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Research Article
Open Access Peer-reviewed

Molecular Cloning and Functional Characterisation of a Polyunsaturated Fatty Acid Elongase in a Marine Bivalve Crassostrea angulata

Hongkuan Zhang, Helu Liu, Dewei Cheng, Hongxing Liu, Huaiping Zheng
Journal of Food and Nutrition Research. 2018, 6(2), 89-95. DOI: 10.12691/jfnr-6-2-4
Published online: February 09, 2018

Abstract

The elongases of fatty acids (ELO) are essential for long chain polyunsaturated fatty acid (LC-PUFA) biosynthesis, and their activities depend on the substrates. The full length cDNA of Crassostrea angulata ELO (CaELO) was cloned by RACE PCR and its function was confirmed. The CaELO encodes a polypeptide of 309 amino acid residues, which containes a histidine box HXXHH motif conserved in all elongases and shares high similarity to the elongases of Chlamys nobilis and Octopus vulgaris. Phylogenetic analysis showed that the putative elongase was placed in the same group with ELOVL2 and ELOVL5, which have been demonstrated to be critical enzymes participating in the biosynthesis of PUFAs in vertebrates. When expressed in Saccharomyces cerevisiae, CaELO was able to elongate n-3 and n-6 PUFA substrates with chain lengths of C18 and C20, indicating that the CaELO had similar substrate specificities to vertebrate ELOVL5. CaELO had lower activity to elongate monounsaturated fatty acids, but had not activity to saturated fatty acids. Interestingly, the conversion rate of PUFAs depended on the length of carbon chain, the number of double bond, and n-3 / n-6 series in the species.

1. Introduction

Polyunsaturated fatty acids (PUFAs) are important components of cellular structure and function and serve as precursor to eicosanoids, including prostaglandins and leukotrienes. These eicosanoids have a number of functions in animals, involving in inflammatory responses, regulation of blood pressure, and reproductive function 1. PUFAs, such as linoleic acid (LA, 18:2n-6) and α-linolenic acid (ALA, 18:3n-3) which are essential fatty acids for biosynthesis of long-chain polyunsaturated fatty acids (LC-PUFA), cannot be endogenously biosynthesized by human and so must be obtained from diet.

In animals, the ability to produce PUFA varies and is dependent on their complement of fatty acyl desaturase and elongases. In vertebrates, the pathway of biosynthesis LC-PUFAs has been clearly described 2, which involves sequential desaturation and elongation of essential PUFA precursors, LA and ALA. While in molluscs, fatty acyl elongase genes with differing fatty acid (FA) substrate specificities are only described in Chlamys nobilis and Octopus vulgaris 3, 4, 5.

Microalgae, the major source of LA, ALA, C20 and C22 PUFAs, are the main feed for the filter-feeder bivalves 6, 7. Previous studies have showed that these PUFAs are crucial to the survival, growth, development and reproduction for molluscs, such as Mytilus galloprovincialis 8, Babylonia areolata 9, Solen marginatus 10, and Ostrea angasi 11, etc. Decades of research on PUFAs of elongation and desaturation activity in bivalves has improved our understanding of the PUFAs synthesis pathway. In the early studies, by using labeled PUFA precursors, clam Mesoderma mactroides and Pacific oyster Crassostrea gigas were demonstrated to have the ability to desaturate and elongate PUFA precursors 12, 13. In addition, Scapharca broughtoni and Mytilus edulis were reported to have the capabilities of de novo synthesis for some peculiar fatty acids (FAs) called nonmethylene interrupted fatty acids (NMIs) 14, 15. More importantly, in recent years, our lab has provided the first compelling evidence that C. nobilis can de novo biosynthesize 20:5n-3 and 20:4n-6 from PUFA precursors though the “Δ8 pathway” 16.

The Crassostrea oysters are important edible marine bivalves, which have been cultured worldwide. It is known that oysters are rich in PUFAs 17, 18, and part of them obtained from their diets. However, whether they have the ability to biosynthesize PUFAs themselves is still unknown.

In the present study, the cDNA of fatty acyl elongase was isolated in the Crassostrea angulata, and function of the encoded polypeptide was characterized in yeast Saccharomyces cerevisiae to investigate whether the gene could elongate PUFA substrates.

2. Materials and Methods

2.1. Cloning of Putative PUFA Elongase from C. angulata

The adult oysters used in the present study were sampled from a cultured population at Nan'ao Island of Guangdong Province, China. The adductor was sampled and total RNA was extracted using TransZol Up reagent (TransGen Biotech, China). Then, 1 μg of total RNA was used as template for cDNA synthesis by using SuperScript III reverse transcriptase PCR and Oligo-dT as primers following the instructions of the manufacturer (Invitrogen). Partial cDNA sequence of elongase was obtained from transcriptome sequencing of C. angulate (SRX481252). Then forward primer CAEF and reverse primer CAER (Table.1) were used to confirm the cDNA fragment. PCR amplification was performed under the following PCR conditions: initial denaturation at 95°C for 2min, 35 cycles of denaturation at 95°C for 30s, annealing at 60°C for 30s, and extension at 72°C for 90s. The PCR products were separated on 1% agarose gel, and then purified with a PCR purification kit (TransGen Biotech). The purified PCR product was ligated with the pEASY- T1 Cloning Kit (TransGen Biotech), and transformed into Trans5α Chemically Competent Cell (TransGen Biotech). The recombinants were identified through blue-white color selection in ampicillin containing LB plates and screened with M13-47 and RV-M primers, and then sequenced on both strands.

Based on the partial cDNA sequence of the elongase, the complete CaELOVL cDNA was amplified through 3′-RACE cDNA and 5′-RACE PCR with the SMARTer™ RACE cDNA Amplification Kit (Clontech, Mountain View, CA) and LA Taq polymerase (TaKaRa, Japan), using touch-down PCR and nested PCR strategy. The touch-down PCR was carried out in a 50 μl volume containing 2.5 μl 5′-RACE cDNA (or 3′-RACE cDNA) template, 1 μl CAE51 (or CAE31) primer (10 mmol L-1), 1 μl UPM (long UPM 10 mmol L-1, short UPM 10 mmol L-1) (Table 1), 0.5 μl LA Taq DNA Polymerase (5 U/μl), 5 μl 10×LA Taq Buffer II (Mg2+ Plus), 8 μl dNTP Mixture (2.5 mM each), and 32 μl sterilized ultrapure water. The PCR reaction program was performed as follows: 94°C for 2 min followed by 35 cycles of 94°C for 30 s, 68°C -60 °C for 30 s in the initial 9 cycles decrementing 1°C /cycle, 60°C for 30 s for the remaining 26 cycles, 72°C for 2 min, and a final extension step at 72°C for 10 min. The products of touch-down PCR were used as template for the subsequent nested PCR. The expected DNA fragment was separated on a 1% agarose gel and then purified with a PCR purification kit (TransGen Biotech). Purified DNA fragment was sub-cloned into pEASY- T1, and then transformed into Trans5α Chemically Competent Cell. Positive recombinant clones were identified by blue-white color selection in ampicillin-containing LB plates and PCR screening with M13-47 and M13-RVM primers, and then sequenced by a commercial company (Sangon Biotech, Shanghai, China). The full length cDNA of CaELOVL was aligned from the overlapping cDNA clones.

2.2. Phylogenetic Analysis of Putative PUFA Elongase from C. angulata

A phylogenetic tree was constructed on the basis of protein sequence of ELOVL from the putative C. angulata ELOVL (CaELOVL) and other organisms, including vertebrate and invertebrate. A multiple sequence alignment was performed using ClustalX2.1 and a phylogenetic tree was made using the neighbor-joining method in MEGA 5.1 package. Confidence in the resulting phylogenetic tree branch topology was measured by bootstrapping through 1000 iterations.

2.3. Real-time PCR Analysis of CaELOVL Transcript Levels

Total RNA from different tissues of adductor, gill, gonad and mantle was isolated as described above. First-strand cDNA was synthesized using RT reagent Kit with gDNA Eraser (Takara, Dalian, China) according to the manufacturer’s instructions. Two gene-specific primers were designed to amplify a product of 158 bp, which was quantified by real-time RT-PCR with the SYBR®Premix Ex Taq™ II Kit (Perfect Real Time) (Takara, Japan) following the manufacturer’s protocol. The reaction was performed in a LightCycler® 480 (Roche). A total reaction volume of 20 μl containing 10 μl SYBR® Premix Ex Taq™ II, 2 μl the four-fold diluted cDNA, 0.8 ml each primer (CAEE-F and CAEE-R to amply CaELOVL gene, β-actin-F and β-actin-R to amply β-actin, 10 mmol L-1, Table 1) and 6.4 μl ultra-pure water was carried out following a denaturation step of 95°C for 30 s, 45 cycles of 95°C for 5 s, 60°C for 30 s, a melting curve analysis from 65°C to 95°C and a cooling step of 40°C for 10 min. Each sample was run in triplicate. Data were analyzed by using the LightCycler480 software (Roche) after the PCR program. Relative mRNA expression level of CaELOVL was determined with the 2-△△Ct algorithm with β-actin from C. angulata as the internal control.

2.4. Heterologous Expression of Elongase ORFs in Yeast

Expression primers were designed for PCR cloning putative elongase cDNA ORF. The forward primer and the reverse primer was CAEF-1, contained a HindIII site (underlined), and CAER-1, contained an EcoRI site (underlined). PCR was performed using TransStart® FastPfu DNA Polymerase (Trans GenBiotech) following the manufacturer's instructions. Amplification involved an initial denaturation step at 95°C for 2 min, followed by 30 cycles of denaturation at 95°C for 20 s, annealing at 56°C for 30 s, and extension at 72°C for 60 s. After PCR, the DNA fragments were cut with HindIII/EcoRI and ligated into the similarly digested yeast expression vector pYES2.0 (Invitrogen), which was then used to transform DH5α E. coli competent cells. Transformation of yeast with recombinant plasmids and yeast culture were carried out using the S.c. EasyComp Transformation Kit (Invitrogen Ltd.). Selection of yeast containing the elongase/pYES2 constructs was on S. cerevisiae minimal medium minusuracil (SCMM-uracil). Culture of the recombinant yeast was carried out in SCMM-uracil broth as described previously 19, using galactose induction of gene expression. Each culture was supplemented with one of the following PUFA substrates: C18:2n-6, C18:3n-3, C20:4n-6,C20:4n-6, C20:5n-3 and C22:3n-6. Yeast cells were harvested, washed, and dried, and then lipid was extracted by homogenization in chloroform/methanol (2:1, vol/vol) containing 0.01% butylated hydroxytoluene (Sigma, Ltd) as antioxidant. FAMEs were prepared, extracted, purified by TLC on 20 cm × 20 cm × 0.25 mm silica gel 60 plate (Merck, Germany), and analyzed using gas chromatography (GC-17A, Shimadzu, Kyoto, Japan) equipped with a hydrogen flame ionization detector and a30 m × 0.25 mm × 0.25 μm capillary column (VF-23ms, Varian, USA), all as described below. The column was temperature programmed from 50 to 150°C at 6°C/min and 150 to 230°C at 5°C/min. Hydrogen was used as carrier gas at a flow rate of 1.1 ml/min. The temperature of detector and injection port was set at 250°C. Individual methylesters were identified by comparison to authentic standards. The proportion of substrate fatty acid converted to the longer chain fatty acid product was calculated from the gas chromatograms as 100·[Product Area / (Product Area + Substrate Area)].

3. Results

3.1. CaELOVL cDNA Sequences and Phylogenetics

A 2238bp full-length cDNA sequence was obtained by 5′- and 3′-RACE PCR and was deposited in the GenBank database under the accession number KY012771. It contains an ORF of 930bp (including stop codon) encoding a putative protein of 309 amino acid (aa), sharing 57% and 54% aa sequence identity with C. nobilis elongase (KF245423) and O. vulgaris (AFM93779.1), respectively. And it has 42-45% identity to Xenopus tropicalis ELOVL5 (NP001011248), Siganus canaliculatus ELOVL5 (ADE34561), Gallus gallus ELOVL2 (NP001184237), Homo sapiens ELOVL5 (NP068586). The newly cloned fatty acyl elongase protein possessed the typical histidine box HXXHH motif (Figure 1) conserved in all elongases. Seven putative transmembrane-spanning domains were predicted using TMHMM 2.0, five of which (I, II, V, VI, VII) had been reported in a mouse ELOVL2 20. A phylogenetic tree was constructed on the basis of the aa sequence alignments between the putative CaELOVL and other ELOVL family members (2, 5) from invertebrates, and as well as from vertebrates (Figure 2). The CaELOVL had the closest relationship with ELOVL from C. nobilis, then O. vulgaris, and formed a group close to ELOVL2 and ELOVL5 from vertebrates.

  • Figure 1. Comparison of the amino acid sequences of PUFA elongases cloning from C. angulata (KY012771) with those from C. nobilis (KF245423), O. vulgaris (AFM93779), Xenopus tropicalis (NP001011248), Siganus canaliculatus (ADE34561), Gallus gallus (NP001184237), and Homo sapiens (NP068586). Identical residues are shaded black and similar residues are gray. The threshold for similarity shading was set at 25%. Indicated is the conserved histidine box motif. Seven (I-VIII) putative membrane-spanning domains (underlined with dashed line) were identified using TMHMM 2.0, five of which (I, II, V, VI, VII) were predicted by Tvrdik et al. (2000)
3.2. Tissue Distribution of CaELOVL

CaELOVL transcripts were detected in all tested tissues (Figure 3), and the level was significantly different among the tissues (P < 0.05). And there was a significantly higher level in gill and mantle than in gonad and adductor.

3.3. Functional Characterization

Yeast transformed with vector containing no insert contained only four endogenous fatty acids (C16:0, C16:1n-7, C18:0, and C18:1n-9) 19, together with whichever exogenous FA was added. The results were consistent with the well-established observations of lack of PUFA elongase activity in S. cerevisiae 21, 22. The FA compositions of yeast transformed with the pYES2-ELOVL construct and grown in the presence of different substrates were shown in Figure 4.

  • Figure 4. Functional characterization of CaELOVL in transgenic S. cerevisiae grown in the presence of fatty acids (FAs). A, B, C, D, E, F and G represent control and adding FA substrate of C18:2n-6, C20:3n-6, C22:3n-6, C20:4n-6, C18:3n-3 and C20:5n-3, respectively. Fatty acids were extracted from yeast transformed with pYES2 vector containing the ORF of the putative fatty acyl elongase cDNA as an insert. Peaks 1-4 represent the main endogenous FAs of S. cerevisiae, namely C16:0 (1), C16:1n-7 (2), C18:0 (3) and C18:1n-9 (4). The remaining main additional peaks (6-17) correspond to the exogenously added FAs and the products of their elongation-C18:2n-6 (7), C20:2n-6 (8), C20:3n-6 (9), C22:3n-6 (10), C22:3n-6 (11), C20:4n-6 (12), C22:4n-6 (13), C18:3n-3 (14), C20:3n-3 (15), C20:5n-3 (16) and C22:5n-3(17). Other minor peaks are 20:1n-9 (5) and 20:1n-7 (6), the latter two resulting from the elongation of 18:1n-9 and 18:1n-7.

All substrates including B (C18:2n-6), C (20:3n-6), E (C20:4n-6), F (C18:3n-3), and G (C20:5n-3) had their corresponding products except for D (C22:3n-6). Moreover, we found that conversion rate is significantly different (P < 0.05), varying from 25% to 0.81% (Table 2).

4. Discussion

Polyunsaturated fatty acids (PUFAs), particularly 18- to 22-carbon PUFAs, have obtained a high-profile position in the biomedical and nutraceutical areas because of their specific therapeutic roles in certain clinical conditions. Moreover, public awareness on eating healthy has also brought these PUFAs to the attention of the consumer.

In the past decades, the synthesis pathways of important PUFAs have been clearly described in vertebrates 23, 24. As the second largest Phylum in animal kingdom, mollusks have provided delicious and health food for human due to their enriched higher protein and PUFAs 23, 25, 26. However, mechanism of their enrichment in PUFAs has not been clearly described, although there have been a few reports about biosynthesis PUFAs mechanism in an octopus 4, 5 and a bivalve 27 in recent years.

In the present study, the activity of CaELOVL was found similar to that of ELOVL5 more than that of ELOVL2, which involves specifically in the elongation of C18 and C20 FAs, respcetively. The result is consistent to the previous study in the mollusks 4, 27 and in the vertebrates 24, 28.

In the past years, bivalves have been regarded as having no or limit ability to biosynthesize n-3 and n-6 long chain PUFAs by elongation and desaturation of the precursors 18:2n-6 and 18:3n-3 17, 29. Instead bivalves have been thought to obtain the PUFAs from their diets 30, 31. However, in the present study, the oyster was identified to have the ability to biosynthesize PUFAs by itself, and previous study in two mollusks of O. vulgaris 4 and C. nobilis 16 had the same findings, respectively. Therefore, we conclude that marine mollusks have the ability to biosynthesize PUFAs by themselves.

More interestingly, from the results listed in Table 2, we found that the conversion rate of PUFAs was related to n-3 / n-6 substrates. Significantly higher conversion rate was found with n-6 PUFA substrates than homologous n-3 substrates; however, in vertebrates, most of functionally characterized PUFA biosynthesis genes/enzymes show more activity toward n-3 PUFA substrates 32, 33, 34, 35. Marine microalgae enrich in n-3 series PUFAs 30, the oyster can obtain n-3 PUFAs by feeding these microalgae, this probably explains why the synthesizing ability of the n-3 series PUFAs became weaker than that of the n-6 series PUFAs. This might be an adaption to environment in the evolution of animals.

In vertebrates, ARA and EPA are biosynthesised from the dietary essential C18 PUFA 18:2n-6 and C18:3n-3 through two alternative pathways, the ‘classical’ Δ6-pathway (Δ6 desaturation → elongation → Δ5 desaturation), or alternatively through the so-called ‘Δ8-pathway’ (elongation → Δ8 desaturation → Δ5 desaturation). In the present, C20:2n-6 and C20:3n-3 can be biosynthesised from the dietary essential 18:2n-6 and C18:3n-3, namely 18:2n-6 → 20:3n-6 and C18:3n-3 → C20:3n-3 for the Δ8-pathway in C. angulata. Although gene responsible for elongation step of this pathway has now been identified in C. angulata, no Fad cDNA with Δ5 and Δ8-desaturase activity has yet been identified and, consequently, it remains unclear whether the C. angulata can biosynthesise ARA and EPA from the dietary essential 18:2n-6. However, in C. nobilis, it has been confirmed that C. nobilis could efficiently biosynthesis ARA and EPA from 18:2n-6 and C18:3n-3through ‘Δ8-pathway’ 27, based on these results, we speculated that the alternative “Δ8-pathway” may be the PUFA biosynthesis pathway in C. angulata.

In conclusion, the oyster Crassostra angulata has the ability to biosynthesize PUFAs by itself. And it has a significantly higher conversion rate in n-6 PUFA substrates than homologous n-3 substrates. Moreover, we speculated that the alternative “Δ8-pathway” may be the PUFA biosynthesis pathway in C. angulata.

Acknowledgements

We are very grateful to Dr. Chiju Wei (Multidisciplinary Research Center, Shantou University) for his careful revision and many constructive comments. Funding for this research was provided by China Modern Agro-industry Technology Research System (CARS-48), Guangdong Province Natural Sciences Foundation (2015A030311027), Science & Technology Plan (2015B090903081) and Yangfan Plan (14600706), China.

References

[1]  Vance, J. E., Vance, D. E. (Eds.). 2008. Biochemistry of lipids, lipoproteins and membranes. Elsevier.
In article      
 
[2]  Hastings, N., Agaba, M., Tocher, D. R., Leaver, M. J., Dick, J. R., Sargent, J. R., & Teale, A. J. 2001. A vertebrate fatty acid desaturase with Δ5 and Δ6 activities. PNAS USA, 98(25), 14304-14309.
In article      View Article  PubMed
 
[3]  Liu, H., Zheng, H., Wang, S., Wang, Y., Li, S., Liu, W., & Zhang, G. 2013. Cloning and functional characterization of a polyunsaturated fatty acid elongase in a marine bivalve noble scallop Chlamys nobilis Reeve. Aquaculture, 416, 146-151.
In article      View Article
 
[4]  Monroig, O., Guinot, D., Hontoria, F., Tocher, D. R., & Navarro, J. C. 2012. Biosynthesis of essential fatty acids in Octopus vulgaris (Cuvier, 1797): Molecular cloning, functional characterisation and tissue distribution of a fatty acyl elongase. Aquaculture, 360, 45-53.
In article      View Article
 
[5]  Monroig, Ó., Tocher, D. R., & Navarro, J. C. 2013. Biosynthesis of polyunsaturated fatty acids in marine invertebrates: recent advances in molecular mechanisms. Mar Drugs, 11(10), 3998-4018.
In article      View Article  PubMed
 
[6]  Bellou, S., Baeshen, M. N., Elazzazy, A. M., Aggeli, D., Sayegh, F., & Aggelis, G. 2014. Microalgal lipids biochemistry and biotechnological perspectives. Biotechnol Adv, 32(8), 1476-1493.
In article      View Article  PubMed
 
[7]  Maadane, A., Merghoub, N., Ainane, T., El Arroussi, H., Benhima, R., Amzazi, S., & Wahby, I. 2015. Antioxidant activity of some Moroccan marine microalgae: Pufa profiles, carotenoids and phenolic content. J Biotechnol, 215, 13-19.
In article      View Article
 
[8]  Pettersen, A. K., Turchini, G. M., Jahangard, S., Ingram, B. A., & Sherman, C. D. 2010. Effects of different dietary microalgae on survival, growth, settlement and fatty acid composition of blue mussel (Mytilus galloprovincialis) larvae. Aquaculture, 309(1), 115-124.
In article      View Article
 
[9]  Sangsawangchote, S., Chaitanawisuti, N., & Piyatiratitivorakul, S. 2010. Reproductive performance, egg and larval quality and egg fatty acid composition of hatchery-reared Spotted Babylon (Babylonia areolata) broodstock fed natural and formulated diets under hatchery conditions. Int J Fish Aquaculture, 2(1), 044-048.
In article      View Article
 
[10]  Da Costa, F., Nóvoa, S., Ojea, J., & Martínez-Patiño, D. 2011. Changes in biochemical and fatty acid composition of the razor clam Solen marginatus (Solenidae: Bivalvia) during larval development. Mar Biol, 158(8), 1829-1840.
In article      View Article
 
[11]  O'Connor, S., Moltschaniwskyj, N., Bolch, C. J., & O'Connor, W. 2012. Dietary influence on growth and development of flat oyster, Ostrea angasi (Sowerby, 1871), larvae. Aquac Res, 43(9), 1317-1327.
In article      View Article
 
[12]  De Moreno, J. E., Moreno, V. J., & Brenner, R. R. 1976. Lipid metabolism of the yellow clam, Mesodesma mactroides: 2-polyunsaturated fatty acid metabolism. Lipids, 11(7), 561-566.
In article      View Article
 
[13]  Waldock, M. J., & Holland, D. L. 1984. Fatty acid metabolism in young oysters, Crassostrea gigas: polyunsaturated fatty acids. Lipids, 19(5), 332-336.
In article      View Article
 
[14]  Zhukova, N. V. 1991. The pathway of the biosynthesis of non-methylene-interrupted dienoic fatty acids in molluscs. Comp Biochem Physiol. 100B (4), 801-804.
In article      View Article
 
[15]  Zhukova, N. V., & Svetashev, V. I. 1986. Non-methylene-interrupted dienoic fatty acids in molluscs from the sea of Japan. Comp Biochem Physiol. 83B (3), 643-646.
In article      View Article
 
[16]  Liu, H., Zhang, H., Zheng, H., Wang, S., Guo, Z., & Zhang, G. 2014. PUFA biosynthesis pathway in marine scallop Chlamys nobilis Reeve. J Agr Food Chem, 62(51), 12384-12391.
In article      View Article  PubMed
 
[17]  Chu, F. L., & Greaves, J. 1991. Metabolism of palmitic, linoleic, and linolenic acids in adult oysters, Crassostrea virginica. Mar Biol, 110 (2), 229-236.
In article      View Article
 
[18]  Cruz-Romero, M. C., Kerry, J. P., & Kelly, A. L. 2008. Fatty acids, volatile compounds and colour changes in high-pressure-treated oysters (Crassostrea gigas). Innov Food Sci Emerg, 9 (1), 54-61.
In article      View Article
 
[19]  Hastings, N., Agaba, M., Tocher, D. R., Leaver, M. J., Dick, J. R., Sargent, J. R., & Teale, A. J. 2001. A vertebrate fatty acid desaturase with Δ5 and Δ6 activities. P Natl Acad Sci USA, 98(25), 14304-14309.
In article      View Article  PubMed
 
[20]  Tvrdik, P., Westerberg, R., Silve, S., Asadi, A., Jakobsson, A., Cannon, B. & Jacobsson, A. 2000. Role of a new mammalian gene family in the biosynthesis of very long chain fatty acids and sphingolipids. J Cell Biol, 149(3), 707-718.
In article      View Article  PubMed
 
[21]  Agaba, M., Tocher, D. R., Dickson, C. A., Dick, J. R., & Teale, A. J. 2004. Zebrafish cDNA encoding multifunctional fatty acid elongase involved in production of eicosapentaenoic (20: 5n-3) and docosahexaenoic (22: 6n-3) acids. Mar Biotechnol, 6(3), 251-261.
In article      View Article
 
[22]  Morais, S., Monroig, O., Zheng, X., Leaver, M. J., & Tocher, D. R. 2009. Highly unsaturated fatty acid synthesis in Atlantic salmon: characterization of ELOVL5-and ELOVL2-like elongases. Mar Biotechnol, 11(5), 627-639.
In article      View Article  PubMed
 
[23]  Li, Y., Monroig, O., Zhang, L., Wang, S., Zheng, X., Dick, J. R., & Tocher, D. R. 2010. Vertebrate fatty acyl desaturase with Δ4 activity. PNAS USA, 107(39), 16840-16845.
In article      View Article  PubMed
 
[24]  Monroig, Ó., Wang, S., Zhang, L., You, C., Tocher, D. R., & Li, Y. 2012. Elongation of long-chain fatty acids in rabbitfish Siganus canaliculatus: Cloning, functional characterisation and tissue distribution of Elovl5-and Elovl4-like elongases. Aquaculture, 350, 63-70.
In article      View Article
 
[25]  Soriguer, F., Serna, S., Valverde, E., Hernando, J., Martín-Reyes, A., Soriguer, M., & Esteva, I. 1997. Lipid, protein, and calorie content of different Atlantic and Mediterranean fish, shellfish, and molluscs commonly eaten in the south of Spain. Eur J Epidemiol, 13(4), 451-463.
In article      View Article  PubMed
 
[26]  Baby, R. L., Hasan, I., Kabir, K. A., & Naser, M. N. 2010. Nutrient analysis of some commercially important molluscs of Bangladesh. J Sci Res, 2(2), 390-396.
In article      View Article
 
[27]  Liu, H., Zhang, H., Zheng, H., Wang, S., Guo, Z., & Zhang, G. 2014. PUFA biosynthesis pathway in marine scallop Chlamys nobilis Reeve. J Agr Food Chem, 62(51), 12384-12391.
In article      View Article  PubMed
 
[28]  Agaba, M. K., Tocher, D. R., Zheng, X., Dickson, C. A., Dick, J. R., & Teale, A. J. 2005. Cloning and functional characterisation of polyunsaturated fatty acid elongases of marine and freshwater teleost fish. Comp Biochem Physiol. 142B (3), 342-352.
In article      View Article  PubMed
 
[29]  Waldock, M. J., & Holland, D. L. 1984. Fatty acid metabolism in young oysters, Crassostrea gigas: polyunsaturated fatty acids. Lipids, 19(5), 332-336.
In article      View Article
 
[30]  Viso, A.C., Marty, J.C., 1993. Fatty acids from 28 marine microalgae. Phytochemistry 34 (6), 1521-1533.
In article      View Article
 
[31]  Zhukova, N.V., Imbs, A.B., Yi, L.F., 1998. Diet-induced changes in lipid and fatty acid composition of Artemia salina. Comp Biochem. Physiol. 120B (3), 499-506.
In article      View Article
 
[32]  Zheng, X., Seiliez, I., Hastings, N., Tocher, D. R., Panserat, S., Dickson, C. A., & Teale, A. J. 2004. Characterization and comparison of fatty acyl Δ6 desaturase cDNAs from freshwater and marine teleost fish species. Comp Bioch Physiol. Part B, 139(2), 269-279.
In article      View Article  PubMed
 
[33]  Hastings, N.; Agaba, M. K.; Tocher, D. R.; Zheng, X.; Dickson, C. A.; Dick, J. R.; Teale, A. J. 2004. Molecular cloning and functional characterization of fatty acyl desaturase and elongase cDNAs involved in the production of eicosapentaenoic and docosahexaenoic acids from α-linolenic acid in Atlantic salmon (Salmo salar). Mar Biotechnol, 6, 463-474.
In article      View Article
 
[34]  Tocher, D. R.; Zheng, X.; Schlechtriem, C.; Hastings, N.; Dick, J. R.; Teale, A. J. 2006. Highly unsaturated fatty acid synthesis in marine fish: Cloning, functional characterization, and nutritional regulation of fatty acyl Δ6 desaturase of Atlantic cod (Gadus morhua L.). Lipids 41, 1003-1016.
In article      View Article
 
[35]  Carmona-Antoñanzas, G.; Monroig, Ó.; Dick, J. R.; Davie, A.; Tocher, D. R. 2011. Biosynthesis of very long-chain fatty acids (C> 24) in Atlantic salmon: Cloning, functional characterisation, and tissue distribution of an Elovl4 elongase. Comp Biochem Physiol. 159B, 122-129.
In article      View Article  PubMed
 

Published with license by Science and Education Publishing, Copyright © 2018 Hongkuan Zhang, Helu Liu, Dewei Cheng, Hongxing Liu and Huaiping Zheng

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Normal Style
Hongkuan Zhang, Helu Liu, Dewei Cheng, Hongxing Liu, Huaiping Zheng. Molecular Cloning and Functional Characterisation of a Polyunsaturated Fatty Acid Elongase in a Marine Bivalve Crassostrea angulata. Journal of Food and Nutrition Research. Vol. 6, No. 2, 2018, pp 89-95. http://pubs.sciepub.com/jfnr/6/2/4
MLA Style
Zhang, Hongkuan, et al. "Molecular Cloning and Functional Characterisation of a Polyunsaturated Fatty Acid Elongase in a Marine Bivalve Crassostrea angulata." Journal of Food and Nutrition Research 6.2 (2018): 89-95.
APA Style
Zhang, H. , Liu, H. , Cheng, D. , Liu, H. , & Zheng, H. (2018). Molecular Cloning and Functional Characterisation of a Polyunsaturated Fatty Acid Elongase in a Marine Bivalve Crassostrea angulata. Journal of Food and Nutrition Research, 6(2), 89-95.
Chicago Style
Zhang, Hongkuan, Helu Liu, Dewei Cheng, Hongxing Liu, and Huaiping Zheng. "Molecular Cloning and Functional Characterisation of a Polyunsaturated Fatty Acid Elongase in a Marine Bivalve Crassostrea angulata." Journal of Food and Nutrition Research 6, no. 2 (2018): 89-95.
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  • Figure 1. Comparison of the amino acid sequences of PUFA elongases cloning from C. angulata (KY012771) with those from C. nobilis (KF245423), O. vulgaris (AFM93779), Xenopus tropicalis (NP001011248), Siganus canaliculatus (ADE34561), Gallus gallus (NP001184237), and Homo sapiens (NP068586). Identical residues are shaded black and similar residues are gray. The threshold for similarity shading was set at 25%. Indicated is the conserved histidine box motif. Seven (I-VIII) putative membrane-spanning domains (underlined with dashed line) were identified using TMHMM 2.0, five of which (I, II, V, VI, VII) were predicted by Tvrdik et al. (2000)
  • Figure 2. Phylogenetic tree comparing the deduced amino acid sequence of the C. angulata elongase of very long-chain fatty acids with a series of protein sequences including representatives of the ELOVL subtypes from other organisms. A multiple sequence alignment was performed using ClustalW and a phylogenetic tree was made using the neighbor-joining method in MEGA 4.1 package. Numbers represent the frequencies with which the tree topology presented was replicated after 1000 bootstrap iterations
  • Figure 3. Tissue distributions of CaELOVL in C. angulata by qRT-PCR analysis. Expression levels in all tissues are presented relative to β-actin. Vertical bars represent ± S.D. (N= 4) for each tissue
  • Figure 4. Functional characterization of CaELOVL in transgenic S. cerevisiae grown in the presence of fatty acids (FAs). A, B, C, D, E, F and G represent control and adding FA substrate of C18:2n-6, C20:3n-6, C22:3n-6, C20:4n-6, C18:3n-3 and C20:5n-3, respectively. Fatty acids were extracted from yeast transformed with pYES2 vector containing the ORF of the putative fatty acyl elongase cDNA as an insert. Peaks 1-4 represent the main endogenous FAs of S. cerevisiae, namely C16:0 (1), C16:1n-7 (2), C18:0 (3) and C18:1n-9 (4). The remaining main additional peaks (6-17) correspond to the exogenously added FAs and the products of their elongation-C18:2n-6 (7), C20:2n-6 (8), C20:3n-6 (9), C22:3n-6 (10), C22:3n-6 (11), C20:4n-6 (12), C22:4n-6 (13), C18:3n-3 (14), C20:3n-3 (15), C20:5n-3 (16) and C22:5n-3(17). Other minor peaks are 20:1n-9 (5) and 20:1n-7 (6), the latter two resulting from the elongation of 18:1n-9 and 18:1n-7.
[1]  Vance, J. E., Vance, D. E. (Eds.). 2008. Biochemistry of lipids, lipoproteins and membranes. Elsevier.
In article      
 
[2]  Hastings, N., Agaba, M., Tocher, D. R., Leaver, M. J., Dick, J. R., Sargent, J. R., & Teale, A. J. 2001. A vertebrate fatty acid desaturase with Δ5 and Δ6 activities. PNAS USA, 98(25), 14304-14309.
In article      View Article  PubMed
 
[3]  Liu, H., Zheng, H., Wang, S., Wang, Y., Li, S., Liu, W., & Zhang, G. 2013. Cloning and functional characterization of a polyunsaturated fatty acid elongase in a marine bivalve noble scallop Chlamys nobilis Reeve. Aquaculture, 416, 146-151.
In article      View Article
 
[4]  Monroig, O., Guinot, D., Hontoria, F., Tocher, D. R., & Navarro, J. C. 2012. Biosynthesis of essential fatty acids in Octopus vulgaris (Cuvier, 1797): Molecular cloning, functional characterisation and tissue distribution of a fatty acyl elongase. Aquaculture, 360, 45-53.
In article      View Article
 
[5]  Monroig, Ó., Tocher, D. R., & Navarro, J. C. 2013. Biosynthesis of polyunsaturated fatty acids in marine invertebrates: recent advances in molecular mechanisms. Mar Drugs, 11(10), 3998-4018.
In article      View Article  PubMed
 
[6]  Bellou, S., Baeshen, M. N., Elazzazy, A. M., Aggeli, D., Sayegh, F., & Aggelis, G. 2014. Microalgal lipids biochemistry and biotechnological perspectives. Biotechnol Adv, 32(8), 1476-1493.
In article      View Article  PubMed
 
[7]  Maadane, A., Merghoub, N., Ainane, T., El Arroussi, H., Benhima, R., Amzazi, S., & Wahby, I. 2015. Antioxidant activity of some Moroccan marine microalgae: Pufa profiles, carotenoids and phenolic content. J Biotechnol, 215, 13-19.
In article      View Article
 
[8]  Pettersen, A. K., Turchini, G. M., Jahangard, S., Ingram, B. A., & Sherman, C. D. 2010. Effects of different dietary microalgae on survival, growth, settlement and fatty acid composition of blue mussel (Mytilus galloprovincialis) larvae. Aquaculture, 309(1), 115-124.
In article      View Article
 
[9]  Sangsawangchote, S., Chaitanawisuti, N., & Piyatiratitivorakul, S. 2010. Reproductive performance, egg and larval quality and egg fatty acid composition of hatchery-reared Spotted Babylon (Babylonia areolata) broodstock fed natural and formulated diets under hatchery conditions. Int J Fish Aquaculture, 2(1), 044-048.
In article      View Article
 
[10]  Da Costa, F., Nóvoa, S., Ojea, J., & Martínez-Patiño, D. 2011. Changes in biochemical and fatty acid composition of the razor clam Solen marginatus (Solenidae: Bivalvia) during larval development. Mar Biol, 158(8), 1829-1840.
In article      View Article
 
[11]  O'Connor, S., Moltschaniwskyj, N., Bolch, C. J., & O'Connor, W. 2012. Dietary influence on growth and development of flat oyster, Ostrea angasi (Sowerby, 1871), larvae. Aquac Res, 43(9), 1317-1327.
In article      View Article
 
[12]  De Moreno, J. E., Moreno, V. J., & Brenner, R. R. 1976. Lipid metabolism of the yellow clam, Mesodesma mactroides: 2-polyunsaturated fatty acid metabolism. Lipids, 11(7), 561-566.
In article      View Article
 
[13]  Waldock, M. J., & Holland, D. L. 1984. Fatty acid metabolism in young oysters, Crassostrea gigas: polyunsaturated fatty acids. Lipids, 19(5), 332-336.
In article      View Article
 
[14]  Zhukova, N. V. 1991. The pathway of the biosynthesis of non-methylene-interrupted dienoic fatty acids in molluscs. Comp Biochem Physiol. 100B (4), 801-804.
In article      View Article
 
[15]  Zhukova, N. V., & Svetashev, V. I. 1986. Non-methylene-interrupted dienoic fatty acids in molluscs from the sea of Japan. Comp Biochem Physiol. 83B (3), 643-646.
In article      View Article
 
[16]  Liu, H., Zhang, H., Zheng, H., Wang, S., Guo, Z., & Zhang, G. 2014. PUFA biosynthesis pathway in marine scallop Chlamys nobilis Reeve. J Agr Food Chem, 62(51), 12384-12391.
In article      View Article  PubMed
 
[17]  Chu, F. L., & Greaves, J. 1991. Metabolism of palmitic, linoleic, and linolenic acids in adult oysters, Crassostrea virginica. Mar Biol, 110 (2), 229-236.
In article      View Article
 
[18]  Cruz-Romero, M. C., Kerry, J. P., & Kelly, A. L. 2008. Fatty acids, volatile compounds and colour changes in high-pressure-treated oysters (Crassostrea gigas). Innov Food Sci Emerg, 9 (1), 54-61.
In article      View Article
 
[19]  Hastings, N., Agaba, M., Tocher, D. R., Leaver, M. J., Dick, J. R., Sargent, J. R., & Teale, A. J. 2001. A vertebrate fatty acid desaturase with Δ5 and Δ6 activities. P Natl Acad Sci USA, 98(25), 14304-14309.
In article      View Article  PubMed
 
[20]  Tvrdik, P., Westerberg, R., Silve, S., Asadi, A., Jakobsson, A., Cannon, B. & Jacobsson, A. 2000. Role of a new mammalian gene family in the biosynthesis of very long chain fatty acids and sphingolipids. J Cell Biol, 149(3), 707-718.
In article      View Article  PubMed
 
[21]  Agaba, M., Tocher, D. R., Dickson, C. A., Dick, J. R., & Teale, A. J. 2004. Zebrafish cDNA encoding multifunctional fatty acid elongase involved in production of eicosapentaenoic (20: 5n-3) and docosahexaenoic (22: 6n-3) acids. Mar Biotechnol, 6(3), 251-261.
In article      View Article
 
[22]  Morais, S., Monroig, O., Zheng, X., Leaver, M. J., & Tocher, D. R. 2009. Highly unsaturated fatty acid synthesis in Atlantic salmon: characterization of ELOVL5-and ELOVL2-like elongases. Mar Biotechnol, 11(5), 627-639.
In article      View Article  PubMed
 
[23]  Li, Y., Monroig, O., Zhang, L., Wang, S., Zheng, X., Dick, J. R., & Tocher, D. R. 2010. Vertebrate fatty acyl desaturase with Δ4 activity. PNAS USA, 107(39), 16840-16845.
In article      View Article  PubMed
 
[24]  Monroig, Ó., Wang, S., Zhang, L., You, C., Tocher, D. R., & Li, Y. 2012. Elongation of long-chain fatty acids in rabbitfish Siganus canaliculatus: Cloning, functional characterisation and tissue distribution of Elovl5-and Elovl4-like elongases. Aquaculture, 350, 63-70.
In article      View Article
 
[25]  Soriguer, F., Serna, S., Valverde, E., Hernando, J., Martín-Reyes, A., Soriguer, M., & Esteva, I. 1997. Lipid, protein, and calorie content of different Atlantic and Mediterranean fish, shellfish, and molluscs commonly eaten in the south of Spain. Eur J Epidemiol, 13(4), 451-463.
In article      View Article  PubMed
 
[26]  Baby, R. L., Hasan, I., Kabir, K. A., & Naser, M. N. 2010. Nutrient analysis of some commercially important molluscs of Bangladesh. J Sci Res, 2(2), 390-396.
In article      View Article
 
[27]  Liu, H., Zhang, H., Zheng, H., Wang, S., Guo, Z., & Zhang, G. 2014. PUFA biosynthesis pathway in marine scallop Chlamys nobilis Reeve. J Agr Food Chem, 62(51), 12384-12391.
In article      View Article  PubMed
 
[28]  Agaba, M. K., Tocher, D. R., Zheng, X., Dickson, C. A., Dick, J. R., & Teale, A. J. 2005. Cloning and functional characterisation of polyunsaturated fatty acid elongases of marine and freshwater teleost fish. Comp Biochem Physiol. 142B (3), 342-352.
In article      View Article  PubMed
 
[29]  Waldock, M. J., & Holland, D. L. 1984. Fatty acid metabolism in young oysters, Crassostrea gigas: polyunsaturated fatty acids. Lipids, 19(5), 332-336.
In article      View Article
 
[30]  Viso, A.C., Marty, J.C., 1993. Fatty acids from 28 marine microalgae. Phytochemistry 34 (6), 1521-1533.
In article      View Article
 
[31]  Zhukova, N.V., Imbs, A.B., Yi, L.F., 1998. Diet-induced changes in lipid and fatty acid composition of Artemia salina. Comp Biochem. Physiol. 120B (3), 499-506.
In article      View Article
 
[32]  Zheng, X., Seiliez, I., Hastings, N., Tocher, D. R., Panserat, S., Dickson, C. A., & Teale, A. J. 2004. Characterization and comparison of fatty acyl Δ6 desaturase cDNAs from freshwater and marine teleost fish species. Comp Bioch Physiol. Part B, 139(2), 269-279.
In article      View Article  PubMed
 
[33]  Hastings, N.; Agaba, M. K.; Tocher, D. R.; Zheng, X.; Dickson, C. A.; Dick, J. R.; Teale, A. J. 2004. Molecular cloning and functional characterization of fatty acyl desaturase and elongase cDNAs involved in the production of eicosapentaenoic and docosahexaenoic acids from α-linolenic acid in Atlantic salmon (Salmo salar). Mar Biotechnol, 6, 463-474.
In article      View Article
 
[34]  Tocher, D. R.; Zheng, X.; Schlechtriem, C.; Hastings, N.; Dick, J. R.; Teale, A. J. 2006. Highly unsaturated fatty acid synthesis in marine fish: Cloning, functional characterization, and nutritional regulation of fatty acyl Δ6 desaturase of Atlantic cod (Gadus morhua L.). Lipids 41, 1003-1016.
In article      View Article
 
[35]  Carmona-Antoñanzas, G.; Monroig, Ó.; Dick, J. R.; Davie, A.; Tocher, D. R. 2011. Biosynthesis of very long-chain fatty acids (C> 24) in Atlantic salmon: Cloning, functional characterisation, and tissue distribution of an Elovl4 elongase. Comp Biochem Physiol. 159B, 122-129.
In article      View Article  PubMed