:20 um).

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sequencing for establishing a close taxonomic relationship to existingThraustochytrium species (Mo et al., 2002). The taxonomic relationshipis further described in Section 4. The 18s rDNA sequencing technique isperformed after obtaining the 18s rDNA gene from the desired microor-ganismand extending the base-pairs using specific primers to get the 18srDNA sequence. This sequence is alignedwith the available thraustochy-trid sequences to establish the evolutionary relationship between iso-lates and available cultures leading to the identification of isolates.

3.3.1. Biochemical identificationA fluorescent staining technique has been developed to give an es-

timation of thraustochytrids cell numbers which involves the reactionbetween fluorochrome (acriflavine), the sulphated cell wall and theirnuclei (Raghukumar and Schaumann, 1993). The cell wall fluorescesred and the nuclei green, differentiating them from other protiststhat fluoresce only green (Moss, 1986). This technique has some lim-itations as there is an absence of cell wall material in the zoospores ofmost thraustochytrid species (Moss, 1986) and thraustochytrid cellswith a thin cell wall are undetectable. A thraustochytrid-specific de-tection method employing a fluorescence in-situ hybridization tech-nique (FISH) has been established using an rRNA targeted ThrFL1probe (Takao et al., 2007). This technique successfully overcomesthe aforementioned limitation of acriflavine direct detection (AFDD)due to the thinner wall of thraustochytrid cells (Moss, 1986). FISHtechnique involved the hybridization between the probe ThrFL1 andrRNA particles which assisted the detection of zoospores and youngsomatic cells. Sudan black staining has also been used for stainingthe thraustochytrid cells (Subramaniam and Chaubal, 1990; Wonget al., 2008).

3.3.2. Molecular identificationMolecular identification of thraustochytrids is based on the 18s

rDNA sequencing technique which characterises the taxonomic rela-tionship of unknown isolates with respect to known isolates. It in-volves PCR based amplification of the 18s rDNA gene requiring aset of primers designed to use sequences of known thraustochytridstrains and comparative phylogenetic analysis to ascertain novel spe-cies (Bongiorni et al., 2005; Mo et al., 2002; Yokoyama and Honda,2007). This is a sensitive tool to determine or establish relationships be-tween thraustochytrid species. The multiple sequence alignment of theisolated 18s rDNA sequences and available sequences gives a branchingrelationship between the new isolates and known thraustochytrid spe-cies. The use of neighbour-joining and maximum likelihood methodsfor determining taxonomic relationships has been assessed with thehelp of analytical software packages such as PHYLIP 3.5c andPHYLOWIN (Bongiorni et al., 2005). PHYLIP 3.5c is the Phylogeny inter-ference package involving the use of 31 programs using molecular se-quence data, distance matrix data, gene frequencies and tree plot data(Felsenstein, 1993). The PHYLOWIN package assists the phylogenetictree constructionwith theuse of neighbour joining,maximumparsimo-ny and maximum likelihood method including bootstrap analysis(Galtier et al., 1996).

3.4. Ultra structure of thraustochytrids

Thraustochytrids can accumulate greater than 50% of their dryweight as lipids, of which more than 25% is normally DHA (Bajpaiet al., 1991; Raghukumar, 2008; Sijtsma and Swaaf, 2004; Singh andWard, 1997; Yaguchi et al., 1997; Yokochi et al., 1998). Transmissionelectron microscopy (TEM) has revealed ample lipid bodies in the cy-toplasm of thraustochytrid cells, which was later confirmed by SudanBlack B staining (Subramaniam and Chaubal, 1990). This ultrastruc-ture study which used TEM illustrated that thraustochytrid cells areheterogeneous in size, approximately 6–21 μm in diameter with agranular cytoplasm containing oil micelles. An association of lipidbodies was observed with the branched, hollow membrane-like fine

structures containing light and dark bands. The pattern of these finestructures has been described as the ratio of saturated and monoun-saturated fatty acids to polyunsaturated fatty acids (Wong et al., 2008).These fine structures were hypothesised to be linked with lipid synthe-sis, although they lose their structure almost completelywith increase inthe size of the lipid bodies (Weete et al., 1997). A relationship betweenlipid body formation and Endoplasmic reticulum (ER) in Schizochytriumlimacinum SR21 was furthermore observed (Morita et al., 2006). Lipidbodies surrounded by ER have been extensively studied in the oleagi-nous fungus Mortierella ramanniana (Kamisaka et al., 1999) whereasanalogous research in Labyrinthulomycota (which includes thrausto-chytrids) has yet to be undertaken.

4. Classification of thraustochytrids

There has always been confusion among researchers regarding thetaxonomical classification of thraustochytrids. The taxonomical struc-tures have been first established and then abolished quite often inthe process of developing taxonomy for thraustochytrids, which hasadded to ambiguity regarding its structural and functional behaviour.During the early 1970s, electron microscopic studies of the ultrastruc-ture of protists determined the phylogenetic relationship of thrausto-chytrids and discovered that they were loosely related to Oomycetes(Kazama, 1972; 1974). With the development of DNA sequencingmethods and electron microscopic studies of ultrastructure, thrausto-chytrids were subsequently designated as a unique group.

Before being incorporated into Oomycetes, thraustochytrids hadbeen included within Phycomycetes (algae-like fungi) (Barr, 1981).Subsequently, thraustochytrids were included within the Phylum Het-erokonta within the Chromophyta (Stramenopiles) kingdom (Cavalier-Smith, 1993; Honda et al., 1999). The genera included in this groupare Thraustochytrium, Schizochytrium, Japonochytrium, Aplanochytrium,Elina, Labyrinthula (or Labyrinthuloides or Labyrinthulomyxa). Labyr-inthula has been included in Labyrinthulaceae family whereas Thrausto-chytriaceae family includes Thraustochytrium. Hence, after numerousrevisions, thraustochytrids proved to be a distinctive and characteristicdivision of protists in which the members can be classified under theThraustochytriales order (Metz et al., 2010).

An example: Domain: EukaryotaKingdom: ChromophytaPhylum: HeterokontaFamily: ThraustochytriaceaeOrder: ThraustochytrialesGenera: Thraustochytrium

5. Production of omega-3 fatty acids

The production of ω-3 fatty acids through the fermentation pro-cess began with the introduction of various nutrient sources andbio-processing strategies. These strategies are aimed at higher pro-ductivity, thus minimising process cost.

5.1. Cultivation medium and bioprocess engineering

Nutrient conditions must be optimised for any new isolate to obtainthe maximum level of omega oil productivity (Shene et al., 2010). Glu-cose is frequently used in microbial fermentation processes as it is con-sidered to be a cheap raw material compared to other carbohydrates(Puri et al., 2011; Singh et al., 1996), as well as being readily targetedfor utilisation by most microorganisms. While monosaccharide's sup-port rapid thraustochytrid growth as well as high DHA yields, poorcell growth was observed when using disaccharides and polysaccha-rides in the fermentation medium (Burja et al., 2006). This observationcontrastedfindingswhere allmono-, di- and polysaccharides supported

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growth and resulted in high lipid content with Thraustochytrium aur-eum ATCC 34304 (Bajpai et al., 1991). Earlier, starch supplementedmedia produced higher DHA yields compared with those using glucose,sucrose, corn and canola oil, although the resulting biomass yields weresimilar (Li and Ward, 1994). This study highlighted the importance ofthe supplied carbon source. Alternating carbon sources such as crudeglycerol, waste water from barley distillation, soybean cake, and liq-uid residues from beer and potato processing were investigated tomake the process economical without compromising on high DHA out-put (Quilodran et al., 2009; Yamasaki et al., 2006; Zhu et al., 2008). Thedifferent carbon and nitrogen sources used with thraustochytrids forproducing omega oil are listed in Table 2.

Many studies used glycerol in the medium while working on Schi-zochytrium limacinum SR21 (Chi et al., 2007; Ethier et al., 2011; Pyleet al., 2008) as well as complex carbon sources such as sweet sorghumjuice (Liang et al., 2010) for DHA production. Liquid residues obtainedfrom the brewing industry and potato chips processing units have alsobeen used to produce DHA (Quilodran et al., 2010). Biodiesel-derivedglycerol has vast potential to minimise production costs because of itslower price compared to other commercially available carbon sources(Pyle et al., 2008). Despite this, crude glycerol obtained from the biodie-sel producing industry requires pre-treatment to remove the soap thatmay retard cell growth and lead to low DHA yields (Chi et al., 2007;Ethier et al., 2011; Liang et al., 2010; Pyle et al., 2008). Glycerol couldstill be superior option for the fermentation process required to produceDHA and EPA in thraustochytrids. Aside from glycerol, carob pulp syruphas been used as a carbon source for the production of DHA in Crypthe-codinium cohniimicroalgae, yielding the best output in terms of biomassand DHA productivity when compared to glucose as the carbon source.Carob syrup proved to be an appropriate nutrient source for DHA pro-duction in Crypthecodinium cohnii (Mendes et al., 2007).

Inorganic nitrogen in the form of ammonium sulphate may beused as a sole nitrogen source but glutamate, aspartate, their amidesand alpha alanine can result in a higher biomass yield (Goldstein,1963). This result is consistent with the research of Jakobsen et al.

(2008) in which the effect of nitrogen in the form of monosodiumglutamate as the sole nitrogen source was studied and observed to in-crease biomass growth and lipid accumulation, relative to inorganicnitrogen growth. Nitrogen depletion from the fermentation mediumelevated the lipid accumulation from 13 to 55% w/w of the dry bio-mass of Aurantiochytrium sp. strain T66. Initiation of the lipid accu-mulation may be attributed to inadequate levels of oxygen and bylimiting the nitrogen and phosphorous sources in the medium usedfor ω-3-fatty acid production. With this strategy, a high lipid contentof 54–63% w/w of dry weight of the Aurantiochytrium sp. cells was ob-served (Jakobsen et al., 2008). Abundant carbon and low nitrogen inthe fermentation medium will therefore result in the high level ofoil accumulation found in thraustochytrids. Due to the minimal levelsof nitrogen, protein and nucleic acid synthesis is limited, converting thecarbon sources into storage oils. The higher C: N ratio was found to sup-port higher DHA production in thraustochytrids (Bowles et al., 1999;Burja et al., 2006; Yokochi et al., 1998). Glucose concentrations in therange of 6–10% resulted in higher levels of DHA (Wong et al., 2008)whereas more than 10% of glucose was reported to retard the growthof thraustochytrids (Yokochi et al., 1998).

Nutrient sources which may result in higher production of ω-3fatty acids in thraustochytrids included polyoxyethylene sorbitanmonooleate (Tween 80), acetyl-CoA and nicotinamide adenine dinu-cleotide phosphate (NADPH). Tween 80 increased biomass growthand total lipid content in Thraustochytrium aureum ATCC 34304 with-out modifying the DHA content (Taoka et al., 2011). The rise in thetotal lipid content was accredited to the presence of oleic acid in theform of oleate in Tween 80. Furthermore, new strategies that in-volved the introduction of Acetyl-CoA and NADPH in the form of eth-anol and malic acid, respectively, during the fermentation process atdifferent periods of lipid accumulation were implemented to improveDHA yield. An increase from 35 to 60% DHA content has beenreported with the addition of malic acid (enhancing NADPH) at therapid lipid accumulation phase (Ren et al., 2009). However, addedNADPH was ineffective on total lipid accumulation but altered the

Table 2Summary of different carbon and nitrogen sources used for omega-3 fatty acid production using different Thraustochytrium species.

Strain Carbon source Nitrogen source % of TFA Cell Mass(gL−1)

Fermentationvolume (ml)

Reference

DHA EPA

T. aureum 34304 Glucose/Fructose Peptone 41–75 1.2–5.2 1.1–5.0 50* Bajpai et al. (1991)Starch TryptoneMaltose Yeast extractLinseed oil Malt extract

MSGT. roseum 28210 Starch (NH4)2SO4 48.3–58.2 – 6.1–17.1 50* Singh and Ward (1997)

MSG (Fed batch)Schizochytrium sp. SR21 Glucose (NH4)2SO4 33.3–38.6 – 21.9–59.2 3000# Yaguchi et al. (1997)

Corn steep liquorS. limacinum SR21 Monosaccharides Yeast extract 6.0–43.1 b1 12 10* Yokochi et al. (1998)

Oleic acid Con steepLinseed oil liquor

S. mangrovei G-13 Glucose Yeast extract Peptone 28 – 14 2000# Bowles et al. (1999)S. limacinum KH 105 Glucose Yeast extract Poly-peptone 34.9 1.6 11.5 50* Aki et al. (2003)Schizochytrium sp. F26-b Glucose Yeast extract 31.8 1.2 3.5 200* Abe et al. (2006)ONC-18 Glucose Yeast extract 16–32 – 10–26 30* Burja et al. (2006)

Glutamic acid 4900#

Schizochytrium sp. KH 105 Glucose Waste water for barley distillery 25.8 – ~26 2000# Yamasaki et al. (2006)S. limacinum SR21 Crude glycerol Yeast extract 33.6 – 22.1 50* Chi et al. (2007)

Ammonium acetateS. mangrovei Sk-02 Coconut water Glucose Yeast extract 20 – 28 100* Unagul et al. (2007)S. limacinum SR21 Crude glycerol Yeast extract Peptone 18.26–53.05 – 2.5–8 50* Pyle et al. (2008)Aurantiochytrium mangrovei MP2 Glucose Yeast extract 3 – 25.4 50* Wong et al. (2008)S. limacinum OUC88 Glucose Soybean cake hydrolysate 4.08 (gL−1) – 25.92 100* Zhu et al. (2008)Thraustochytriidae sp., M12-X1 Beer residues Yeast extract MSG 39.5–61.7 7.4–11.3 2.3 100* Quilodran et al. (2009)S. limacinum SR21 Sweet sorghum juice – 34.28 0.61 9.4 100* Liang et al. (2010)S. limacinum SR21 Crude glycerol Yeast extract 24.86– – 4–15 4500# Ethier et al. (2011)

Peptone 31.09 (Continuous)

*Shake flask cultivation, #Bioreactor.

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fatty acid composition, whereas use of acetyl-CoA demonstrated anincrease of 35% in lipid accumulation with only a minor change inDHA content. The combination of an acetyl-CoA and NADPH strategymay offer both high lipid accumulation and increased DHA yield.

5.2. Factors affecting omega-3 fatty acid production

Fermentation conditions have been altered to enhance the pro-duction of PUFAs in thraustochytrids and their optimisation hasresulted in a high level of omega oil production (Bajpai et al., 1991;Wu and Lin, 2003). In testing for optimal PUFA yield, chemical param-eters, such as carbon and nitrogen sources in different concentrations,and physical parameters such as initial pH, temperature and inoculumage were adjusted. While temperature was found to have no notableeffect, all other parameters were found to affect fatty acid production(Bajpai et al., 1991). With the aim of high DHA productivity, a temper-ature shift regime resulted in a DHA content of 52% of total fatty acids.A decline in temperature from 30 to 20 °C yielded high levels of DHAmaking 30 °C the optimal culture temperature for the highest biomassgrowth (Zeng et al., 2011). A low culture temperature may affect thelipid synthesis in the marine microbes isolated from low temperateregions (Fang et al., 2000). DHA production might be higher or lowerin the previously reported thraustochytrid strains (Burja et al., 2006;Huang et al., 2003; Unagul et al., 2005) as they have been isolatedfrom different environments, naturally altering their physiology. Thisdifference in the natural conditions may contribute to variable produc-tion of omega oils, ranging from strain to strain.

The transformation of saturated fatty acids into unsaturated fattyacids was found to occur when oil producing strains grow in oxygenabundant environments (Raghukumar, 2008). The volume of the liq-uid medium was also assessed, and it was found that the dissolvedoxygen was affected, thus resulting in a lower DHA yield (Zhou et al.,2010). A large volume of liquid medium (more than 40 mL in 250 mLflask) resulted in a low DHA yield. Response surface methodology wasfound to be suited for optimising the effect of multiple factors on fer-mentation yields (Kalil et al., 2000). The experimental value was com-parable to the predicted yields with 97% closeness. This technique canassist in the optimisation of DHA yields by using various models suchas fractional factorial, central composite design and RSM (Wu and Lin,2003).

The production of omega-3 fatty acids in large-scale fermentorshas been reported elsewhere (Bajpai et al., 1991; Chi et al., 2009;Hong et al., 2011; Jakobsen et al., 2008). The growth of thraustochy-trid strains in a bioreactor differ significantly, depending on the straincharacteristics. T. roseumwas found to grow better in a flask than in astirred tank fermentor (Ilda et al., 1996). However, Schizochytrium sp.SR21 exhibited significantly better growth under bioreactor conditions(Nakahara et al., 1996). Of sixty-eight isolates comprising the genera

Schizochytrium and Thraustochytrium, 54 were found to produce DHAin the range of 22–80% of total C20 to C22 content of the cells (Burjaet al., 2006). This observed DHA yield is in agreement with the findingsof Huang et al (2003), who reported 53%DHAof total fatty acids presentin the thraustochytrids isolated from cold environments. Some thraus-tochytrid isolates with DHA content are summarised in Table 3.

Growth-dependent lipid accumulation was achieved by introduc-ing a nitrogen source in the medium in the form of ammonia(300–400 mg L−1) in a fed batch system using Schizochytrium sp.G13/2S (Ganuza et al., 2008) which resulted in the highest growthrate irrespective of the degree of lipid accumulation. This study ex-plored the lipid accumulation as a result of nitrogen limitation in thefermentation medium (Jakobsen et al., 2008). Ganuza and co-workersalso demonstrated the use of sodium chloride as the only salt sourcein a defined medium. A two stage growth process of Schizochytriumsp. SR21was employed, which enabled a design to optimise the param-eters separately for biomass growth and lipid accumulation (Chi et al.,2009).

With the change in fermentation process, a continuous fermenta-tion was designed to study lipid accumulation by Shizochytrium sp.G13/2S in different glucose (7–40 g L−1) and glutamate (4–6 g L−1)concentrations (Ganuza and Izquierdo, 2007). This process stimulat-ed more lipid accumulation as compared to the batch fermentationwhere lipid accumulation ceased as a result of stalled cell growth. Afed batch system using Aurantiochytrium sp. KRS101 that resulted ina DHA productivity of 40% of TFA was reported (Hong et al., 2011).

6. Lipid extraction methods

When referring to a classic example of a lipid extraction technique,the Bligh and Dyer method (1959) is usually the first to be citeddue in large part to its simplicity in using common solvents such aschloroform, methanol and water mixtures. In the Bligh and Dyer meth-od, the lipids remain in the chloroform layer while themethanolic layercontains the non-lipids. This method has been modified by researchersto increase the actual output of PUFAs frommicrobes by combining lipidextraction with trans-esterification followed by recovery using a hex-ane and chloroform mixture (Burja et al., 2007; Lewis et al., 2000).The fatty acid quantification of PUFAs is complex due to their propensityfor oxidation. To avoid this, various modified methods using differentsolvent mixtures and disruption techniques based on sonication havebeen reported. Direct saponification and direct trans-esterification,the acid Bligh and Dyer method coupled with trichloroacetic acid andthe miniature Bligh and Dyer method were some of the modified ex-traction methods reported recently (Burja et al., 2007). Such methodsyielded higher amounts of lipids from Thraustochytrium and Schizochy-trium species. Among these, the miniature Bligh and Dyer method wasfound to be the easiest whereas the direct esterification method hadconstraints such as low fatty acid recovery. Often, modifications involv-ing some of the variables such as the time period of trans-esterificationand volume of the trans-esterification mixture are required to enhancethe extraction of fatty acids (Burja et al., 2007).

A modified trans-esterification method using sodium hydroxide inmethanol and Boron trifluoride in methanol was used to extract lipidsfrom fish oil (Armenta et al., 2009). Direct trans-esterification offreeze-dried thraustochytrid cells was performed and analysed froma calibration graph of standards. Both of these methods were foundto be efficient in measuring PUFAs in fish oil and thraustochytrid cells.

A solid phase extraction method was designed for fast and quanti-tative separation of lipid classes from microbial oils (Pinkart et al.,1998). The method proved to be a surprisingly effortless and cost-effective technique characterised by high yield and a purified formof lipid classes when operated with appropriate solvents, and wassubsequently performed for PUFA separation as well (Burja et al.,2007).

Table 3Several thraustochytrid strains isolated from different environments and their respec-tive DHA (% Total fatty acids, TFA) content.

Strain DHA(% of TFA)

Reference

Thraustochytrium aureum ATCC 34304 51 Bajpai et al. (1991)Schizochytrium sp. ATCC 20888 32 Barclay et al. (1994)T. roseum ATCC 28210 46–49 Li and Ward (1994)Schizochytrium sp. SR21 34 Nakahara et al. (1996)S. limacinum MYA 1381 34 Yokochi et al. (2003)Schizochytrium mangrovei Sk-02 28 Unagul et al. (2005)Thraustochytrium sp. ONC -T18 31.5 Burja et al. (2006)Schizochytrium sp.G13/2S 43–47 Ganuza and Izquierdo (2007)Schizochytrium sp. HX-308 60 Ren et al. (2009)Thraustochytriidae sp. C41 18.4–27.1 Quilodran et al. (2009)Thraustochytriidae sp. M12-X1 31.5–61.7Thraustochytriidae sp. Z105 32 Zhou et al. (2010)Aurantiochytrium BL10 29 Yang et al. (2010)

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The use of microbial samples for fatty acid analysis may be prob-lematic and ultimately inaccurate unless the process is performedfastidiously. The main steps in the process involve extraction, theconstructing of fatty acid methyl esters (FAMEs), GC optimisationand the usage of appropriate internal and external standards for de-termining the concentration (Masood et al., 2005; Schreiner, 2005).An improved protocol was derived with the addition of acetyl chlo-ride on a dry ice bath, followed by trans-esterification performed atroom temperature, and an analysis of GC data using relative responsefactors. This procedure was found to be applicable in the recovery ofω-3 fatty acids, DHA and EPA (Xu et al., 2010).

A new technique has also been formulated by Jacobsen and co-workers for lipid extraction with a modification of the previously de-scribed protocols. This modification involves heat treatment and pro-tease digestion of freeze dried thraustochytrid cells. The use of heatand an enzyme in the lipid extraction was a novel concept to beimplemented in the lipid extraction process (Jakobsen et al., 2008).

7. Quantification of fatty acids

Various analytical techniques have been reported for the quantifi-cation of fatty acids. These include urea fractionation (Abu-Nasr et al.,1954), thin layer chromatography (TLC) and preparative scale gaschromatography (Hardy and Keay, 1967). This was followed by highperformance liquid chromatography (HPLC) methods with somemodifications such as reverse phase HPLC using C18 column and sil-ver nitrate HPLC (Aveldano et al., 1983; Ozcimder and Hammers,1980; Scholfield, 1979). However, silver nitrate HPLC was found tobe unsuitable for mono and di-unsaturated fatty acids, as PUFAs areinadequately determined due to the effect of the double bond onthe carbon chain length. A method involving low temperature frac-tional crystallization of unwanted lipids was reported but was notapplicable to highly unsaturated fatty acids (Ackman, 1981). Otherchromatography techniques such as silica gel thin layer chromatogra-phy (Shantha and Ackman, 1991) and open column chromatography(Hayashi and Kishimura, 1993) were applied to large scale prepara-tions (Medina et al., 1995) following urea fractionation. A super criticalfluid extraction technology associated with the separation of PUFAsby gas chromatographywas reported in late 1990s (Walker et al., 1999).

Some limitations of these techniques should be noted, namelyi) low PUFA yield, ii) exhaustive preparation and iii) scale-up difficul-ty (Cartens et al., 1996; Medina et al., 1995).

The fatty acid methyl esters produced after the lipid extractionwere analysed by a more efficient and reliable technique using gaschromatography coupled with mass spectrometry using differentfatty acids as the internal standards. The samples could be identifiedwith the help of a chromatogram peak related to the retention timeof the particular fatty acid comparable to that of the internal stan-dards. Recently, this technology has been applied extensively (Renet al., 2009; Zeng et al., 2011).

8. Other metabolites produced by thraustochytrids

Thraustochytrids have been known to produce various secondarymetabolites such as carotenoids, sterols, steroids and surfactants (Fanand Chen, 2007; Lewis et al., 2001). In addition to this, they are alsoa potential source of enzymes and extracellular polysaccharides.

8.1. Carotenoids

Thraustochytrids are a promising source of carotenoids as well asPUFAs. The production of carotenoid pigments such as astaxanthin, ze-axanthin, canthaxanthin, echinenone, phoenicoxanthin and β-caroteneby Thraustochytrium sp. has been reported (Burja et al., 2006; Carmonaet al., 2003). Thraustochytrid strain KH-105was reported byAki and co-workers as β-carotene and xanthophyll accumulator (astaxanthin and

canthaxanthin). Xanthophyll productionwas found to respond inverse-ly proportional to nitrogen concentrations. They also observed thatmoderate nitrogen concentrations of 0.3 and 6% support the productionof astaxanthin and canthaxanthin, respectively (Aki et al., 2003). Astax-anthin and canthaxanthin exhibit antioxidant and chemo-protectiveproperties, making them potential candidates as food additives thatmay act against the development of neurodegenerative disorders(Yuan et al., 2011). Various methods, using the combination of organicsolvents and ultrasonic bath, have been examined to determine carot-enoid content present in the Thraustochytrium strain ONC-T18(Armenta et al., 2006). Semi-fragile cell walls in thraustochytrid strainKH-105 aided the downstream processing of the pigments with com-prehensive recovery using a solvent such as acetone. Amutation strategywas furthermore carried out to enhance the astaxanthin productivity of S.limacinum by treating the cells with N-methyl-N′-nitro-N-nitrosoguanidine (NTG), resulting in intense colour development in thecolonies (Chatdumrong et al., 2007).

As β-carotene acts as a precursor for vitamin A, sufficient intake ofβ-carotene can help to prevent diseases caused by vitamin A deficien-cy, including blindness, immune dysfunction and skin disorders(Fierce et al., 2008). More than 80% of β-carotene is synthesized bychemical processes. The chemosynthetic pathway produces relativelymore trans-stereoisomers of beta-carotene than microbial processes.Trans-stereoisomers are less competent antioxidants, and thereforeless desirable for medical applications (Del Campo et al., 2007; Shaishet al., 2006). Carotenoids produced via microorganisms are widely ac-cepted and considered to be a safe drug component. The halo-tolerantmarinemicroalgaDunaliella sp. is considered as a large carotenoid accu-mulator but high carbon dioxide consumption combined with low pro-duction efficiency and poor control, and the high cost and requirementsof land, limit the expansion ofmass cultures ofDunaliella for β-caroteneproduction (Ye et al., 2008). Thraustochytrids can be used as an alterna-tive for carotenoid production.

8.2. Enzymes

Thraustochytrids have been known to produce a range of enzymessuch as proteases, lipases, esterase, acid and alkaline phosphatase,cellulases and xylanases (Raghukumar, 2008). The extracellular cellu-lolytic activity was studied in 19 strains of thraustochytrids using car-boxymethyl cellulose (CMC) as the substrate, with 14 strainsconfirming cellulase activity by hydrolysing the substrate (Naganoet al., 2011). A three-fold increase was observed in the extracellularlipase activity of two thraustochytrid strains AH-2 and TZ with thehelp of various optimisations of the culture conditions (Kanchana etal., 2011).

8.3. Extracellular polysaccharides

Thraustochytrids are reported to produce Extracellular Polysaccha-ride (EPS) that include sugars as the major component (39–53%) withthe presence of proteins, lipids, uronic acids and sulphates (Jain et al.,2005). Specifically, glucose formed the main fraction of sugars in EPSfollowed by galactose, mannose and arabinose. The molecular weightof EPSwas determined as 94 kDa and 320 kDa. However, it has been ob-served that many bacterial EPS possess a 3-D structure (Dogsa et al.,2005). Different structures have been reported in microbes includinglarge net structures in hyaluronan (carbohydrate polymer), an EPSfrom Streptococcus equi (Gamini et al., 2002), fibrous structures fromBacillus sp. CP912 (Yun and Park, 2003), and elastic coiled structuresfrom Erwinia chrysanthemi spp. (Ding et al., 2003). EPS can act as anti-tumour and antiviral agents and can also be used in the cosmetic andfood industries (Sutherland, 1998). EPS are found to play an importantrole in the cell life of thraustochytrids. They may protect thraustochy-trids from desiccation, assist in cell adherence to the marine substrataand serve as an energy source during periods of starvation (Jain et al.,

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2005). Thraustochytrids may also afford protection against metal andtoxin contamination (Colaco et al., 2006).

9. Thraustochytrids and their potential

Most PUFAs (particularly DHA and EPA) presently used as foodsupplements and medicines are obtained from fish oil. These oilsare mostly accumulated in fish through bioaccumulation in the foodchain as fish feed on zooplanktons which have consumed microalgaecontaining PUFAs (Bell et al., 1986; Saito et al., 1995). However, fishoil is associated with heavy metal contamination and extractioninvolves chemical processes such as fractional distillation and ureaconcentration which alter the fatty acids chain by oxidative degrada-tion. Additionally, waste products in the form of urea are costly to dis-pose of (Zuta et al., 2003). The fish oil in food supplements is alsolinked to a distinctive smell, unpleasant taste and poor oxidativestrength. This has encouraged researchers to look for alternative, con-taminant free, eco-friendly and high quality DHA producing sources(Spolaore et al., 2006). Thraustochytrids are known to be producersof high levels of ω-3 PUFAs, DHA and EPA (Raghukumar, 2002). Itwould also be of interest to investigate the tolerant behaviour ofthraustochytrids towards metals such as iron, manganese and lead(Colaco et al., 2006; Goldstein, 1973). In addition, thraustochytridshave been shown to be potential sources of novel drugs and second-ary metabolites (Raghukumar, 2008).

10. Limitations of marine exploration

Human activity is polluting the marine ecosystem hindering theefforts of scientists to extract information or products in their pureform from the marine world. Limitations imposed on scientists bythe consequences of human activity and contamination include:

1. Decreased biodiversity due to disturbed ecosystems.2. Heavy metal contamination of fish oils.3. A relatively small fraction of marine bio-reserves which have been

declared, limiting comparative studies with analogous sites.4. Global environment change such as the harmful effect of global

warming has affected the oceanic ecology leading to a decline inthe growth of some marine microbes potentially impacting accu-mulation of omega oils.

11. Conclusions and future prospects

Researchers have a great opportunity to discover and exploit manyas yet unidentified marine microbes capable of producing higher levelsof omega-3 fatty acids and other valuable products. One of the mainchallenges is developing optimum culture conditions for rapidly grow-ing marine microbes that produce high levels of DHA and EPA, as com-pared to the limited number of commercially useful species presentlyavailable in the international depository. Culture collections must beeffectively preserved and maintained in centralised locations as doneby ATCC so that a diversity of cultures can be extensively evaluated bya wide range of researchers. The appropriate selection of microbesand their identification, plus suitably controlled fermentation methodsare required for cost-effective production of omega-3 oils such asDHA. Currently the cost of fermentation of DHA and EPA is significantlyhigher than obtaining these fatty acids from fish oil. However, fermen-tation enables better control of EPA andDHA ratios, aswell as decreasedcontaminant levels. The cold environments and high pressure associat-ed with the psychrophilic and piezophilic microbes might also have aneffect on phospholipid synthesis including the highly desirable PUFAscontaining phospholipids. For example, phosphatidyl serine DHA iscurrently available from krill oil, but is expensive and involves harvest-ing krill in large numbers. Krill are the main food for whales and so areimportant in the early food chain. Thraustochytrids can produce

phospholipids and therefore have the potential to provide a ferment-able source of an oil similar to krill oil. The application of metabolic en-gineering techniques to improve thewild-type strain is in an early stageof development. In fact there is no full genomic sequence of a thrausto-chytrid yet published. Metabolic engineering could further improveyields of DHA, EPA and other compounds from thraustochytrids. Thecost of fermentation can be minimised using low cost carbon sourcessuch as glycerol and further work on the use of low cost carbon sourcesfor DHA production from these organisms is required.

In addition to this, the metal tolerance activity of thraustochytridsmay make them applicable for use in the bioremediation of some formsof metal, such as lead. This is a currently unexplored area in the applica-tion of thraustochytrids. The identification of currently undiscoveredstrains, the development of improved fermentation techniques andmet-abolic engineering approaches to improve lipid synthesis are key areas ofresearch for thraustochytrids. Thraustochytrids can be a useful source ofomega-3 fatty acids, carotenoids, enzymes and other valuable com-pounds. The study of thraustochytrids is still in an early stage and thereis much left to do in the biotechnology of these marine microorganisms.

NomenclatureDHA docosahexaenoic acidEPA eicosapentaenoic acidDPA Docosapentaenoic acidω omegaΔ deltaα alphaγ gammaβ betaDCW dry cell weightFISH fluorescence in-situ hybridization techniqueAFDD acriflavine direct detection methodrDNA ribosomal DNAER endoplasmic reticulumNADPH nicotinamide adenine dinucleotide phosphateAcetyl Co-A acetyl co-enzyme A(NH4)2SO4 ammonium sulphateMSG monosodium glutamateFAMEs fatty acid methyl estersCO2 carbon dioxideCMC carboxy-methylcelluloseEPS extracellular polysaccharidesATCC American Type Culture Collection

Note added in Proof

The authors propose to provide additional information thatstrengthens the biosynthetic pathway followed in marine microor-ganism such as Schizochytrium for the synthesis of lipids.

Besides the conventional fatty acid synthase (FAS) system in thraus-tochytrids for fatty acid synthesis, one distinct pathway named polyke-tide synthase (PKS) system had been observed in Schizochytrium sp.which does not require molecular oxygen in contrast to FAS system(Metz et al., 2001). PKS system still involves acetyl-CoA and malonyl-CoA being the essential building blocks however does not involve insitu reduction of the intermediates. The short chain fatty acids (C14:0and C16:0) synthesis in Schizochytrium sp. might have followed theFAS system but the long chain fatty acids was found to be synthesisedthrough PKS system (Ratledge, 2004; Fig. 2a) which involved the useof 8 enzymes; 3-ketoacyl synthase (KS), malonyl-CoA:ACP acyltransfer-ase (MAT), acyl carrier protein (ACP), 3-ketoacyl-ACP reductase (KR),acyltransferase (AT), chain length factor (CLF), enoyl reducatase (ER)and dehydrase or isomerase (DH). Unlike the use of several elongasesand desaturases enzymes in FAS pathway, PKS pathway exhibited

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dehydration and isomerization reactions involving fatty acyl intermedi-ates for carbon chain elongation. Though the identification of Δ4 fattyacid desaturase in Thraustochytrium sp. (Qiu et al., 2001) suggestedthat, the synthesis of docosahexaenoic acid (DHA) may be the result ofusual FAS system, however, no other desaturase enzyme has beenfound to ascertain thisfinding (Ratledge, 2004). Thus, thedefinitemech-anism involved, would take more time to unfold to give a clear pictureabout the fatty acid biosynthesis in thraustochytrids.

Acknowledgements

The authors are thankful to the Strategic Research Centre (SRC) forBiotechnology, Chemistry and System Biology, Deakin University,Australia for providing support to pursue research work.

Also, the authors thank Professor Colin Ratledge (University ofHull, UK) for his observations, which emphasized the relevance ofan alternative pathway i.e. PKS pathway in marine microbes foromega-3 oils synthesis.

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