Open Access

The codon-optimized Δ6-desaturase gene of Pythium sp. as an empowering tool for engineering n3/n6 polyunsaturated fatty acid biosynthesis

  • Sukanya Jeennor1,
  • Pattsarun Cheawchanlertfa1,
  • Sarinya Suttiwattanakul1,
  • Sarocha Panchanawaporn2,
  • Chanikul Chutrakul2 and
  • Kobkul Laoteng1Email author
BMC Biotechnology201515:82

Received: 10 March 2015

Accepted: 29 August 2015

Published: 15 September 2015



The ∆6-desaturase gene, encoding a key enzyme in the biosynthesis of polyunsaturated fatty acids, has potential in pharmaceutical and nutraceutical applications.


The ∆6-desaturase gene has been isolated from a selected strain of Oomycetes, Pythium sp. BCC53698. The cloned gene (PyDes6) contained an open reading frame (ORF) of 1401 bp encoding 466 amino acid residues. The deduced amino acid sequence shared a high similarity to those of other ∆6-desaturases that contained the signature features of a membrane-bound ∆6-desaturase, including a cytochrome b 5 and three histidine-rich motifs and membrane-spanning regions. Heterologous expression in Saccharomyces cerevisiae showed that monoene, diene and triene fatty acids having ∆9-double bond were substrates for PyDes6. No distinct preference between the n-3 and n-6 polyunsaturated fatty acyl substrates was found. The ∆6-desaturated products were markedly increased by codon optimization of PyDes6.


The codon-optimized ∆6-desaturase gene generated in this study is a promising tool for further reconstitution of the fatty acid profile, in a host system of choice, for the production of economically important fatty acids, particularly the n-3 and n-6 polyunsaturated fatty acids.


Polyunsaturated fatty acids (PUFAs) are important metabolites, which have benefits on human and animal health. Besides being a metabolic fuel, they also play crucial roles in membrane biology and signaling processes in living cells [13]. As a consequence, the demand for PUFAs has continually increased in recent years. Although PUFAs are widely distributed in natural resources, such as plant seed, fungi and marine organisms [4], the search for economical and renewable resources of PUFAs has been extensively persued due to a concern for either an insecurity of supply or healthier performance of PUFA products. Metabolic engineering of the PUFA biosynthetic pathway has been of considerable interest as an alternative approach to produce biomass rich in specific PUFAs [5, 6]. However, this modern technology requires potent genes involved in relevant metabolic pathways.

PUFA biosynthesis is generally associated with a set of membrane-bound enzymes, namely fatty acid desaturase and elongase, which catalyze the introduction of a double bond and the 2-carbon chain extension of fatty acids, respectively. These enzymes have different specificities on fatty acid substrates, relating to acyl chain length and double bond position [7]. ∆6-Desaturase is a key enzyme involved in the biosynthesis of n-3 and n-6 PUFAs, which is responsible for the conversion of essential fatty acids, linoleic acid (LA, C18:2∆9,12) and α-linolenic acid (ALA, C18:3 ∆9,12,15) to γ-linolenic acid (GLA, C18:3∆6,9,12) and stearidonic acid (STA, C18:4∆6,9,12,15), respectively. Subsequently, the desaturated C18 PUFAs can be further metabolized to longer-chain PUFAs, such as arachidonic acid (ARA, C20:4∆5,8,11,14) and eicosapentaenoic acid (EPA, C20:5∆5,8,11,14,17), through alternating series of desaturation and elongation. The highly unsaturated fatty acid, EPA, is one of the nutritionally important n-3 PUFAs, which can be synthesized via either n-3 or n-6 pathways. To manipulate the oil composition of organisms of choice, by a metabolic engineering approach, a high expression level of the ∆6-desaturase enzyme in a heterologous host is required. As well as optimized culture conditions, an efficient promoter and substrate availability, codon optimization is one of the most common approaches for improving heterologous gene expression in some host organisms [813]. Although genes encoding for ∆6-desaturase have been cloned and characterized from various organisms [1420], research on codon optimization has been limited [13].

Very recently we employed Pythium sp. BCC53698 as a genetic resource for the isolation of the ∆6-elongase gene, based on its fatty acid profile [21]. This Oomycete fungus synthesizes ARA and EPA as the end products of n-6 and n-3 PUFAs, respectively, indicating that it would be a potential source for the ∆6-desaturase gene. In this work, we have identified and functionally characterized the ∆6-desaturase gene of Pythium sp. BCC53698. Substrate specificity and preference were investigated by heterologous expression in S. cerevisiae. Codon optimization of the ∆6-desaturase gene was also performed to enhance the product yield.


Identification and characterization of the Pythium6-desaturase gene

The gene coding for ∆6-desaturase was cloned from Pythium sp. using PCR technology. A 700-bp fragment was obtained, and its deduced amino acid sequence showed a high sequence similarity to ∆6-desaturases of other organisms. Inverse PCR and RACE techniques were then performed to obtain the full-length cDNA. Under the optimized PCR conditions (annealing temperature of 55 °C) the product targets approximately 550 and 400 bp in length, were derived from inverse PCR and 5′-RACE, respectively. Taken together, the full-length PyDes6 gene contained an ORF of 1401 bp encoding 466 amino acid residues with a calculated molecular mass of 52.8 kDa. The deduced amino acid sequence of PyDes6 shared the highest homology with Oomycete ∆6-desaturases, which had 68 % identity with the functionally characterized ∆6-desaturase from Phytophthora infestans [22] and Pythium splendens [23].

The PyDes6 sequence contained a conserved characteristic of membrane-bound desaturases, which included three histidine-rich motifs (HXXXH, HXXHH and QXXHH). In addition, the cytochrome b 5-like motif (HPGG) was found at its N-terminus as shown in Additional file 1: Figure S1. Hydropathy analysis of the Pythium desaturase revealed five hydrophobic regions (Fig. 1). All histidine-rich motifs were present in the hydrophilic portion that might be a location at the cytoplasmic surface of the membrane. These features coincide with a model of the topology of membrane-bound desaturases [24]. The phylogenetic tree revealed that PyDES6 belongs to the ∆6-desaturase of Oomycetes (Additional file 1: Figure S2) close to the subgroup of marine algae. This result is in agreement with the fatty acid profile, which is used as a chemotaxonomic marker, showing that Pythium sp. accumulates a n-3 long-chain PUFA (EPA) similar to some marine algae, such as Nannochloropsis oculata [25] and Phaeodactylum tricornutum [26]. These results suggest that this gene encodes a putative ∆6-desaturase, which might be responsible for the introduction of ∆6-double bond into acyl chains.
Fig. 1

Topology model of PyDES6. Enzyme topology was predicted based on TMHMM and Phobius programs. The cylinders represent transmembrane regions of the enzyme, Cytb 5 indicates the N-terminal cytochrom b 5 domain, and three histidine-rich motifs are represented by H1, H2 and H3

Functional analysis of the Pythium6-desaturase

To verify the function of the cloned Pythium desaturase, heterologous expression in S. cerevisiae, under the control of GAL1 promoter, was performed. The transformed yeast cells were supplemented with linoleic acid (C18:2∆9,12, LA) as a fatty acid substrate. After the induction of gene expression fatty acid analysis revealed an extra peak, with the retention time corresponding to γ-linolenic acid (C18:3 ∆6,9,12, GLA), was detected in the yeast transformants carrying the PyDes6, which was absent in the yeast containing empty vector pYES2 (Fig. 2 and Table 1). This result confirmed that PyDes6 encodes for ∆6-desaturase that catalyzes the insertion of double bond into LA yielding GLA.
Fig. 2

Chromatographic profiles of the yeast transformants carrying pYES2 (upper), pPyDes6 (middle) and pMPyDes6 (lower) plasmids. The transgenic yeasts were cultured in the presence of exogenous fatty acid substrates, LA (a) and ALA (b), and the gene expression was induced by addition of 2 % (w/v) galactose

Table 1

Fatty acid composition of yeast transformants containing the empty plasmid (pYES2) and recombinant plasmids (pPyDes6 and pMPyDes6) grown in SD medium supplemented with 50 μM fatty acid substrates

Yeast transformant

Fatty acid composition in total fatty acids (% w/w)







C18:2∆9,12 (LA)

C18:3∆6,9,12 (GLA)

C18:3∆9,12,15 (ALA)

C18:4∆6,9,12,15 (STA)

pYES2 (control)


 No fatty acid addition

22.1 ± 0.4

43.7 ± 0.5


7.5 ± 0.1

26.7 ± 0.2







23.5 ± 0.3

34.3 ± 0.9


8.5 ± 0.1

20.2 ± 0.7


13.6 ± 1.2





22.7 ± 0.3

32.8 ± 0.3


8.7 ± 0.1

20.9 ± 0.6




14.9 ± 0.7




 No fatty acid addition

19.1 ± 1.3

43.7 ± 3.6

1.4 ± 0.1

7.9 ± 0.5

27.8 ± 1.9







22.4 ± 0.5

37.6 ± 0.4

0.2 ± 0.0

7.1 ± 0.2

21.1 ± 0.3


11.2 ± 0.0

0.6 ± 0.0




21.0 ± 2.1

29.9 ± 8.1

0.2 ± 0.1

5.8 ± 2.0

28.9 ± 11.4




13.8 ± 1.0

0.6 ± 0.2



 No fatty acid addition

22.6 ± 0.2

31.9 ± 0.1

11.4 ± 0.2

7.7 ± 0.0

23.3 ± 0.1

3.1 ± 0.0






24.8 ± 0.2

26.6 ± 0.0

5.7 ± 0.3

9.6 ± 0.1

20.8 ± 0.0

1.6 ± 0.0

4.1 ± 0.1

6.9 ± 0.1




24.6 ± 0.3

25.2 ± 0.2

5.9 ± 0.0

10.3 ± 0.1

20.3 ± 0.4

1.6 ± 0.1



4.7 ± 0.3

7.3 ± 0.1

Substrate utilization of PyDes6 was investigated by feeding the yeast transformants with fatty acids with different chain lengths and double bond positions, including saturated and monounsaturated fatty acids and PUFAs. In addition to LA substrate, PyDes6 could catalyze the ∆6-desaturation of n-3 PUFA, α-linolenic acid (C18:3∆9,12,15, ALA), yielding stearidonic acid (C18:4∆6,9,12,15, STA) (Fig. 2 and Table 1). In addition, the ∆6-desaturated 16C fatty acid (C16:2∆6,9) was detectable indicating that the endogenous monoene fatty acid (C16:1∆9,) was a substrate for PyDes6. However, the endogenous and exogenous saturated fatty acids tested were not desaturated. Considering substrate preference, there was not much difference between the conversion rates of LA (5.4 %) and ALA (4.2 %) to the respective ∆6-desaturated products (Table 2).
Table 2

Conversion rate of fatty acyl substrates of yeast transformants containing the recombinant plasmids (pPyDes6 and pMPyDes6)


Conversion rate of substrate to product (%)




3.2 ± 0.0

26.3 ± 0.2



11.8 ± 0.1


5.4 ± 0.0

62.7 ± 0.6


4.2 ± 1.8

60.9 ± 1.3

a nd not detectable

Influence of codon optimization on Pythium6-desaturase activity

Codon usage of the PyDes6 for its expression in fungal system was optimized. Using the OptimumGene™ algorithm, 21.1 % of the 1401-bp coding region was changed, which led to 56.9 % codons being optimized. The GC content was reduced from 64.9 % to 51.9 %. Codon adaptation index (CAI) of the optimized desaturase gene (MPyDes6) was increased from 0.5 to 0.9, which is regarded as good in terms of high gene expression in the desired expression organism. The full-length cDNA of MPyDes6 was ligated to pYES2 vector under the control of GAL1 promoter, generating the pMPyDes6 plasmid. Heterologous expression of the codon-optimized desaturase showed that the MPyDes6 retained the function of ∆6-desaturase, which could convert LA and ALA to GLA and STA, respectively (Fig. 2 and Table 1). It could also convert endogenous monoene fatty acids, C16:1∆9 and C18:1∆9 to hexadecadienoic acid (C16:2∆6,9) and octadecadienoic acid (C18:2∆6,9), respectively, whereas only C16:2∆6,9 was slightly accumulated in the pPyDes6 transformant. Compared with the native enzyme (PyDes6), the conversion of LA to GLA by the MPyDes6 transformant sharply increased from 5.4 to 62.7 %. Similarly, the higher conversion rate of ALA (60.9 %) was found in the transformant carrying the codon-optimized desaturase (Table 2). Thus, the increase of ∆6-desaturated products was a result of codon optimization showing the effective approach for enhancement of the PUFA production in fungal system. The yeast transformants with the empty vector (control), native (PyDes6) and codon-optimized (MPyDes6) genes did not show difference in the cell growth and biomass (Additional file 1: Figure S3).


The biosynthesis of long-chain PUFAs through a series of desaturation and elongation, the ∆6-desaturase has been documented to be a rate-limiting enzyme, and its expression is regulated by several factors [27]. Among ∆6-desaturases from diverse organisms, there is a differentiation in catalytic activity in terms of utilization of acyl substrates that might be a result of genetic variation. The PyDes6 gene identified from this work showed conserved characteristics of membrane-bound desaturases, including a cytochrome b 5 and three histidine-rich motifs, and transmembrane domains [28]. It has been reported that the cytochrome b 5 motif contributes as an electron donor in the electron transport system of fatty acid desaturation by forming the core of a heme binding domain [29]. The histidine-rich motifs are known to be the catalytically essential residues [30].

From the study of substrate specificity, the Pythium enzyme was specific to 16C and 18C fatty acid substrates having ∆9-double bond. Thus, we classified the PyDes6 into the front-end desaturase family, which catalyzes the addition of a double bond between the pre-existing double bond and the carboxyl end of PUFAs [28]. Although the structural and functional characteristics of PyDes6 shared common features of ∆6-desaturases for catalyzing PUFA synthesis, the discrimination in substrate preference was found. Interestingly, both n-3 (ALA) and n-6 (LA) PUFAs having ∆9-double bond were substrates for PyDes6 enzyme at a similar level of substrate conversion rate in contrast to Phytophthora6-desaturase, which prefers LA over ALA [22]. In the plant Primula, ALA was a preferred substrate for the ∆6-desaturase [31].

The synthesis of long-chain PUFAs is derived from either the n-3 or n-6 pathway. The capability of PyDES6 to utilize the n-6 fatty acid (LA) and n-3 fatty acid (ALA) at a similar levels, facilitates the application of this gene for engineering of PUFA pathway of choice in a broad range of organisms (plants and microorganisms). The outcome will depend on substrate availability or the predominant fatty acid accumulated in the host cells. The low proportion of ∆6-desaturated products (GLA and STA) found in the yeast transformant might be a result of a difference in codon usage between Pythium and S. cerevisiae. This could be explained by the evolutionary relationship of the ∆6-desaturase gene derived from Pythium which is closer to marine algae and diatoms than other fungi. This phenomenon is consistent with the taxonomic classification of fungal-like Oomycetes [32].

To enhance the production yields in engineered strain, the optimization of codon usage complemented to a host machinery of target was implemented in this study. The significant increase (P < 0.05) in the substrate conversion rate observed in the yeast harboring codon-optimized gene was presumably derived from an improved translation efficiency in the host system. The increased GLA content obtained by expressing the codon-optimized ∆6-desaturase of Pythium in S. cerevisiae was relatively higher than the yeast cultures carrying ∆6-desaturases of other organisms [31, 33, 34]. These results suggest that codon usage of the host organism had a profound effect on the expression of Pythium enzyme. Additionally, substrate availability is also an important criterion for increasing the production yield of PUFAs, which can be achieved either through genetic or physiological manipulation.

The low lipid content found in S. cerevisiae [35] is seen as a limitation to its use for high yield lipid production on a large scale. Consequently the development of other known oleaginous strains is key to our target in the exploitation of metabolic engineering to develop organisms which deliver high product yield.


This study describes the cloning of a ∆6-desaturase gene from Pythium sp. The gene encoded an enzyme which catalyzed the ∆6-desaturation of the fatty acyl substrates having ∆9-double bond. The product yields were markedly enhanced by codon optimization of the Pythium gene. The redundancy in substrate utilization of the enzyme the codon-optimized gene could be exploited as potential genetic tool for production of nutritionally important PUFA(s) by reconstituting fatty acid profile in biological systems of commercial interest through n-3 or n-6 pathway.


Strains and growth conditions

Pythium sp. BCC53698, deposited in the BIOTEC culture collection (BCC), National Center for Genetic Engineering and Biotechnology, was used as the genetic resource for the isolation of a ∆6-desaturase gene. The fungus was grown in a semi-synthetic medium [36] at 30 °C to logarithmic phase. S. cerevisiae DBY746 (MATα, his3-∆1, leu2-3, leu2-112, ura3-52, trp1-289) was employed as a host for functional analysis of the cloned gene. The yeast was generally grown in a complete medium, YPD (1 % bacto-yeast extract, 2 % bacto-peptone and 2 % glucose). For transformant selection, a minimal medium, SD (0.67 % bacto-yeast nitrogen base without amino acids and 2 % glucose) supplemented with 20 mg/l L-tryptophane, 20 mg/l L-histidine-HCl and 30 mg/l L-leucine was used. Escherichia coli DH5α was used for plasmid propagation.

Nucleic acid manipulation

Genomic DNA from Pythium sp. was extracted from young mycelia (16-hr culture) using the protocol modified from Raeder and Broda [37]. Total RNA was extracted using TRI reagent (Molecular Research Center, Inc., Ohio) according to manufacturer’s instruction. First-strand cDNA was synthesized using P29-oligo-dT-AP primer (Table 1) and SuperScript II first-strand synthesis system (Invitrogen, CA). Then, approximately 50 ng of the first-strand cDNA were further used as templates for 5′-RACE.

Cloning of full-length cDNA of ∆6-desaturase from Pythium sp.

To clone the gene coding for Pythium6-desaturase (PyDes6), a portion of gene was amplified by PCR using a genomic DNA template and degenerate primers, P14-PyDes6-F and P15-PyDes6-R (Table 3), which were designed based on conserved amino acid sequences of ∆6-desaturases of several fungi, F(W/Y)QQSGWLAH and (Q/N)YQ(I/V)(E/D)HHLFP, respectively. The reaction was carried out as follows; an initial denaturation step at 94 °C for 3 mins; followed by 35 cycles at 94 °C for 35 s; primer-specific annealing temperature for 40 s and 72 °C for 1 min; and a final extension step of 72 °C for 5 mins. The expected PCR fragment (700 bp) was subcloned using TOPO TA cloning kit (Invitrogen, CA) following the manufacturer’s instructions. Plasmids were then extracted and purified using the QIAprep mini kit (Qiagen), and sequenced.
Table 3

Oligonucleotide primers used for amplifying ∆6-desaturase gene of Pythium sp.

Primer name

Oligonucleotide sequence (5′-3′)









































aSense and anti-sense strands are denoted by plus (+) and minus (-) symbols, respectively

The inverse PCR and RACE techniques were used for cloning of the full-length PyDes6 gene. According to the derived DNA sequences of the PCR product, six gene-specific primers (Table 3) were designed for amplification of 3′- and 5′-cDNA ends. For inverse PCR, 150 ng of genomic DNA were digested with BamHI (Thermo scientific) in a final reaction volume of 30 μl. The reaction was incubated for about 1–2 h at 37 °C and then inactivated by heating at 80 °C for 15 mins. Digested genomic DNA was purified using the GenepHlow Gel/PCR kit (Geneaid) and circularized by self-ligation with T4 ligase as recommended by the manufacturer (Promega). The reaction was incubated at 4 °C for 16 h [38]. Subsequently, inverse PCR reaction was set by using the self-ligated DNA as a template and the combination of appropriate primer pairs (Table 3) in a final volume of 25 μl. Thermal cycles were as follows; denaturation at 94 °C for 3 mins; followed by 35 cycles of 94 °C for 40 s; 55 °C for 45 s and 72 °C for 2 mins; and the final extension step of 72 °C for 7 mins. To obtain the 5′-end cDNA fragment, the 5′-RACE technique was carried out using 5′-RACE cDNA amplification kit (Invitrogen, CA). PCR was performed using the AAP primer (Table 3) and antisense primer (P89-PyDes6-R). Nested PCR was also conducted to derive a specific product using P91-PyDes6-RN and P96-PyDes6-RN primers. All PCR fragments were purified using the GenepHlow Gel/PCR kit (Geneaid) following the manufacturer’s protocol and were then subcloned into pGEM-T easy vector (Promega, USA) for further sequencing using a service of Macrogen (Korea). The sequences obtained were analyzed against known nucleotide or amino acid sequences available in NCBI GenBank database using the BLAST program ( The cDNA sequence of PyDes6 has been deposited in GenBank and assigned the accession number of KM609327

Structural characterization and phylogenetic analysis

Multiple amino acid alignments of PyDes6 and the ∆6-desaturases of other organisms were performed by using the ClustalW [39] and GeneDoc programs [40]. Transmembrane regions were predicted by the TMHMM algorithm [41] and Phobius [42]. An unrooted phylogenetic tree was constructed based on alignment of amino acid sequences using the neighbour-joining method in MEGA5 software [43]. A total of 1000 bootstrap tests were sampled to determine the confidence in each node on the consensus tree.

Codon optimization of the Pythium6-desaturase

The coding region of the PyDes6 was optimized based on the general rule of RNA stability and the codon usage of the host system. In this study, the OptimumGene TM algorithm was implemented for optimizing a variety of parameters, which are critical to the efficiency of PyDes6 expression in fungal system. The DNA fragment coding for the optimized codon of PyDes6 was synthesized by a service of Genscript (Piscataway, USA). The sequence of MPyDes6 has been deposited in GenBank and assigned the accession number of KT438838. The BamHI and EcoRI sites were incorporated at the start and stop codons, respectively, to further facilitate subcloning into the expression vector, pYES2. The fragment was located downstream of the GAL1 promoter yielding pMPyDes6 plasmid.

Heterologous expression of native and codon-optimized PyDes6 cDNAs in S. cerevisiae

For functional analysis of the native and codon-optimized gene in the heterologous host, the native PyDes6 cDNA fragment was amplified by RT-PCR using high fidelity Taq polymerase (Invitrogen, CA) with the specific primers, P99-PyDes6-BamHI-F and P100-PyDes6-EcoRI-R (Table 3) that contained BamHI or EcoRI sites, respectively, to facilitate subsequent cloning. The amplified product was subcloned into pYES2 expression vector (Invitrogen, CA) downstream of the GAL1 promoter to generate pPyDes6 plasmid. The recombinant plasmids (pPyDes6 and pMPyDes6) and the empty plasmid (pYES2) were then individually transformed to S. cerevisiae using PEG/lithium acetate method following the manufacturer’s protocol (Invitrogen, CA). Transformed cells were selected by uracil prototrophy on SD medium lacking uracil. The yeast transformants were then grown in SD medium containing 2 % (w/v) raffinose and 50 μM of individual fatty acids (Sigma, St. Louis, MO) at 30 °C for 48 h. These fatty acids included saturated fatty acids (myristic acid; C14:0, pentadecanoic acid; 15:0, margaric acid; C17:0, nonadecanoic acid; C19:0, arachidic acid; C20:0 and behenic acid; C22:0), monounsaturated fatty acid (myristoleic acid; C14:1∆9, cis-vaccenic acid; C18:1∆11,cis-eicosenoic acid; C20:1∆11 and erucic acid; C22:1∆13) and PUFAs (LA and ALA). Subsequently, gene expression was induced by adding galactose to a final concentration of 2 % (w/v), and further cultivated for 48 h at 25 °C. Cell growth was determined by spectrophotometry with absorbance at 600 nm (OD600). Cells were harvested by centrifugation, washed twice with 0.1 % tritonX 100, dried to derive a constant weight. Three independent experiments were carried out for each culture. The rate of substrate conversion was calculated as follows; percentage of conversion rate = [product formed/(substrate + product formed)] × 100. Comparison of the substrate conversion rates between the yeast transformants carrying the native and codon-optimized gene was performed to determine the efficiency and substrate preference. The statistical analysis of substrate conversion rates was done using the program SPSS 11.5.

Fatty acid analysis

To determine the fatty acid composition of the yeast transformants, fatty acid methyl esters (FAMEs) were prepared using the method modified from Lepage and Roy [44]. The dried yeast cells were directly transmethylated with 2 ml of 5 % HCl in methanol at 80 °C for 90 mins. After the samples were cooled to room temperature, 1 ml of distilled water and 1 ml of 0.01 % (v/v) butylated hydroxytoluene (BHT) in n-hexane were added and the suspensions were shaken vigorously. The n-hexane phase containing FAMEs was collected and dried under N2 stream. Then, the samples were resuspended in 100 μl hexane and analyzed by gas chromatography using a GC-17A gas chromatograph (Shimadzu, Tokyo) equipped with a capillary column Type OMEGAWAX™250 (Supelco, USA) (30 m × 0.25 μm) and a flame ionization detection. Helium was used as a carrier gas at a constant flow rate of 1.0 mlmin-1. The column and detector temperatures were set at 150–230 °C and 260 °C, respectively. Area measurements of the chromatographic peaks were used to calculate the relative amount of the individual fatty acids. FAMEs were identified by reference to the retention time of FAME standards (Sigma, St. Louis, MO).



α-Linolenic acid


Arachidonic acid


BIOTEC culture collection


Basic local alignment sequencing tool


Codon adaptation index


Eicosapentaenoic acid


Fatty acid methyl esters


Gas chromatography


γ-Linolenic acid


Linoleic acid


Long-chain polyunsaturated fatty acids


Rapid amplification of cDNA ends


Stearidonic acid.



This work was supported by a research grant (P-12-01636) from National Center for Genetic Engineering and Biotechnology (BIOTEC), Thailand. Jeennor, S. and Cheawchanlertfa, P. have an equal contribution on this work. We are grateful to Emeritus Professor John Peberdy, Nottingham University Business School and Dr. Pornkamol Unrean, BIOTEC for language proofreading.

Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (, which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver ( applies to the data made available in this article, unless otherwise stated.

Authors’ Affiliations

Bioprocess Technology Laboratory, Bioresources Technology Unit, National Center for Genetic Engineering and Biotechnology, National Science and Technology Development Agency
Bioassay Laboratory, Bioresources Technology Unit, National Center for Genetic Engineering and Biotechnology, National Science and Technology Development Agency


  1. Fan YY, Chapkin RS. Importance of dietary gamma-linolenic acid in human health and nutrition. J Nutr. 1998;128:1411–4.Google Scholar
  2. Wright S, Burton JL. Oral evening primrose seed oil improves atopic eczema. Lancet. 1982;20:1120–2.Google Scholar
  3. Wang X, Lin H, Gu Y. Multiple roles of dihomo-gamma-linolenic acid against proliferation diseases. Lipids Health Dis. 2012;11:25.Google Scholar
  4. Gunstone FD, Harwood JL, Padley FB. The Lipid Handbook. 2nd ed. London: Chapman & Hill; 1994.Google Scholar
  5. Laoteng K, Certík M, Cheevadhanark S. Mechanisms controlling lipid accumulation and polyunsaturated fatty acid synthesis in oleaginous fungi. Chem Pap. 2011;65:97–103.Google Scholar
  6. Courchesne NM, Parisien A, Wang B, Lan CQ. Enhancement of lipid production using biochemical, genetic and transcription factor engineering approaches. J Biotechnol. 2009;141:31–41.Google Scholar
  7. Leonard A, Pereira S, Sprecher H, Huang Y. Elongation of long-chain fatty acids. Prog Lipids Res. 2004;43:36–54.Google Scholar
  8. Cormack BP, Gertram G, Egerton M, Gow NA, Falkow S, Brown AJ. Yeast-enhance green fluorescent protein (yGFP): a reporter of gene expression in Candida albicans. Microbiology. 1997;143:303–11.Google Scholar
  9. De Rocher EJ, Vargo-Gogola TC, Diehn SH, Green PJ. Direct evidence for rapid degradation of Bacillus thuringiensis toxin mRNA as a cause of poor expression in plants. Plant Physiol. 1998;117:1445–61.Google Scholar
  10. Hung F, Deng L, Ravnikar P, Condon R, Li B, Do L, et al. mRNA stability and antibody production in CHO cells: improvement through gene optimization. Biotechnol J. 2010;5:393–401.Google Scholar
  11. Outchkourov NS, Stiekema WJ, Jongsma MA. Optimization of the expression of equistatin in Pichia pastoris. Protein Expr Purif. 2002;24:18–24.Google Scholar
  12. Tokuoka M, Tanaka M, Ono K, Takagi S, Shintani T, Gomi K. Codon optimization increases steady-state mRNA levels in Aspergillus oryzae heterologous gene expression. Appl Environ Microbiol. 2008;74:6538–46.Google Scholar
  13. Chen Q, Nimal J, Li W, Liu X, Cao W. Delta-6 desaturase from borage converts linoleic acid to gamma-linolenic acid in HEK293 cells. Biochem Biophys Res Commun. 2011;410:484–8.Google Scholar
  14. Sayanova O, Smith MA, Lapinskas P, Stobart AK, Dobson G, Christie WW, et al. Expression of a borage desaturase cDNA containing an N-terminal cytochrome b5 domain results in the accumulation of ∆6-desaturase fatty acids in transgenic tobacco. Proc Natl Acad Sci U S A. 1997;94:4211–6.Google Scholar
  15. Napier JA, Hey SJ, Lacey DJ, Shewry PR. Identification of a Caenorhabditis elegans ∆6-fatty-acid-desaturase by heterologous expression in Saccharomyces cerevisiae. Biochem J. 1998;330:611–4.Google Scholar
  16. Cho PH, Nakamura TM, Clarke DS. Cloning, expression and nutritional regulation of the mammalian ∆6-desaturase. J Biol Chem. 1999;274:471–7.Google Scholar
  17. Huang YS, Chaudhary S, Thurmond JM, Bobik EG, Yuan L, Chan GM, et al. Cloning of ∆12 and ∆6-desaturases from Mortierella alpina and recombinant production of γ-linolenic acid in Saccharomyces cerevisiae. Lipids. 1999;34:649–59.Google Scholar
  18. Zhang Q, Li M, Ma H, Sun Y, Xing L. Identification and characterization of a novel ∆6-fatty acid desaturase gene from Rhizopus arrhizus. FEBS Lett. 2004;556:81–5.Google Scholar
  19. Na-Ranong S, Laoteng K, Kittakoop P, Tantichareon M, Cheevadhanarak S. Substrate specificity and preference of ∆6-desaturase of Mucor rouxii. FEBS Lett. 2005;579:2744–8.Google Scholar
  20. Tan L, Meesapyodsuk D, Qiu X. Molecular analysis of ∆6 desaturase and ∆6 elongase from Conidiobolus obscurus in the biosynthesis of eicosatetraenoic acid, a ω3 fatty acid with nutraceutical potentials. Appl Microbiol Biotechnol. 2011;90:591–601.Google Scholar
  21. Jeennor S, Cheawchanlertfa P, Suttiwattanakul S, Panchanawaporn S, Chutrakul C, Laoteng K. Novel elongase of Pythium sp. with high specificity on ∆6-18C desaturated fatty acid. Biochem Biophys Res Commun. 2014;450:507–12.Google Scholar
  22. Sun Q, Liu J, Zhang Q, Qing X, Dobson G, Li X, et al. Characterization of three novel desaturases involved in the delta-6 desaturation pathways for polyunsaturated fatty acid biosynthesis from Phytophthora infestans. Appl Microbiol Biotechnol. 2013;97:7689–97.Google Scholar
  23. Zhang R, Zhu Y, Ren L, Zhou P, Hu J, Yu L. Identification of a fatty acid ∆6-desaturase gene from the eicosapentaenoic acid-producing fungus Pythium splendens RBB-5. Biotechnol Lett. 2013;35:431–8.Google Scholar
  24. Stukey JE, McDonough VM, Martin CE. The OLE1 gene of Saccharomyces cerevisiae encodes the ∆9-fatty acid desaturase and can be functionally replaced by the rat stearoyl-CoA desaturase gene. J Biol Chem. 1991;265:20144–9.Google Scholar
  25. Roncarati A, Meluzzi A, Acciarri S, Tallarico N, Meloti P. Fatty acid composition of different microalgae strains (Nannochloropsis sp., Nannochloropsis oculata (Droop) Hibberd, Nannochloris atomus Butcher and Isochrysis sp.) according to the culture phase and the carbon dioxide concentration. J World Aquaculture Soc. 2004;35:401–11.Google Scholar
  26. Stiron R, Giusti G, Berland B. Changes in the fatty acid composition of Phaeodactylum tricornutum and Dunuliella tertiolecta during growth and under phosphorus deficiency. Mar Ecol Prog Ser. 1989;55:95–100.Google Scholar
  27. Zhang Q, Li MC, Sun HY, Ma HT, Xing LJ. Progress on molecular biology of delta6-fatty acid desaturases. Sheng Wu Gong Cheng XueBao. 2004;20:319–24.Google Scholar
  28. Meesapyodsuk D, Qiu X. The front-end desaturase: structure, function, evolution and biotechnological use. Lipids. 2012;47:227–37.Google Scholar
  29. Sayanova O, Shewry PR, Napier JA. Histidine-41 of the cytochrome b 5 domain of the borage ∆6-fatty acid desaturase is essential for enzyme activity. Plant Physiol. 1999;121:641–6.Google Scholar
  30. Shanklin J, Whittle E, Fox BG. Eight histidine residues are catalytically essential in a membrane-associated iron enzyme, steroyl-CoA desaturase, and are conserved in alkane hydroxylase and xylene monooygenase. Biochemistry. 1994;33:12787–94.Google Scholar
  31. Sayanova VO, Beaudoin F, Michaelson VL, Shewry RP, Napier JA. Identification of Primula fatty acid ∆6-desaturase with n-3 substrate preference. FEBS Lett. 2003;542:100–4.Google Scholar
  32. Gunderson JH, Elwood H, Ingld A, Kindle K, Sogin ML. Phylogenetic relationships between chlorophytes, chrysophytes, and oomycetes. Proc Natl Acad Sci U S A. 1987;84:5823–7.Google Scholar
  33. Kurdrid P, Subudhi S, Hongsthong A, Ruengjitchatchawalya M, Tanticharoen M. Functional expression of Spirulina-∆6 desaturase gene in yeast, Saccharomyces cerevisiae. Mol Biol Rep. 2005;32:215–26.Google Scholar
  34. Li JN, Chai YR, Zhang XK. Identification and characterization of a novel ∆6-fatty acid desaturase gene from Rhizopus nigricans. Mol Biol Rep. 2009;36:2291–7.Google Scholar
  35. Runguphan W, Keasling JD. Metabolic engineering of Saccharomyces cerevisiae for production of fatty acid-derived biofuels and chemicals. Metab Eng. 2014;21:103–13.Google Scholar
  36. Laoteng K, Jitsue S, Dandusitapunth Y, Cheevadhanarak S. Ethanol-induced changes in expression profiles of cell growth, fatty acid and desaturase genes of Mucor rouxii. Fungal Genet Biol. 2008;45:61–7.Google Scholar
  37. Raeder U, Broda P. Rapid preparation of DNA from filamentous fungi. Lett Appl Microbiol. 1985;1:17–20.Google Scholar
  38. Boulin T, Besserean JL. Mos1-mediated insertional mutagenesis in Caenorhabditis elegans. Nat Protoc. 2007;2:1276–87.Google Scholar
  39. Thompson JD, Higgins DG, Gibson TJ. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res. 1994;22:4673–80.Google Scholar
  40. Nicholas KB, Nicholas Jr HB, Deerfield DW. GeneDoc: analysis and visualization of genetic variation. EMBNET News. 1997;4:1–4.Google Scholar
  41. Krough A, Larsson B, von Heijne G, Sonnhammer EL. Predicting transmembrane protein topology with a Hidden Markov model: application to complete genome. J Mol Biol. 2001;305:567–80.Google Scholar
  42. Kall L, Krough A, Sonnhammer EL. A combined transmembrane topology and signal peptide prediction method. J Mol Biol. 2004;338:1027–36.Google Scholar
  43. Tamura K, Peterson D, Peterson N, Stecher G, Nei M, Kumar S. MEGA5: Molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony. Mol Biol Evol. 2011;28:2731–9.Google Scholar
  44. Lepage G, Roy CC. Improved recovery of fatty acid through direct transesterification without prior extraction or purification. J Lipid Res. 1984;25:1391–5.Google Scholar


© Jeennor et al. 2015