Transgenic Biosynthesis of Polyunsaturated Fatty Acids: A Sustainable

Review pubs.acs.org/CR

Transgenic Biosynthesis of Polyunsaturated Fatty Acids: A Sustainable Biochemical Engineering Approach for Making Essential Fatty Acids in Plants and Animals Jingjing Jiao† and Yu Zhang*,‡ †

Chronic Disease Research Institute, Department of Nutrition and Food Hygiene, School of Public Health, Zhejiang University, Hangzhou 310058, China ‡ Department of Food Science and Nutrition, School of Biosystems Engineering and Food Science, Zhejiang University, Hangzhou 310058, China 6.2. Strategies for Enhancing the Production of PUFAs in Transgenic Organisms 7. Future Prospects Author Information Corresponding Author Notes Biographies Acknowledgments Abbreviations References

1. INTRODUCTION The dietary equilibrium of fatty acid composition is an important consideration for the daily fatty acid consumption of humans and provides considerable benefits to human nutrition.1 Different types of fatty acids found in lipid compounds have distinct functions for human health. However, the rapid development of the food industry, the diversification of food processing methodologies, and the impact of dietary habits may inevitably induce nutritional imbalances (e.g., imbalance of the daily consumption ratio of ω-6 to ω-3 fatty acids) and raise a series of health issues.2 Polyunsaturated fatty acids (PUFAs), including ω-6 and ω-3 fatty acids, are essential nutrients for humans; however, C18 PUFA cannot be synthesized de novo in mammals.3 Dietary PUFAs are important determinants of the structure and function of animal cells, and these nutrients are critical human nutrition sources for improving health and preventing diseases, especially cardiovascular diseases.4 The role of the ratio of ω-6 to ω-3 fatty acids and ω-3 PUFA in the prevention and control of disease has attracted wide attention and has recently become the focus of recent investigations. Moreover, the tissue content of ω-3 PUFA is highly related to brain and retinal development in humans, especially children. Excessive amounts of ω-6 PUFA and high ω-6/ω-3 ratios are highly associated with the pathogenesis of cardiovascular diseases,5 cancer,6 inflammatory disorders,7 and neural diseases,8 whereas an enhanced level of ω-3 PUFA and low ω-6/ω-3 ratios appear to induce suppressive effects on the incidence of various diseases.9 Besides, high levels of ω-3 PUFA and low dietary ratio of ω-6 to ω-3 PUFA are also related to the control and

CONTENTS 1. Introduction 2. Mapping PUFA Biosynthesis Pathways 2.1. Δ6-Desaturation Pathway 2.2. Δ8-Desaturation Pathway 2.3. Δ4-Desaturation Pathway 2.4. Sprecher Pathway 2.5. Polyketide Synthases Pathway 3. Lipidomic Analysis of PUFAs 3.1. Shotgun Lipidomics and PUFA Analysis 3.2. Targeted Lipidomics and PUFA Analysis 4. Sources of PUFA Biosynthesis Enzymes 4.1. Fatty Acid Desaturases 4.2. Fatty Acid Elongases 4.3. Polyketide Synthases 5. Heterologous Expression of Genes Encoding PUFA Biosynthesis Enzymes 5.1. Plant FAD Gene Family: Contributing to the Synthesis of Mono-, Di-, and Triunsaturated Fatty Acids 5.2. Human and Animal FAT/FADS Genes: Acting as ω-3/ω-6 and Δ5/Δ6 Desaturases for PUFA Biosynthesis 5.3. Fatty Acid Elongase Genes: Encoding the Elongation of the Carbon Chain Skeleton of PUFAs 5.4. Effect of Extraneous Factors on PUFA Biosynthesis 6. Enhanced Production of PUFAs via Transgenic Approaches 6.1. Biochemical Transgenic Engineering and Production Yields of PUFAs © XXXX American Chemical Society

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dx.doi.org/10.1021/cr300007p | Chem. Rev. XXXX, XXX, XXX−XXX

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Figure 1. Biosynthesis of PUFAs. The aerobic biosynthesis pathway is shown via different routes, including the Δ6, Δ8, Δ4, and “Sprecher” pathways. AA, arachidonic acid; ALA, α-linolenic acid; DGLA, dihomo-γ-linolenic acid; DHA, docosahexaenoic acid; DPA, docosapentaenoic acid; EDA, eicosadienoic acid; EPA, eicosapentaenoic acid; ETA, eicosatetraenoic acid; ETrA, eicosatrienoic acid; GLA, γ-linolenic acid; LA, linoleic acid; OA, oleic acid; OTA, octadecatetraenoic acid; SA, stearic acid; THA, tetracosahexaenoic acid; TPA, tetracosapentaenoic acid; Des, desaturase; Elo, elongase.

prevention of periodontal,10 nonalcoholic fatty liver,11 and Crohn’s diseases.12 Unfortunately, imbalances between ω-3 and ω-6 PUFA consumption are commonly found, and ω-3 PUFA deficiency has become a serious health problem, especially in the elderly, alcoholic patients, and low-weight newborns.13 In general, plants can synthesize a variety of unsaturated fatty acids, such as oleic acid (OA, C18:1, Δ9), linoleic acid (LA, C18:2, Δ9,12), and α-linolenic acid (ALA, C18:3, Δ9,12,15) via lipid metabolism.14 Most mammals have the capacity to synthesize PUFAs from precursor fatty acids derived from the diet, such as LA and ALA.15 However, the synthetic efficiency of ω-3 PUFAs in humans is low, especially for the conversion from ALA to ω-3 long-chain polyunsaturated fatty acids (LCPUFAs). Importantly, these poor ω-3 LC-PUFA synthesis efficiencies make it difficult to satisfy the daily needs of ω-3 LCPUFA nutrients for humans. Therefore, daily supplements of ω-3 PUFAs are needed for maintaining human health. Currently the main source of ω-3 LC-PUFAs for humans has been deepwater fish oils, which are accumulated via the intake of ω-3 LC-PUFA-rich microalgae by fish (mackerel, herring, salmon, sardines, etc.) that do not synthesize these ω-3 LCPUFAs by themselves. Changing the fatty acid profile by elevating the synthesis of ω-3 PUFA (via modifying the original genotype and improving the expression of target genes in vivo) is an effective way of rebalancing the contents of ω-3 and ω-6 fatty acids. Significant advances have been made in developing higher plants or animals that biosynthesize LC-PUFAs via transgenic technol-

ogies.16 It has become increasingly important to develop new LC-PUFA sources because excessive exploitation has made an impact in the reduction of marine fish products. Moreover, unexpected environmental pollution has led to undesirable heavy metal residues in fish oils. Biosynthesis and transgenic approaches have the potential to provide safe, affordable, and renewable sources of nutritional LC-PUFAs, such as arachidonic acid (AA, C20:4, Δ5, 8, 11, 14), eicosapentaenoic acid (EPA, C20:5, Δ5, 8, 11, 14, 17), and docosahexaenoic acid (DHA, C22:6, Δ4, 7, 10, 13, 16, 19), and these could make up for decreasing and unsustainable supplies of LC-PUFAs from fish. Thus, by using transgenic technology, ω-3 and ω-6 PUFA consumption can eventually be kept in balance in humans. Previous work has indicated that some lower species, such as nematodes, fungi, and algae, have demonstrated a promising capacity for synthesizing LC-PUFAs.17 Thus, many researchers have devoted resources to cultivate new or improved species as sources for the synthesis of LC-PUFAs and for further developing transgenic plants and animals via molecular cloning. Also, scientists focused on simultaneous, rapid, and highthroughput analysis of saturated fatty acids, monounsaturated fatty acids, PUFAs, LC-PUFAs, and their in vivo metabolites, which is an important branch of lipidomics. Lipidomics is the emerging field of systems-level analysis of lipids and factors that interact with lipids.18 Although important, the study of lipids has been hampered by analytical limitations.19 Major advances in lipidomic technology come from combining powerful separation techniques (e.g., high-performance liquid chromaB


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Figure 2. Processive biosynthesis of PUFAs via the PKS pathway.

followed by Δ6-specific elongation to produce dihomo-γlinolenic acid (DGLA, C20:3, Δ8, 11, 14) and eicosatetraenoic acid (ETA, C20:4, Δ8, 11, 14, 17), respectively. Finally, the fatty acids AA and EPA are generated after Δ5-desaturation, respectively.15,23 Many plants have the ω-3 desaturase ability of transforming ω-6 fatty acids (such as LA) into ω-3 fatty acids (such as ALA) via dehydrogenization. Unfortunately, plants do not encode the capability to do Δ6-elongation and Δ5- or Δ6desaturation; therefore, these pathways need to be obtained from other species via transgenosis.

tography (HPLC)) with sophisticated detection methods (e.g., mass spectrometry) and make this field a promising area of biomedical research, with a variety of applications in the development of transgenic biosynthesis and biomarker analysis.20 In addition, there is a lot of interest in genes encoding desaturation and elongation enzymes of the PUFA biosynthetic pathway, and considerable efforts have been expended to identify these genes. Similar to the well-studied engineering of transgenic oilseeds, research progress on the biosynthesis of PUFAs in transgenic animals has attracted wide attention and has led to the development of FAT-1 transgenic mice,17a pigs,21 marine fish,22 etc. This review highlights the key features and recent developments in PUFA biosynthesis pathways, in lipidomic analysis of PUFAs and their metabolites, in species sources of biosynthetic enzymes, and in heterologous expression of transgenes. Furthermore, this review presents insights and strategies for the sustainable biochemical engineering of PUFA in multifarious transgenic organisms.

2.2. Δ8-Desaturation Pathway


This alternative pathway is usually present in protozoan, and similar to the Δ6-desaturation pathway, the Δ8-desaturation pathway is composed of multiple desaturation and elongation steps, including continuous Δ9-elongation, Δ8-desaturation, and Δ5-desaturation. When LA is used as the substrate in this pathway, eicosadienoic acid (EDA, C20:2, Δ11, 14) is synthesized first, followed by the formation of DGLA. AA is ultimately generated after Δ5-desaturation. Alternatively, eicosatrienoic acid (ETrA, C20:3, Δ11, 14, 17) is initially synthesized when ALA is used as the substrate, and ETA followed by EPA are then formed via Δ8- and Δ5-desaturation, respectively.15,24 The role of this alternative route, in terms of physiological significance in human health, is still unclear. However, this metabolic approach, especially for the synthesis of EPA, could be used when the conventional Δ6-desaturation route is genetically or pathologically impaired.

This is the major and conventional aerobic pathway widely present in most of eukaryotic organisms. The pathway initiates with Δ6-desaturation and results in the synthesis of γ-linolenic acid (GLA, C18:3, Δ6, 9, 12) and octadecatetraenoic acid (OTA, C18:4, Δ6, 9, 12, 15) when LA and ALA act as substrates, respectively. The Δ6-desaturation reaction is

The existence of this Δ4-specific pathway was discovered via the isolation of a Δ4-desaturase from the marine protist Thraustochytrium spp. and the freshwater species Euglena.17b,25 This pathway involves a Δ5-elongation effect on EPA to generate docosapentaenoic acid (DPA, C22:5, Δ7, 10, 13, 16,

2. MAPPING PUFA BIOSYNTHESIS PATHWAYS Several alternative pathways have been found for the biosynthesis of PUFAs, especially those that allow for the generation of EPA and DHA. The genes encoding various critical enzymatic reactions have also been identified, including desaturase and elongase, both of which contribute to different biosynthesis pathways of PUFAs (Figure 1).


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19), which is then desaturated by Δ4-desaturation to yield DHA.

of shotgun lipidomics is expected to increase sensitivity, mass accuracy, and integrated array analysis of the metabolic networks involved in neutral lipid metabolism. Furthermore, it seems that the increased use of stable isotope methods in both intact cells and organs will continue to provide rapid improvements in the understanding of the alterations in neutral lipid metabolic networks that participate in cellular bioenergetics, membrane synthesis, and cellular signaling in health and disease.41 To comprehensively understand the efficiency of each desaturase and elongase during PUFA biosynthesis, the simultaneous determination of participating fatty acids and their isomers is requested. Fatty acids and their isomers are traditionally analyzed by gas chromatography (GC) and HPLC.42 However, the need for derivatization, limited resolution, the time-consuming sample pretreatment steps and large amount of analytical tasks of samples are some of the difficulties associated with those methods.43 The MS-based shotgun lipidomics methods for simultaneous determination of PUFAs with different unsaturated degrees and identification of the location of double bonds in PUFAs have rapidly been developed. For example, the method based on the fragmentation pattern analysis after high-energy collision-induced dissociation (CID) is one of previously developed MS methods for the direct analysis of PUFAs and fatty acid isomers.44 Hsu and Turk45 have investigated low-energy CID for identification of the double bond location in fatty acids. Ozonolysis is the other developed method that is associated with MS analysis of fatty acids.33,46 Although current shotgun lipidomics technology is capable of simultaneously quantifying hundreds of molecular species covering >20 major lipid classes from lipid extracts of biological samples, the targeted analyte PUFAs and their isomers are difficultly identified from numerous lipid analytes.47 Sometimes similar MS fragmentation patterns between PUFAs and other lipid analytes will interfere with the identification task. To cope with this problem, technologies of multidimensional MS-based shotgun lipidomics analysis,43,48 matrixassisted laser desorption and ionization time-of-flight (MALDI-TOF) MS,49 and quadrupole time-of-flight (QTOF) MS50 have been recently developed. However, the instruments of the above methods are relatively expensive compared with the classically used GC or GC-MS instruments. Therefore, identification and quantification of PUFAs and fatty acid isomers still remain challenging in shotgun lipidomics.

2.4. Sprecher Pathway

Compared to the Δ4-desaturation pathway, the Sprecher Δ4independent pathway is a more complicated biosynthesis route for the production of DHA. Previous studies have demonstrated that this is the predominant route used in mammals.26 As an alternative pathway to produce DHA, the Sprecher route is independent of Δ4-desaturation but involves consecutive Δ5and Δ7-elongation cycles to produce tetracosapentaenoic acid (TPA, C24:5, Δ7, 10, 13, 16, 19), followed by Δ6-desaturation and one cycle of C2-shorteninig via β-oxidation in the peroxisome to yield DHA.27 Recent evidence has been reported by several groups, indicating that C24 Δ6-desaturase is the same enzyme as C18 Δ6-desaturase, which is responsible for the synthesis of GLA and OTA.28 In addition, the Sprecher pathway is also used by nonmammalian vertebrates such as fish; however, it is still unclear whether all chordates use this pathway to yield DHA.29 2.5. Polyketide Synthases Pathway

Compared to the aerobic pathways that use desaturase and elongase, the polyketide synthases (PKS) system is an anaerobic biosynthetic pathway that has been identified for producing PUFAs in both prokaryotic and eukaryotic marine organisms.30 Such biosynthesis does not require the multiple desaturase and elongase enzymes as in the aerobic biosynthesis pathways but instead uses a PKS-like gene cluster to synthesize PUFAs.31 In detail, the PKS pathway uses acyl carrier protein (ACP) as a covalent attachment for fatty acid chain synthesis, proceeding with reiterative cycles. The full cycle of biosynthesis includes condensation of an acyl-ACP and a malonyl-ACP to generate a ketoacyl-ACP, ketoreduction to convert ketoacylACP to hydroxyacyl-ACP, dehydration to remove a water molecule from hydroxyacyl-ACP resulting in unsaturated enoylACP, and reduction of enoyl-ACP to a saturated acyl chain (Figure 2).25 However, unlike fatty acid synthesis, PKS often omits steps of the full cycle such as dehydration and reduction. Thus, the products of the PKS pathway are highly varied in structure and often contain keto and hydroxyl groups and double bonds.32

3. LIPIDOMIC ANALYSIS OF PUFAS Lipidomics, the systematic study of lipids and their function in biological systems, has emerged as an active area of interest. Although it is often considered as a branch of metabolomics, it is unique in terms of the analytical and bioinformatic techniques employed.33 In recent years, review publications focused on lipidomics in the context of their structural roles in biological membranes,20 exosomes,34 and cytoskeletal dynamics,35 or their functional roles in pain modulation,36 brain signaling,37 and toxicology.38 Mass spectrometry or tandem mass spectrometry (MS or MS/MS)-based separation methods are becoming increasingly important in biomolecule analysis and are mostly pronounced in the emerging field of lipidomics.33,39 The lipidomic analysis of PUFA profiles plays an important role in the evaluation of desaturase or elongase efficiencies and biosynthesis production yields.

3.2. Targeted Lipidomics and PUFA Analysis

Lipidomics has been widely acknowledged as a tool of the full characterization of lipid molecular species and of their biological roles with respect to expression of proteins involved in lipid metabolism and function, including gene regulation.51 Although shotgun lipidomics is used for the large-scale analysis of as many as possible lipids, targeted lipidomics aims to focus on one specific lipid pathway.52 The identification and quantification of eicosanoids, a large family of AA oxidation products that contain 20 carbon atoms, are mostly studied via targeted lipidomics analysis.53 GC-MS combined with electroncapture negative chemical ionization (ECNCI) was commonly used in the past eicosanoid analyses.54 However, extremely tedious derivatization procedures are compulsory, making this method very time-consuming for the analysis of large sample amounts. Subsequently, the analytical methods of endogenous eicosanoids were significantly expanded by discoveries of electrospray ionization (ESI),55 atmospheric-pressure chemical ionization (APCI),56 and electron-capture atmospheric-pres-

3.1. Shotgun Lipidomics and PUFA Analysis

Shotgun lipidomics is a rapidly developing technology, which identifies and quantifies individual lipid molecular species directly from lipid extracts of biological samples.40 The power D


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Figure 3. Representative sources and species of both aerobic and anaerobic enzymes for the biosynthesis of PUFAs. Light circle, aerobic desaturases (Des's); medium circle, anaerobic polyketide synthases (PKS); dark circle, aerobic elongases (Elo's), respectively. Des: Ac, Acanthamoeba castellanii;62,63 At, Arabidopsis thaliana;64 Cb, Caenorhabditis briggsae;65 Cc, Coprinus cinereus;66 Ce, Caenorhabditis elegans;67 Ci, Ctenopharyngodon idella;68 Cr, Chlamydomonas reinhardtii;69 Dl, Dicentrarchus labrax L.;70 Eg, Euglena gracilis;17b Gm, Gadus morhua L.;71 Lc, Lates calcarifer;72 Ma, Mortierella alpina;73 Pl, Pavlova lutheri;74 Po, Portulaca oleracea L.;75 Pt, Phaeodactylum tricornutum;76 Ss, Salmo salar.77 Elo: Ig, Isochrysis galbana;78 Mp, Marchantia polymorpha L.;79 Pi, Parietochloris incise;80 Pp, Physcomitrella patens;81 Ps, Pavlova salina;82 Ta, Thraustochytrium aureum.83 PKS: S, Schizochytrium sp.;30 Sp, Shewanella putrefaciens.84

sure chemical ionization (ECAPCI).57 Liquid chromatography (LC) using reverse-phase solvents with microbore columns or normal-phase solvents with chiral columns is now able to separate complex mixtures of eicosanoid regioisomers and enantiomers.53,58 Advanced tandem MS technology can provide the highest sensitivity and specificity as well as the separation of the isomers that LC is unable to resolve. The sample preparation for LC-MS analysis of eicosanoids usually consists of a single solid-phase extraction (SPE) or liquid−liquid extraction procedure with or without one-step derivatization, which saves time and labor and decreases loss of analytes.53 Compared to shotgun lipidomics analysis, the targeted lipidomics method usually refers to one of the lipid classes and eliminates the interference from other coanalytes. Most of the targeted lipidomics studies focus on the identification and quantification of eicosanoids59 and endocannabinoids.52,60 Few studies61 have used targeted lipidomics to analyze PUFAs, free fatty acids, and their derivatives so far. For the analysis of PUFAs, the targeted lipidomics method is capable of analyzing saturated fatty acids, monounsaturated fatty acids, and PUFAs in one of the specific biosynthesis pathways. However, it is difficult to quantify all of the fatty acid analytes present in all of the PUFA biosynthesis pathways simultaneously using this technology.

4. SOURCES OF PUFA BIOSYNTHESIS ENZYMES 4.1. Fatty Acid Desaturases

Most of the desaturation enzymes participating in PUFA biosynthesis have been cloned and characterized from various organisms, including alga, fungi, mosses, plants, and mammals (Figure 3). For instance, fungal desaturase mutants with unique enzyme systems are used not only for the regulation and overproduction of valuable PUFAs but also for the elucidation of fungal lipogenesis.85 On the basis of the presence of different catalysates, the fatty acid desaturases (FADs) could be divided into three groups: (i) the acyl-coenzyme A (acyl-CoA) desaturase that catalyzes CoA-bound fatty acids and that is widely present in mammals, yeast, and fungi; (ii) the acyl−acyl carrier protein (acyl-ACP) desaturase that catalyzes ACPbound fatty acids for the generation of the first double bond and that is present in plant plastids; and (iii) the acyl-lipid desaturase that catalyzes lipid-bound fatty acids for most of desaturation steps and that exists in various plants and blue alga.86 On the basis of regional selectivity, FADs can also be separated into two groups: the “methyl-end” desaturase (e.g., ω-3 and ω-6 desaturases), indicating the next double bond between the existing one and the methyl end of the fatty acid chain, and the “front-end” desaturases (e.g., Δ5, Δ6, Δ8, Δ9, Δ12, and Δ15 desaturases), inserting a new double bond between an existing one and the carboxyl end of the acyl group in a methylene-interrupted style.15,87 These “front-end” E


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desaturases are all members of the cytochrome b5 fusion desaturase superfamily, because they contain an N-terminal domain that is orthologous to the microsomal cytochrome b5 and serves as the electron donor for desaturation.88 Recently, FADs (mainly Δ6- and Δ12-desaturases) were isolated from some new microorganisms, plants, and animals. In detail, Δ6desaturases were successfully isolated and characterized from blackcurrant (Ribes nigrum L.),89 black seabream (Acanthopagrus schlegeli),90 borage (Borago of f icinalis),91 chicken,92 entomopathogenic fungus (Conidiobolus obscurus),93 European sea bass (Dicentrarchus labrax L.),70 grass carp (Ctenopharyngodon idella),68 and unicellular marine microalga (Nannochloropsis oculata).94 Δ12-Desaturases were molecularly cloned and characterized from the Antarctic microalga (Chlorella vulgaris NJ-7),95 Chinese dove tree (Davidia involucrata Baill),96 marine alga (Pinguiochrysis pyriformis),97 peanut (Arachis hypogaea L.),98 and white-rot fungus (Ceriporiopsis subvermispora).99 Identification and characterization of some other FADs (e.g., Δ4-desaturases) were also studied.100 4.2. Fatty Acid Elongases

The identification and cloning of the main enzymes catalyzing the Δ5, Δ6, and Δ9 elongation pathways of plants, fungi, and microalga have been reported (Figure 3). Biochemical evidence revealed that four principal successive steps of each C2 elongation are performed by individual proteins.101 The elongation process starts with the condensation of acyl-CoA and malonyl-CoA to form 3-ketoacyl-CoA. Then, a reduction reaction occurs that converts 3-ketoacyl-CoA into 3-hydroxyacyl-CoA in the presence of nicotinamide adenine dinucleotide phosphate (NADPH). 3-Hydroxyacyl-CoA is subsequently dehydrated in the third step, resulting in enoyl-CoA, which is reduced again by enoyl-CoA reductase in the final step to complete the elongation cycle and generate an extended acylchain (Figure 4).102 The four-step elongation process requires the participation of related enzymes. In the first step, the 3ketoacyl-CoA synthases (KCS), identified as the enzymes responsible for the condensation step, can be divided into two groups: (i) the plant fatty acid elongase (FAE)-like enzymes from higher plants (e.g., Arabidopsis thaliana) involved in the biosynthesis of saturated and monounsaturated fatty acids with C18−22 chain length,103 and (ii) the elongase (ELO)-type class of elongases involved in PUFA biosynthesis.104 However, the elongating activity of a FAE-like KCS from the protozoan Perkinsus marinus (not a member of the ELO family) has recently been demonstrated to be involved in PUFA biosynthesis.104 In the remaining steps, previous studies have reported that the three required enzymes, including 3-ketoacyl-CoA reductase (KCR), 3-hydroxyacyl-CoA dehydratase, and trans2,3-enoyl-CoA reductase, were successfully isolated and functionally characterized from cotton Gossypium hirsutum L. fibers,105 yeast Saccharomyces cerevisiae,106 and Gram negative spirochete Treponema denticola,107 respectively.

Figure 4. Intermediate enzymatic reactions involved in acyl-CoA elongation. The blue, red, yellow, and green balls indicate coenzyme A (CoA), synthases, reductases, and dehydrases, respectively. HCD, 3hydroxyacyl-CoA dehydratase; KCR, 3-ketoacyl-CoA reductase; KCS, 3-ketoacyl-CoA synthase; TER, trans-2,3-enoyl-CoA reductase.

of phosphopantetheine transferase and identified sequences related to fatty acid synthases (FASs), in the form of enoyl reductase and dehydratases.31b,84,109

5. HETEROLOGOUS EXPRESSION OF GENES ENCODING PUFA BIOSYNTHESIS ENZYMES 5.1. Plant FAD Gene Family: Contributing to the Synthesis of Mono-, Di-, and Triunsaturated Fatty Acids

The small crucifer, Arabidopsis thaliana L., is appropriate for the map-based cloning of FAD genes because it has a small nuclear genome that is almost devoid of interspersed, highly repetitive DNA. The FAD2 and FAD6 encoding ω-6 fatty acid desaturases were first isolated from Arabidopsis and characterized.110 FAD2, designated as the microsomal oleate desaturase, is located in the endoplasmic reticulum (ER). FAD2 uses phospholipids as acyl substrates and introduces nicotinamide adenine dinucleotide (NADH), NADH-cytochrome b5 reductase, and cytochrome b5 as electron donors. In contrast, FAD6, designated as the plastidial oleate desaturase, is located in the chloroplast. FAD6 uses primarily glycolipids as acyl carriers and introduces NAD(P)H, ferredoxin-NAD(P) reductase, and ferredoxin as electron donors. Moreover, different FAD gene repeats, especially FAD2, have different

4.3. Polyketide Synthases

Previously, several PKS gene clusters derived from marine bacteria Shewanella and Vibriomarinus resulted in the accumulation of EPA and DHA when expressed in E. coli or Synechococcus.84,108 Further analysis of some of the Shewanella open-reading frames (ORFs) revealed several regions with similarity to synthesis domains of PKS, including 3-ketoacyl synthases, 3-ketoacyl reductase, ACP, chain length factor, and acyl transferases. In addition, other researchers working with Shewanella have reported accessory enzyme activity in the form F


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Figure 5. Biosynthesis and conversion of ω-6 PUFA into ω-3 PUFA in transgenic animals and plants using the FAT-1 transgene.

a milestone in the era of postgenomic biology for multicellular organisms. The entire C. elegans genome yielded seven ORFs with similarity to fatty acid desaturases from other species. Functional characterization has defined the precise enzymatic roles of each of these ORFs and has provided a comprehensive description of the C. elegans genes (FAT) involved in the PUFA biosynthetic pathway.123 The most important genes consist of FAT-1 and FAT-2. The FAT-1 gene of C. elegans encodes a ω-3 fatty acid desaturase that converts ω-6 into ω-3 fatty acids, which are absent in most animals, including mammals.4a,17a FAT-1 was shown to exert its specificity for both C18 and C20 unsaturated substrates,124 although the very small quantities of ALA in C. elegans may indicate a preference for C20 fatty acids.125 The dual C18/C20 substrate preference of FAT-1 provides two alternative pathways for the order of desaturation required for EPA synthesis.123 On the basis of this functionality of FAT-1, Kang et al.17a generated a transgenic mouse that is capable of converting ω-6 into ω-3 fatty acids using the FAT-1 gene to reduce the ratio of ω-6 to ω-3 from 20:1−50:1 to almost 1:1 (Figure 5). The FAT-1 transgenic mice are now being widely used as a new and emerging tool for studying the benefits of ω-3 fatty acids and the molecular mechanisms of their functionality.126 In addition, an sFAT-1 gene synthesized using modified and optimized codons from another type of nematode C. briggsae also encoded the capacity to produce ω-3 PUFAs (especially DHA and DPA) in transgenic mice.65 FAT-2 displays a high degree of sequence identity (72%) to FAT-1 and encodes a Δ12/ω-6 fatty acid desaturase that desaturates both 16:1 and 18:1 substrates.127 Therefore, the desaturation reaction of OA is responsible for the synthesis of LA. Moreover, Kong et al.128 revealed that the FAT-2 gene isolated from C. elegans could be successfully integrated into the genome of Chinese hamster ovary cells and transgenically expressed, resulting in the accumulation of LA. FAT-3 and FAT-4 carry out “front-end” desaturation and, like Δ5- and Δ6desaturases from other systems,129 differ from the other C. elegans desaturases in the presence of an N-terminal cytochrome b5 domain.123 FAT-3, encoding the Δ6 fatty acid

expression patterns as well as functionality in the same plants. For example, other than other FAD2 gene repeats, FAD2−1, FAD2−2, and FAD2−4 isolated from turnip (Brassica rapa L.),111 olives (Olea europaea L.),112 and upland cottons (Gossypium hirsutum L.),113 respectively, are responsible for encoding a functional Δ12-desaturase and converting OA into LA. Therefore, the orientation and isolation of functional FAD2 gene repeats play an important role in the improvement of LA contents in recombinant eukaryotes. A recent study on the generation of FAD2 transgenic mice showed that these mice exhibit an 87% and a 9% increase of AA contents in muscle and liver, respectively, indicating an active conversion of OA into LA.114 The FAD3, FAD7, and FAD8 genes encoding ω-3 fatty acid desaturases were also first identified in Arabidopsis.115 These three genes, designated as linoleate desaturases, are responsible for the conversion of LA into ALA. The FAD3 gene is located in ER membrane in Arabidopsis, and silencing this gene by small interfering RNA (siRNA) can induce the low ALA of FAD3 mutant phenotype.116 Recently, Zhang et al.117 reported that the FAD3 gene was highly expressed in leaves, roots, and stems of cold-sensitive lima bean (Phaseolus lunatus L.) Although the localization of the FAD7 gene in the chloroplast has long been established, it was only recently reported in a molecular identification study that FAD7 is preferentially localized in the thylakoid membranes beside the chloroplast envelope.118 As a plastidial isoenzyme of FAD7, the regulation of the post-translational stability of FAD8 provides an important regulatory mechanism in response to temperature, mediating a decrease in ALA levels at elevated temperatures.119 Besides high expression of genes in Arabidopsis, FAD7 and FAD8 genes encoding plastidial ω-3 fatty acid desaturases, which respond to temperature and pathogen stress, were recently characterized from purslane (Portulaca oleracea L.)120 and soybean [Glycine max (L.) Merr.].121 5.2. Human and Animal FAT/FADS Genes: Acting as ω-3/ω-6 and Δ5/Δ6 Desaturases for PUFA Biosynthesis

The successful completion of the nematode Caenorhabditis elegans genome sequencing program122 has been recognized as G


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biosynthesize C20/22 PUFAs.141 ELOVL4 was first discovered as a target gene in patients with eye diseases, including Stargardtlike and autosomal dominant macular dystrophy. The high expression of ELOVL4 in the human retina reveals that it might be involved in the elongation steps required for DHA synthesis.142 However, high levels of DHA cannot protect ELOVL4 transgenic mice against retinal degradation.143 A study on retinal fatty acids of diabetic animals showed that a decrease in C20+ PUFA is highly correlated to a reduction in ELOVL4 expression.144 In addition, salmon ELOVL4 effectively converted C20/22 PUFAs to elongated polyenoic products up to C36. Tissue distribution showed that salmon ELOVL4 mRNA transcripts are rich in eye, brain, and testes, indicating that these tissues are important metabolic sites for the biosynthesis of LCPUFAs.145 ELOVL5 encodes the elongation of C18−C20 PUFAs but does not appear to have the capacity to elongate PUFA substrates beyond C22.146 Until recently, most of the characterized fish fatty acyl elongases were considered to be ELOVL5-like, as they prefer C18/20 PUFA substrates.147 Studies using liver microsomal protein from wild-type and knockout mice demonstrated that the elongation of GLA to DGLA requires ELOVL5 activity. Moreover, ELOVL5-induced and endogenously synthesized PUFAs are key regulators of sterol regulatory element binding protein (SREBP)-1c activation and fatty acid synthesis in the livers of mice.148 Peroxisomal βoxidation is also involved in C22 PUFA synthesis, where elongation, presumably by ELOVL2 and/or ELOVL5, in the ER is followed by one round of chain-shortening in peroxisomes to produce the end products (e.g., DHA) destined for esterification in the ER.102,149 Because the ELOVL enzymes are proposed to be key enzymes in lipid metabolism, the diversity in transcriptional regulation of ELOVL2, ELOVL4, and ELOVL5 genes may provide some hints as to the physiological functions of downstream PUFA products in mammals.

desaturase, is responsible for the synthesis of GLA from LA or OTA from ALA, while FAT-4 encoding the Δ5 fatty acid desaturase is responsible for the final desaturation step in the synthesis of AA from DGLA or EPA from ETA. The similarity between the amino acid sequences of FAT-3 and FAT-4 is 66%. Mutational studies have demonstrated that C. elegans FAT-3 mutants lacking Δ6-desaturase activity fail to produce C20+ PUFAs and cause behavioral and developmental defects.130 Also, previous work demonstrated that a trans fat diet could cause decreased brood size and shortened lifespan in C. elegans Δ6-desaturase mutant FAT-3.131 Characterization of the ORFs in yeast via heterologous expression indicated that FAT-6 and FAT-7 are stearoyl Δ9-desaturases that also desaturate PA. However, FAT-5 displays a strict substrate preference for PA and shows minor activity toward stearate.132 FAT-6 and FAT-7 are 92% identical at the amino acid sequence level and 78−80% identical with FAT-5. In animals and humans, the Δ5- and Δ6-desaturases are the key enzymes introducing de novo unsaturations in the carbon chain of precursors leading to the biosynthesis of LC-PUFAs. Previous studies described the mammalian genomic structure of the fatty acid desaturase (FADS) cluster including FADS1 and FADS2 genes encoding the activities of Δ5- and Δ6desaturases, respectively.133 A third gene, named FADS3, was identified, indicating 62% and 70% nucleotide sequence homology with FADS1 and FADS2, respectively. Further studies showed a significant correlation between FADS3 polymorphism and lipid metabolism biomarkers such as PUFAs, high-density- or low-density-lipoprotein cholesterol, and triglyceride levels.134 Compared to confirmed functionalities of FADS genes in mammals, bifunctional activities of FAD genes were found in vertebrates. Like mammals, Atlantic salmon possesses separate genes for Δ5 and Δ6 FADS.77,135 Distinct Δ6 FAD cDNAs have been isolated from all fish species studied to date including freshwater and marine species.71,136 A recent study found that a FAD gene with Δ5 activity isolated from zebrafish is the bifunctional Δ6/Δ5 FAD.137 Further study showed that FAD1 and FAD2 genes isolated from the herbivorous marine teleost fish (Siganus canaliculatus) possess bifunctional Δ6/Δ5 and Δ4/Δ5 activities, respectively. This is a unique report of a vertebrate FAD with Δ4 activity and suggests that there is potentially more than one possible pathway for the biosynthesis of DHA in vertebrates.138

5.4. Effect of Extraneous Factors on PUFA Biosynthesis

Compared to the activities of desaturases and elongases, extraneous factors (e.g., dietary PUFA supplementation, chemicals, biochemicals, drugs, oxidative stress, etc.) are also crucial for the step-by-step PUFA synthesis in both mammals and plants. First, dietary fatty acids, especially LA and ALA, are promising candidate sources for synthesis in mammals because mammals are incapable of synthesizing ω-6 and ω-3 PUFAs de novo and are entirely dependent on dietary sources to procure and maintain adequate levels of these fatty acids in the peripheral and central tissues. Many studies have revealed that low dietary PUFA supplementation upregulates the expression of various desaturase and elongase genes. Increasing levels of dietary linseed oil lowers deposition of DHA, EPA, and AA in all tissues of female zebrafish.150 The expression of both Δ6and Δ5-desaturase genes is generally higher in Atlantic salmon fed with vegetable oil-substituted diets compared to the salmon fed with fish oil, particularly in the seawater phase.151 Similarly, the expression of mRNA for Δ9-desaturase and SREBP-1c genes decreases in mammals fed with ω-3 PUFA-enriched fish oil compared to the animals fed with soybean oil.152 Dietary fish oil supplements could increase tissue ω-3 PUFA composition and expression of Δ6-desaturase and elongase 2 in Jade Tiger hybrid abalone.153 Dietary fish oils and water salinity were highly related to the fatty acid composition and expression of FADS6-like and ELOVL5-like genes of red sea bream Pagrus major.154 Previous work has also demonstrated

5.3. Fatty Acid Elongase Genes: Encoding the Elongation of the Carbon Chain Skeleton of PUFAs

The genes involved in the elongation of PUFAs were investigated in both mammals and plants. To date, six genes named ELOVL1−6 have been identified and shown to conduct the condensation reaction during the elongation cycle. Among these genes, ELOVL2, ELOVL4, and ELOVL5 have functional roles in PUFA synthesis. ELOVL2 was first described in 2000, although the functional characterization of this gene was not clear at that time.139 Subsequent studies have shown that both mouse and human ELOVL are capable of elongating AA, EPA, docosatetraenoic acid (DTA, C22:4, Δ7, 10, 13, 16), and DPA in transfected yeast and mammalian HEK293 cells, whereas no activity was found for saturated or monounsaturated fatty acids.140 However, mouse ELOVL2, but not human ELOVL2, was able to elongate GLA to some degree, indicating a minor divergence between the species. The presence of ELOVL2 in Atlantic salmon also explains the ability of this species to H


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Figure 6. Production of PUFAs and yields of AA, EPA, and DHA via transgenes in different host organisms. At, Arabidopsis thaliana; Ce, Caenorhabditis elegans; Dr, Danio rerio; Eg, Euglena gracilis; Ig, Isochrysis galbana; Ma, Mortierella alpina; Mp, Marchantia polymorpha L; Nt, Nicotiana tabacum; Pp, Pichia pastoris; Pt, Phaeodactylum tricornutum; Ps, Pavlova salina; Om, Oncorhynchus masou; Des, desaturase; Elo, elongase.

that the deprivation of dietary ω-3 PUFA increases the coefficients of conversion of circulating ALA to DHA in the rat liver, but not brain, in the presence of upregulated mRNA and increased activity levels of Δ5- and Δ6-desaturases and elongases 2 and 5.155 Chemicals, biochemicals, and drugs are also considered as important foreign factors. Different carbon sources could also affect the biosynthesis of PUFAs and gene expression of desaturases on the molecular and biochemical levels.156 In addition, the use of oxyacetamide, chloroacetanilide, pyroxasulfone, and other herbicides of Herbicide Resistance Action Committee groups K3 and N could inhibit fatty acid elongases during plant cultivation.157 Furthermore, the stress induced by chemical contamination residues (e.g., cadmium) provokes an inhibition of triunsaturated fatty acids and reduces the desaturation process.158 In mammalian studies, ethanol was shown to strongly inhibit the AA synthesis in hepatocytes from spontaneously hypertensive rats, likely by an inhibition of Δ5 desaturation.159 Benzoxazinones and indolediones were evaluated as long-chain fatty acid ELOVL6 inhibitors.160 On the basis of the reduced microsomal Δ6desaturase activity, soybean protein exerts its inhibitory effect on the linoleate desaturation index in liver microsomal lipids of rats.161 Furthermore, some drugs (e.g., atypical antipsychotic medications),162 hormones (e.g., testosterone),163 and even the gender of animals164 have been shown to modify the fatty acid synthesis patterns and augment PUFA biosynthesis in animals or cultured cells via the modulation of Δ5- and/or Δ6desaturase activities. Hitherto, few studies have focused on developing a mechanistic understanding of the impact of desaturase/elongase activities on relative gene expression during PUFA biosynthesis. Furthermore, it is still unclear whether the modulation of PUFA synthesis by the above extraneous factors can be used in transgenic animals and plants.

6. ENHANCED PRODUCTION OF PUFAS VIA TRANSGENIC APPROACHES 6.1. Biochemical Transgenic Engineering and Production Yields of PUFAs

In the past decade, great progress has been made in engineering PUFA production in transgenic plants and mammals. Notably, increasing levels of EPA, DHA, and total PUFA production have been ascribed to the appropriate choices of heterologous enzymes, relative species, and host plants or mammals.165 Some representative engineering studies on the production of PUFAs (mainly EPA and DHA) and yields are shown in Figure 6. Agricultural oil production has been highly efficient and has the potential to become a sustainable ω-3 PUFA resource. Many studies have reported the enhanced production of terrestrial ω3 PUFA-rich oils via the transfer of genes from marine microorganisms (e.g., microalgae) or fish (e.g., salmon) into oilseed crops.172 The “reverse engineering” of any PUFA biosynthetic pathway requires the transgenic mobilization of multiple enzymatic activities, which requires the heterologous expression of a minimum of three transgenes.15 Figure 6 indicates that considerable effort has focused on the combined expression of Δ6-desaturase with Δ6-elongase and Δ5desaturase, the so-called “conventional” Δ6-desaturation pathway, in plants or animals. However, the production efficiency of AA and EPA following this pathway was low.166,173 To improve the biosynthesis yields of EPA and/or DHA, some novel sources of desaturases (Δ6-desaturase from the microalga Ostreococcus lucimarinus174 and Micromonas pusilla,175 and Δ5desaturase from ciliate protozoan Paramecium tetraurelia176) have been developed. The Δ8-desaturation pathway, which includes the combined expression of Δ9-elongase with Δ8desaturase and Δ5-desaturase, is potentially more effective in the biosynthesis of AA and EPA due to a reduced requirement for acyl exchange between phospholipids and the acyl-CoA pool. In fact, the Δ8-pathway produced much higher levels of AA and EPA than those obtained from the “conventional” pathway described previously.14 Besides the production in transgenic plants, the PUFA biosynthesis in mammals using FAT transgenes has also been highlighted. The elevated I


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expression of FAT-2 genes encoding ω-6 desaturase from C. elegans significantly enhanced LA content in transgenic cells (2.4-fold higher than those in wild-type cells).128 Many studies also contributed to the biosynthesis of PUFA in transgenic mammals. Kang et al.17a showed that mice engineered to carry a FAT-1 gene from C. elegans can add a double bond into an unsaturated fatty-acid hydrocarbon chain and convert ω-6 to ω3 fatty acids. The FAT-1 transgenic mice are capable of producing ω-3 fatty acids from the ω-6 type, leading to abundant ω-3 fatty acids with reduced levels of ω-6 fatty acids in their organs and tissues, without the need of a dietary ω-3 supply.126 Zhu et al.65 generated sFAT-1 transgenic mice by introducing mammal expression vector DNAs containing the sFAT-1 gene, which was synthesized from revised and optimized codons based on C. briggsae genomic gene for enhanced expression in mammals, into regular mice. Besides the biosynthesis of PUFAs in mice, corresponding transgenic engineering work in pigs was also highlighted. Lai et al.168 described the generation of hFAT-1 transgenic pigs, which express a humanized C. elegans gene and produce high levels of ω-3 fatty acids from ω-6 analogues (EPA 4.2%, DPA 1.7%, and DHA 1.8%). Their tissues have a significantly reduced ratio of ω-6/ω-3 fatty acids, which was decreased from 8.52 in the control to 1.69 in hFAT-1 transgenic pigs. Meanwhile, a recent study demonstrated that the sFAT-1 gene from C. briggsae could be functionally expressed in transgenic pigs, which could synthesize high-quality ω-3 PUFAs endogenously. Furthermore, the fatty acid compositions in the transgenic pigs were altered, and the levels of ω-6/ω-3 ratios were decreased from 14.53 in the control to 2.62 in sFAT-1 transgenic pigs.177 Besides the findings of novel desaturase and elongase sources for the optimal transgenic production of ω-3 PUFAs via the classical Δ6- or Δ8-desaturation pathway, studies also focused on the development of new sources of enzymes and related gene expressions for improving the production yields of ω-3 PUFAs via “Sprecher” and PKS pathways.178 Besides, the stateof-the-art progess in biosynthesis engineering studies pays attention to the biochemical production of ω-3 PUFAs other than EPA and DHA (e.g., ETA, C20:4, Δ8, 11, 14, 17),179 biosynthesis of PUFAs in new animals (e.g., zebrafish Danio rerio180 and polyps of corals181), and relationships between PUFA biosynthesis and the occurrence of nonalcoholic fatty liver disease in Western populations.182 All of the relative plant or animal studies revealed that the transgenic approach is an effective strategy for changing fatty acid composition of organisms and improving nutritional values of PUFA-rich products.

triacylglycerol (TAG).14,172c To overcome the limitations of some species, the incorporation of genes that encode appropriate acyltransferase enzymes, such as lyso-phosphatidylcholine acyltransferase (LPCAT), are of particular importance.183 The activity of LPCAT might be to preferentially remove fatty acids from the site of desaturation and directly incorporate them into the acyl-CoA pool, making them available for transgene-mediated elongation and for generating the carbon skeleton of LC-PUFAs.14 Although transgenic oilseeds are more common, limited studies have focused on “making fish oils in transgenic mammals”. FAT-1 transgenic mice are a representative model for enhanced biosynthesis of ω-3 PUFAs and relative fatty acids.4a,17a,126 Second, appropriate selection and expression of specific promoters benefit the efficient synthesis of PUFAs. The seedspecific promoters, such as napin, are recommended for use to ensure efficient expression of transgenes in target plants and coordinate with fatty acid synthesis and TAG assembly. A stepwise engineering study emphasized the uniform use of the napin promoter for nine transgenes for the effective production of ω-3 PUFAs in oilseed crops.23 Some studies have also demonstrated successful synthesis of PUFAs via the use of constitutive promoters, such as USP (promoter region of the unknown seed protein of Vicia faba) and CaMV (cauliflower mosaic virus) 35S; however, these studies reported poor yields.24,184 A typical success of the increased production of ω-3 PUFAs in FAT-1 transgenic mice and pigs was achieved by the chicken β-actin promoter.17a,168 Another successful enhancement of the production of EPA and DHA in E. coli and transgenic mice was also accomplished by the use of LacZ (βgalactosidase gene) and CMV (cytomegalovirus) promoters, respectively.65,185 Therefore, both plant and animal studies have clearly indicated that an appropriate choice of promoter is important for suitable lipid modification in the target species. Third, pertinent metabolism pathways should be fully considered for high-efficiency synthesis of PUFAs. LA and ALA can both become substrates during the synthesis of EPA. However, more intermediates may be generated in the presence of both substrates. The expression of FAT-1 transgenes could effectively convert ω-6 PUFAs into ω-3 PUFAs in animals.17a Compared to the FAT-1 genes, lower catalytic efficiency of plant-endogenetic ω-3 desaturases indicates inefficient synthesis. Therefore, highly efficient synthesis of EPA can be achieved by one or more of these approaches: (i) selection of target plants that lack ALA, such as sunflower, safflower, and cottonseed;186 (ii) the use of Δ15- and Δ17-desaturases for their powerful ability of converting ω-6 into ω-3 fatty acids;187 or (iii) following the Δ8-desaturation pathway. In addition, the fine-tuning of the fatty acid flux between the acyl-CoA, phospholipid, and TAG pools is capable of maximizing PUFA yields. Current progress indicates that yields above 20% have not yet been achieved because of bottlenecks that became obvious in the endogenous biosynthetic pathways when heterologous genes were expressed.188 The main bottleneck was the suboptimal acyl-exchange transference of desaturated intermediates from the lipid class pool to the acylCoA pool. For example, a previous study revealed that the lack of available Δ6-desaturated acyl-CoA substrates in the acyl-CoA pool limits the biosynthesis of elongated C20 fatty acids and disrupts the alternating sequence of lipid-linked desaturations and acyl-CoA dependent elongations.184 This problem can be solved by (i) investigating the acyl-exchange mechanism in organisms with the ability to biosynthesize EPA and DHA by

6.2. Strategies for Enhancing the Production of PUFAs in Transgenic Organisms

Different yields of PUFAs achieved in plants and animals expressing alternative transgenes for aerobic PUFA biosynthetic pathways indicate several strategies that may be used to enhance overall PUFA synthesis and direct biosynthesis toward specific PUFA end products. First, it is very important to consider whether the codon matches with the species in order to enhance PUFA production in transferring genes from one species to another. Also, the choice of a target plant or animal for the introduction of a transgenic PUFA pathway is important because of the species diversity of various endogenous metabolic pathways, especially those that pertain to the transfer of PUFAs between phosphatidylcholine and CoA and the accumulation of J


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construction, and microinjections, will provide substantial support for the development of new transgenic animal models. Finally, further studies will address the nutrigenetic and nutrigenomic research aspects, which consider PUFA−gene interaction, genetic variation, and dietary response.

themselves (e.g., marine microalgae) and identify the LPCAT species with the capacity of effective transference of PUFAs or (ii) directly inserting acyl-CoA desaturase transgenes and improving the elongation efficiency. Most plant desaturases are acyl-lipid desaturases, whereas all of the well-known mammalian desaturases are acyl-CoA desaturases.189 Recently, the first acyl-CoA Δ6-desaturase from the microalga Ostreococcus tauri was identified.190 When this desaturase was coexpressed with the acyl-CoA Δ6-elongase from the moss Physcomitrella patens and the lipid-linked Δ5-desaturase from the diatom Phaeodactylum tricornutum, high proportions of AA or EPA were obtained because nearly all of Δ6-desaturated products were elongated. However, it is still unclear whether this desaturase could improve the Δ6-elongation efficiency in transgenic organisms. Moreover, the yield of DHA was unusually low, especially for transgenic plants. Future efforts focusing on the identification of an acyl-CoA Δ5 desaturase will likely improve overall elongation rates, particularly given that Δ5-elongation appears to be the narrowest point en route to DHA.191 Venegas-Calerón et al.192 summarized that the use of acyl-CoAdependent desaturases is predicted to bypass the metabolic bottleneck generated by substrate dichotomy between the desaturase and the elongase, and thus the biosynthesis and homeostasis of the acyl-CoA pool deserve to be further investigated.

AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest. Biographies

7. FUTURE PROSPECTS The use of transgenic plants to synthesize “fish oils” or for the biosynthesis of PUFAs in transgenic animals may prove to be a sustainable source for the production of these important fatty acids and may demonstrate the utility of genetically modified technology to improve the state of human health and nutrition. Although increased production of PUFAs has succeeded in various transgenic studies, there are still many challenges for the optimal use of transgenic approaches in the field. Depending on the transgenic mechanism used, different substrate requirements for the alternating desaturation and elongation steps appear to be a major limiting factor in PUFA production. This problem is highlighted by the lack of a logistical understanding of the acyl trafficking between the phospholipid and acyl-CoA pools. Clear recognition of the mechanisms underlying the exchange between these two pools, and the isolation of genes encoding enzymes involved in this process that prefer or at least do not discriminate against PUFA acyls, will be very useful.192 Moreover, the effect of the elongation complex on PUFA synthesis has not been thoroughly studied. Four consecutive reactions are involved in the extension of the fatty-acyl chain, and the condensing enzymes or elongases represent just one component of the elongation complex. A further understanding of this process may be beneficial to coordinating heterologous condensing enzymes with other components of the endogenous complex.191 Data about the synthesis of PUFAs from transgenic animals is less varied than plant transgenic biosynthesis studies. The transgenic animal model (e.g., FAT1 transgenic pigs and mice) provides a new strategy for producing ω-3 fatty acid-rich foodstuffs (e.g., meat, milk, and eggs). This genetic approach is a cost-effective and sustainable way of producing ω-3 essential fatty acids to meet increasing demand in future.4a On the basis of the current technology, future efforts will focus on the engineering of enhanced PUFA production in other transgenic animals, such as cows, sheep, chickens, etc. In addition, improvements to related technologies, such as isolation and characterization of transgenes, vector

Dr. Jingjing Jiao received her Ph.D. degree in Food Science at Zhejiang University in China in 2008. She completed her postdoctoral experience in lipid medicine and technology at Massachusetts General Hospital and Harvard Medical School (Boston, Massachusetts, U.S.A.) in 2011. She is currently a lecturer of School of Public Health at Zhejiang University. Her research interests include (i) nutrigenomic studies on the prevention and control of cancer and cardiovascular diseases via nutrient metabolism; (ii) molecular cloning and transgenic engineering of polyunsaturated fatty acids; and (iii) control and risk assessment of in vivo toxicity of chemical contaminants and heavy metals.

Dr. Yu Zhang obtained his Ph.D. degree in Food Science at Zhejiang Univeristy in China in 2008. He finished his postdoctoral research in lipid medicine and technology at Massachusetts General Hospital and Harvard Medical School (Boston, Massachusetts, U.S.A.) in 2011. He has been an Associate Professor of Department of Food Science and Nutrition at Zhejiang University since 2011. He received the International Union of Pure and Applied Chemistry (IUPAC) Honorable Mention Award for Young Chemists in 2009 and China National Top 100 Excellent Ph.D. Dissertation Award in 2011. His K


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research interests include (i) analytical biochemistry, transgenic engineering, and nutrigenomics of polyunsaturated fatty acids; (ii) analytical chemistry, formation pathway, prevention mechanism, in vivo toxicity control, and risk assessment of chemical contaminants in food and related food safety researches; and (iii) extraction, separation, purification, and functionality of phytochemicals from natural products.

ACKNOWLEDGMENTS This study was financially supported by the National High Technology Research and Development Program of China (863 Program, Grant No. 2010AA023003) and National Natural Science Foundation of China (Grant No. 31201307). ABBREVIATIONS AA arachidonic acid ACP acyl carrier protein acyl-ACP acyl−acyl carrier protein acyl-CoA acyl-coenzyme A ALA α-linolenic acid DGLA dihomo-γ-linolenic acid DHA docosahexaenoic acid DPA docosapentaenoic acid EDA eicosadienoic acid ELO elongase EPA eicosapentaenoic acid ER endoplasmic reticulum ETA eicosatetraenoic acid ETrA eicosatrienoic acid FAD fatty acid desaturase FAE fatty acid elongase FAS fatty acid synthase GLA γ-linolenic acid LA linoleic acid LC-PUFA long-chain polyunsaturated fatty acid LPCAT lyso-phosphatidylcholine acyltransferase KCR 3-ketoacyl-CoA reductase KCS 3-ketoacyl-CoA synthase NADH nicotinamide adenine dinucleotide NADPH nicotinamide adenine dinucleotide phosphate OA oleic acid ORF open-reading-frame OTA octadecatetraenoic acid PA palmitic acid PKS polyketide synthase PUFA polyunsaturated fatty acid SA stearic acid siRNA small interfering RNA SREBP sterol regulatory element binding protein TAG triacylglycerol TPA tetracosapentaenoic acid REFERENCES (1) Paoloni-Giacobino, A.; Grimble, R.; Pichard, C. Clin. Nutr. 2003, 22, 429. (2) (a) Kummerow, F. A. Am. J. Clin. Nutr. 1979, 32, 58. (b) Weylandt, K. H.; Kang, J. X. Lancet 2005, 366, 618. (3) (a) Salem, N., Jr.; Simopoulos, A. P.; Galli, C.; Lagarde, M.; Knapp, H. R. Lipids 1996, 31, S1. (b) Simopoulos, A. P. Am. J. Clin. Nutr. 1999, 70, 560S. (4) (a) Kang, J. X. J. Membr. Biol. 2005, 206, 165. (b) Connor, W. E. Am. J. Clin. Nutr. 2000, 71, 171S. (c) Leaf, A.; Kang, J. X. World Rev. Nutr. Diet. 2001, 89, 161. L


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