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. 2025 Dec 11;12:172–182. doi: 10.1016/j.synbio.2025.11.013

Enhancing delta-tocotrienol production in Saccharomyces cerevisiae via metabolic engineering strategies in conjunction with the mutagenesis of tocopherol cyclase

Ziming Liu a,b,c,d,1, Min Tang a,b,c,d,1, Wanze Zhang a,b,c,d,1, Yanjie Tian a,b,c,d, Jianjun Qiao a,b,c,d, Mingzhang Wen a,b,c,d, Weiguo Li a,b,c,d,⁎, Qinggele Caiyin a,b,c,d,⁎⁎
PMCID: PMC12754398  PMID: 41480008

Abstract

Delta (δ)-tocotrienol is a member of the vitamin E family and exhibits bioactivities such as antioxidant, anti-inflammatory, and neuroprotective activities. As a nutrient with protective effects on human health, δ-tocotrienol has broad application prospects in food, cosmetic, and pharmaceutical industries. The construction of efficient microbial cell factories capable of δ-tocotrienol production using synthetic biology approaches is an effective strategy for supplementing or even replacing the vitamin E supply chain in the future. The current study successfully enhanced the biosynthesis of δ-tocotrienol in Saccharomyces cerevisiae by combining metabolic engineering and enzyme engineering strategies. Specifically, the substrate channel constructed by the sequential fusion of the enzymes PaCrtE and SyHPT successfully increased the supply of the key precursor MGGBQ, resulting in a significant increase in the production of δ-tocotrienol. In situ extraction and optimization of the expression of transporter protein PDR1 increased the efflux of δ-tocotrienol, directing the metabolic flux toward the product δ-tocotrienol. To enhance the catalytic activity of the key rate-limiting enzyme tocopherol cyclase from Arabidopsis thaliana (AtTC), semirational protein design was conducted herein. The mutant AtTCT87S was found to increase the production of δ-tocotrienol by 2.3 times compared to that obtained with the wild-type enzyme. AtTCT87S can thus be universally used for synthetic biology strategies in future studies to enhance the microbial heterologous production of δ-tocotrienol. The strain T08 was finally obtained herein; the numerous metabolic engineering strategies discussed in this study were integrated into this strain, allowing the production of 4337.3 μg/L of δ-tocotrienol in a shake-flask fermentation, which is 8.9 times that of the yield obtained with the initial strain T03. Scaling up to a 5-L fermentation tank resulted in a δ-tocotrienol yield of 16.9 mg/L.

Keywords: δ-tocotrienol, Vitamin E, Tocopherol cyclase, Microbial cell factories, Saccharomyces cerevisiae

1. Introduction

Vitamin E is a golden or yellow oily material, which was discovered in green leafy vegetables by Evans and Bishop (1922) [1]. Vitamin E is a fat-soluble vitamin and an essential nutrient for humans. It is mainly found in the leaves and seeds of plants, as well as in eggs and poultry [2]. Vitamin E represents a group of eight compounds, including alpha (α)-, beta (β)-, gamma (γ)-, and delta (δ)-tocopherols along with α, β, γ, and δ-tocotrienols (Fig. 1a). Both tocopherol and tocotrienol contain a hydrophilic chromophore ring and a hydrophobic isopentenyl side chain. The isopentenyl side chain of tocopherol is completely saturated, while that of tocotrienol has three double bonds [3]. The commercial production of vitamin E globally is over 75,000 tons per year currently [4] and is chiefly reliant on plant extraction as well as chemical, microbial, and biochemical methods of synthesis. Approximately 80 % of the commercially available vitamin E is produced via chemical synthesis [5]. The chemical synthesis of vitamin E side chains involves the steps of catalytic hydrogenation and proton acid catalysis, which are dependent on the utilization of specific equipments. Moreover, the synthesis route for the main ring of vitamin E is limited by the high cost of raw materials. Therefore, the development of a green, environmentally friendly, and cost-effective method of vitamin E synthesis is of tremendous importance. A combination of biological and chemical routes allows for a relatively rapid synthesis of vitamin E while circumventing the disadvantages associated with its chemical synthesis. The synthesis and secretion of farnesene in microorganisms was achieved in 2014, followed by in vitro chemical synthesis processes to generate the vitamin E precursor isophytol [5]. This method had provided a convenient alternative to the traditional synthesis method of isophytol and revolutionized the production of vitamin E.

Fig. 1.

Fig. 1

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Chemical structure (a) and biosynthetic pathway (b) of δ-tocotrienol.

Tocopherol represents the main synthetic configuration of vitamin E presently in the market. Tocotrienol is another subtype of vitamin E with physiological functions characteristic of vitamin E, including antioxidant, anti-inflammatory, anticancer, and neuroprotective activities [6]. The current study focuses on δ-tocotrienol and its two biosynthetic pathways, the mevalonate (MVA) and shikimic acid pathways. Geranyl geranyl pyrophosphate (GGPP), the isopentenyl side chain of δ-tocotrienol, is synthesized via the MVA pathway. Similarly, the aromatic receptor homogentisic acid (HGA) is synthesized from 4-hydroxyphenylpyruvate (4-HPP) generated via the shikimic acid pathway in a reaction catalyzed by 4-hydroxyphenylpyruvate dioxygenase (HPPD). The conversion of GGPP and HGA to the intermediate 2-methyl-6-geranylgeranyl benzoquinone (MGGBQ) is catalyzed by homogentisate geranylgeranyl transferase (HGGT); the conversion of MGGBQ to δ-tocotrienol is then catalyzed by tocopherol cyclase (TC) [7], as shown in Fig. 1b. Interestingly, the heterologous production of δ-tocotrienol in microorganisms has now been achieved. Albermann et al. were the first to express the entire synthesis pathway of δ-tocotrienol in Escherichia coli and successfully achieve its heterologous synthesis, obtaining δ-tocotrienol yields of 15 μg/g DCW [8]. The low levels of GGPP synthesis in E. coli limit options for increasing the yield of δ-tocotrienols in this system. Saccharomyces cerevisiae has proven to be an excellent system for the study of terpenes and aromatic compounds, and its tolerance to isoprene compounds such as farnesyl pyrophosphate (FPP) and GGPP is higher than that of E. coli [[9], [10], [11]]. Moreover, the eukaryotic expression system of S. cerevisiae may be more suitable for the expression of plant-derived enzymes. Sun et al. first expressed heterologous genes encoding HPPD, homogentisate phytyltransferase (HPT), and TC at a multicopy rDNA loci in S. cerevisiae in conjunction with enhancing the MVA pathway; the optimization of fermentation conditions using response surface methodology allowed them to ultimately obtain δ-tocotrienol yields of 4.10 ± 0.10 mg/L [12]. Subsequently, Jiao et al. achieved heterologous synthesis of tocotrienols in S. cerevisiae by optimizing the biosynthesis pathway and performing metabolic engineering of product efflux [13]. In another study, Jiao et al. achieved the heterologous synthesis of a single configuration of δ-tocotrienol in S. cerevisiae. Specifically, they enhanced the synthesis pathways of the precursor molecules GGPP and HGA by optimizing the rate limiting steps, weakening the competing pathways, and mitigating feedback inhibition. The optimal copy number ratio of the heterologous genes encoding TC, HPT, and HPPD was adjusted to 2:3:1, and the transporter protein drug-responsive transcription factor (PDR1) was overexpressed herein. In addition, olive oil and 2-hydroxypropyl-β-cyclodextrin were added during fermentation to increase the efflux of δ-tocotrienol, resulting in a final yield of 241.7 mg/L and 85.6 % efflux of δ-tocotrienol into the extracellular environment [14]. Han et al. constructed a substrate channel via the fusion of the three heterologous genes HPPD, HPT, and truncated TC (tTC) using linker and scaffold proteins to improve the catalytic efficiency of the key enzymes for the biosynthesis of tocotrienols. Moreover, computer simulation was employed for modifying tTC to enhance its activity, resulting in a δ-tocotrienol yield of 3262.2 μg/L [15]. Recently, Xiang et al. synthesized δ-tocotrienol in Yarrowia lipolytica and achieved a yield of 466.8 mg/L in a 5-L bioreactor by carrying out direct fusion of the gene encoding SyHPT (from Synechocystis sp. PCC 6803) with tocopherol cyclase from Arabidopsis thaliana (AtTC) and conducting a semirational design on SyHPT [16].

Despite the remarkable achievements reported in previous studies, the catalytic activities of enzymes involved in δ-tocotrienol biosynthesis and the supply of precursors continue to pose limitations in its production. The key rate limiting nodes in the biosynthesis pathway of δ-tocotrienol were modified herein. Specifically, a fusion protein of CrtE and SyHPT was constructed to increase the catalytic efficiency of the enzyme through substrate channel effects. The efflux of δ-tocotrienol was increased via biphasic fermentation and overexpression of the transporter protein PDR1, leading to increased metabolic flux toward the product. Additionally, semirational design was applied for the key rate limiting enzyme TC; the nonconserved amino acids identified by sequence alignment with other tocopherol cyclase were mutated to obtain a mutant with higher catalytic activity, which can be universally employed for synthetic biology approaches in future studies on the microbial heterologous production of δ-tocotrienol. The abovementioned strategies used herein increased the yield of δ-tocotrienol and lay the groundwork for future research to increase the yield of δ-tocotrienol.

2. Materials and methods

2.1. Strains, media, and chemical reagents

E. coli DH5α (Tsingke Biotechnology, China) was used for plasmid construction. The bacterial cells were cultured in Luria–Bertani (LB) medium (10 g/L tryptone, 5 g/L yeast extract, and 10 g/L sodium chloride). S. cerevisiae Sc027 [17] was used as the initial strain. All yeast strains used in this study are listed in Table S1. Yeast extract peptone dextrose (YPD; 20 g/L glucose, 20 g/L tryptone, and 10 g/L yeast extract) and synthetic dropout (SD; 20 g/L glucose, 6.7 g/L yeast nitrogen base, and 2 g/L amino acid dropout mixed powder) media were employed for the culture of yeast cells and selection of recombinant yeast strains.

Polymerase chain reaction (PCR) amplification was carried out using 2 × Phanta Max Master Mix and 2 × Rapid Taq Master Mix from Vazyme Biotech, China. The amplicons so obtained were purified using the Universal DNA Purification Kit from Tiangen Biotech, China. Plasmid extraction was performed using the TIANprep Mini Plasmid Kit (Tiangen Biotech). All the chemicals, including analytical standard HGA and δ-tocotrienol, were purchased from Beijing Solarbio Science & Technology Co., Ltd. (China), unless stated otherwise. All the primers and codon-optimized heterologous genes were synthesized by Genewiz, China.

2.2. Construction of plasmids and strains

All the plasmids used in this study are listed in Table S2. The genes encoding PpHPPD from Pseudomonas putida, AtHPPD from Arabidopsis thaliana, the GGPP synthase PaCrtE from Pantoea ananatis, SyHPT from Synechocystis sp. PCC 6803, and four TCs, including AhTC from Arachis hypogaea, TaTC from Triticum aestivum, GmTC from Glycine max, and PdTC from Prunus dulcis, were codon optimized and synthesized by Genewiz, China. All the abovementioned genes were inserted in the multiple cloning site of plasmid pESC-URA under the bidirectional promoter pGAL1-pGAL10 for overexpression in S. cerevisiae strains.

For the site-directed mutagenesis of AtTC, linearized plasmids containing the desired mutations were obtained via the PCR amplification of the template plasmid pESC-URA- AtTC using primers containing the desired mutations. The linearized plasmids were subsequently recircularized using ClonExpress Ultra One Step Cloning Kit-C115 from Vazyme Biotech, China. The primers employed herein have been listed in Table S3. Genomic DNA from S. cerevisiae CEN.pK2-1C was used for the PCR amplification of endogenous genes. The templates for homologous recombination included homologous arms flanking the desired inserting site and gene expression cassettes (including the promoters, open reading frame, and terminator sequences). These sequences were amplified by PCR and purified to yield the initial fragments. All the adjacent basic fragments sharing homologous sequences of 40–60 bp were employed for overlap extension PCR and subsequent homologous recombination. The adjacent 2–4 initial fragments were assembled using fusion PCR. The primers employed herein have been listed in Table S3. The fragments obtained by fusion PCR were purified, quantified, and employed for the transformation of yeast cells and subsequent integration into its chromosome [18]. Yeast transformation was carried out following the lithium acetate protocol [19].

2.3. Fermentation conditions

A two-phase fermentation method was used herein for producing δ-tocotrienol. For shake-flask fermentation, a single colony of the transformants was picked up from YPD or SD agar plates and used for the inoculation of YPD or SD medium (3 mL) in 10-mL culture tubes; the cultures were incubated at 30 °C under rotary conditions (200 rpm) for 20 h until the exponential phase of growth was reached. Then, the preculture was used for the inoculation of 50 mL of SD-URA or YPD media such that an initial optical density at 600 nm (OD600) of 0.05 was attained. The cultures were subsequently grown for 96 h. After 10 h, 1 % (w/v) final concentration of d-galactose and 5 mL of the extractant (n-dodecane) were added.

A 5-L bioreactor was used herein for fed-batch fermentation using 2 L of YPD medium supplemented with solutions of vitamins and trace metals. For preparing a fresh seed culture, the strains stored in −80 °C were revived on a YPD agar plate. For obtaining the seed culture, the cells were first grown in 30 mL of YPD medium followed by subculturing in 200 mL of YPD medium and incubation at 30 °C for 18 h under rotary conditions (220 rpm). At the start of fermentation, the seed culture was employed for the inoculation (10 % v/v) of the fermentation medium in 5-L bioreactor. The pH, temperature, and dissolved oxygen levels were controlled at 5.50, 30 °C, and above 30 %, respectively. Glucose and ethanol were used as carbon sources. The glucose feeding rate was adjusted such that the concentration of residual glucose remained below 3 g/L. The ethanol concentration was maintained below 5 g/L until the end of fermentation.

2.4. Measurements of biomass and metabolite concentrations

The biomass (OD600 of the cells) was determined using an ultraviolet-1800PC spectrophotometer (Shanghai MAPADA Instruments Co., Ltd., China). The contents of residual glucose and ethanol were measured using an SBA-90 biosensor.

For HGA analysis, 2 mL of the cell culture after fermentation was centrifuged at 12,000 rpm for 5 min to promote phase separation. The supernatant was collected, and 40 μL of glacial acetic acid was added to it. The mixture was then filtered using 0.22-μm polyethersulfone syringe filters for high performance liquid chromatography (HPLC) analysis. The HPLC analysis for HGA was performed using the 1260 Infinity II Prime Online LC System. Chromatographic separation of HGA was carried out at 30 °C using an Agilent InfinityLab Poroshell 120 EC-C18 column. Elution was carried out using a mixture consisting of 90 % (v/v) 0.01 M monopotassium phosphate (A) and 10 % methanol (B) at a flow rate of 0.8 mL/min, followed by UV detection at a wavelength of 290 nm.

Following its production and enrichment in cells, the fat-soluble δ-tocotrienol needs to be extracted from the cells and subsequently detected. For HPLC analysis, 30 mL of the cell culture was collected and centrifuged at 8000 rpm for 5 min. The supernatant was discarded, and the cells were washed twice with distilled water and centrifuged after each wash, with the supernatant discarded each time. Then, 1 mL of ethyl acetate and 3 g of quartz sand (16–30 mesh) were added to the 10-mL centrifuge tube containing yeast cells. The mixture was shaken in a vortex mixer for 30 min and then cooled on ice for 10 min; this process was carried out for a total of two times. Subsequently, the centrifuge tube containing the mixture was subjected to centrifugation at 12,000 rpm for 5 min. Then, the supernatant was collected and filtered using 0.22-μm nylon syringe filters for subsequent analyses. The HPLC analysis of δ-tocotrienol was carried out using the same device employed for that of HGA. Elution was carried out using acetonitrile (A) and water (B) at a flow rate of 0.8 mL/min, followed by UV detection at a wavelength of 292 nm. Gradient elution was carried out under the following conditions: 70 % A/30 % B to 90 % A/10 % B (0–10 min), 90 % A/10 % B to 100 % A/0 % B (10–40 min), 100 % A/0 % B (40–70 min), and 100 % A/0 % B to 70 % A/30 % B (70–80 min).

2.5. Molecular docking analysis and multiple sequence alignments

Clustal X (http://www.clustal.org) and WebLogo (https://weblogo.threeplusone.com) were used for multiple sequence alignments. The protein structures of AtTC and its mutants were predicted using AlphaFold 3 (https://alphafoldserver.com). The three-dimensional structure of MGGBQ (PubChem CID 14759579) was downloaded from the website https://pubchem.ncbi.nlm.nih.gov. The models were then docked with the substrate MGGBQ using AutoDock Vina.

3. Results and discussion

3.1. Screening of key enzymes and generation of S. cerevisiae strains harboring the δ-tocotrienol biosynthesis pathway

GGPP and HGA are the two main precursors for the synthesis of δ-tocotrienol, and a supply of GGPP and HGA should be established for the production of δ-tocotrienol. In yeast, GGPP can be synthesized from FPP by the farnesyltranstransferase BTS1, while HGA is synthesized from the product 4-HPP of the shikimic acid pathway. The S. cerevisiae strain Sc027 employed herein allows for the enhanced expression of enzymes of the MVA pathway using the Gal promoters [17]. For ensuring a supply of HGA, tyrosine was added to the culture medium; this amino acid is utilized by the shikimic acid pathway in yeast to generate 4-HPP, which can then be converted to HGA upon the overexpression of the gene encoding HPPD.

Previous studies suggest that heterologous expression of HPPD from two sources can be achieved in S. cerevisiae; these include PpHPPD from P. putida and AtHPPD from A. thaliana. To compare the ability of the two HPPDs to catalyze the synthesis of HGA in the strain Sc027, the plasmids pESC-Ura-PpHPPD and pESC-Ura-AtHPPD were constructed. These plasmids can be expressed at high copy numbers in S. cerevisiae and used for the transformation of the strain Sc027. The strain Sc027 harboring either of the two plasmids (pESC-Ura-PpHPPD or pESC-Ura-AtHPPD) was cultured in 50 mL of SD-URA liquid medium containing 0.1 % (W/V) tyrosine for 96 h. After fermentation, the products were extracted from the fermentation broth and analyzed using HPLC. A comparison of the positions of the peaks in the sample from the fermentation broth with those in the HGA standard revealed that both PpHPPD and AtHPPD can catalyze the production of HGA in the strain Sc027 (Fig. 2a). Subsequent calculations revealed that the yield of HGA generated by PpHPPD was approximately 6.1 mg/L, which was 1.4 times higher than that by AtHPPD (Fig. 2b). Thus, the performance of PpHPPD was found to be superior to that of AtHPPD in the strain Sc027; PpHPPD was therefore selected for use in subsequent experiments for catalyzing the synthesis of HGA and constructing the biosynthesis pathway for δ-tocotrienol in yeast.

Fig. 2.

Fig. 2

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Construction of the biosynthetic pathway of δ-tocotrienol in S. cerevisiae. (a) Comparison of HPLC chromatograms between HGA standard and S. cerevisiae strains expressing PpHPPD or AtHPPD. (b) HGA production of S. cerevisiae strains expressing PpHPPD or AtHPPD in plasmids. (c) Comparison of HPLC chromatograms between δ-tocotrienol standard and S. cerevisiae strains expressing genes of TCs from different sources. (d) δ-tocotrienol production of S. cerevisiae strains expressing genes of TCs from different sources in plasmids.

SyHPT from Synechocystis sp. PCC 6803, which has shown promising results in previous studies [14,16], was selected herein for catalyzing the synthesis of MGGBQ from GGPP and HGA. In addition, the endogenous farnesyltranstransferase BTS1 catalyzing the synthesis of GGPP in S. cerevisiae has a low catalytic efficiency due to its high Km for FPP and low turnover number [20,21], which limits the subsequent synthesis of δ-tocotrienol. Therefore, the GGPP synthase PaCrtE from P. ananatis was employed herein for enhancing the supply of GGPP in Sc027 [22,23]. The expression cassette constructed with PaCrtE, SyHPT, and PpHPPD was inserted into the MET17 locus of the Sc027 genome, and the resulting strain (confirmed upon sequencing) was named T01.

TC is the final enzyme of the biosynthesis pathway and catalyzes MGGBQ cyclization to produce δ-tocotrienol. Moreover, it catalyzes one of the key rate-limiting steps in the δ-tocotrienol biosynthesis pathway. To increase δ-tocotrienol production, TCs from different sources were screened in yeast with the hope of identifying an enzyme with higher catalytic activity. Four TCs from plants with high vitamin E content were screened herein, including AhTC, TaTC, GmTC, and PdTC [24]. Together with AtTC, the five TCs were individually cloned into the yeast expression plasmid pESC-Ura to yield the plasmids pESC-Ura-AhTC, pESC-Ura-TaTC, pESC-Ura-GmTC, pESC-Ura-PdTC, and pESC-Ura-AtTC, respectively. These plasmids were then employed for the transformation of the strain T01. The obtained strains were cultured in 50 mL SD-URA medium containing 0.1 % (W/V) tyrosine for 96 h. After fermentation, the intracellular products were extracted and analyzed using HPLC to compare the catalytic performance of the five TCs. A comparison of the HPLC chromatogram of these samples with that of the δ-tocotrienol standard revealed that AtTC, AhTC, PdTC, and GmTC could catalyze MGGBQ cyclization to generate δ-tocotrienol in the yeast strain T01 (Fig. 2c); however, δ-tocotrienol production was undetectable in the strain expressing TaTC. Moreover, δ-tocotrienol production demonstrated the catalytic activities of PaCrtE and SyHPT in strain T01. In terms of δ-tocotrienol production, AtTC showed optimal performance in the T01 strain, with a δ-tocotrienol yield of 679.2 μg/L, which is 3.2, 25.2, and 12.2 times that of AhTC, PdTC, and GmTC, respectively (Fig. 2d). Therefore, AtTC was selected for use in subsequent experiments.

The use of plasmids as vector for gene expression in brewing yeast is known to result in plasmid loss and genetic instability [25]. Therefore, chromosomal integration of the genes encoding the rate-limiting enzymes AtTC and SyHPT of the δ-tocotrienol biosynthesis pathway was conducted herein to support sustained production. DPP1 was initially selected as the insertion site for the genes encoding AtTC and SyHPT. Accordingly, the expression cassette AtTC-PGal10-PGal1-SyHPT constructed herein was inserted into the DPP1 locus of the T01 genome. The strain so obtained was verified by sequencing and named as T02. Following fermentation with the strain T02 for 96 h, the intracellular products were extracted and analyzed using HPLC; however, δ-tocotrienol was not detected. We speculate that the low expression intensity of the gene AtTC is insufficient to support the production of adequate levels of δ-tocotrienol to meet the HPLC detection limit. This result suggests that a further increase in the copy number of key genes associated with the biosynthesis pathway is required.

The insertion of genes into the multicopy sites of S. cerevisiae can lead to higher levels of gene expression, thereby improving the overall efficiency of catalysis by the encoded enzymes. For example, Qi et al. integrated genes encoding the enzymes p-coumaric acid 3-hydroxylase and cytochrome P450 reductase one of the caffeic acid biosynthesis pathway into the δ site of S. cerevisiae, resulting in a 50-fold increase in caffeic acid production [26]. Peng et al. introduced HapAmp, a novel methodology that utilizes haploinsufficiency as an evolutionary force to drive gene amplification. This technique facilitates efficient, titratable, and stable integration of up to 47 copies of heterologous genes into the yeast genome, as exemplified by the significantly increased production of nerolidol [27]. Herein, RPL25 was selected as the gene integration site for driving the spontaneous gene replication. The expression cassette AtTC-PGal10-PGal1-SyHPT constructed herein (Fig. 3a) was used for the transformation of strain T01, resulting in six strains that were verified by sequencing; these were named strains R01–R06. Following fermentation in 50 mL YPD medium containing 0.1 % (W/V) tyrosine, all the strains produced sufficient quantities of δ-tocotrienol to allow detection using HPLC; the results are shown in Fig. 3b and Fig. S1. The highest yield of δ-tocotrienol (488.9 μg/L) was produced by the strain R05, which is hereinafter referred to as strain T03. As a yeast strain harboring the complete pathway for δ-tocopherol biosynthesis, the strain T03 was employed for subsequent research.

Fig. 3.

Fig. 3

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The spontaneous gene replication of AtTC and SyHPT in RPL25 site to increase the expression level of key enzymes. (a) The chromosome integration mode of AtTC and SyHPT at RPL25 site. (b) δ-tocotrienol production of so obtained strains R01–R06.

3.2. Enhancing precursor supply and expressing the fusion protein PaCrtE–SyHPT enhances the production of δ-tocotrienol

Enhancing the supply of δ-tocotrienol precursors in the biosynthesis pathway is a feasible and effective means of enhancing its production. GGPP is one such important precursor. In S. cerevisiae, GGPP is synthesized via the MVA pathway, which begins with acetyl coenzyme A (acetyl CoA). Previous studies have demonstrated that enhancing the production of acetyl CoA can effectively increase the yield of downstream products such as lycopene [23,28]. The pathway for the generation of acetyl CoA in the cytoplasm of S. cerevisiae involves the production of acetate via the activity of the enzyme acetaldehyde dehydrogenase ALD6; the subsequent conversion of acetate to acetyl CoA is catalyzed by acetyl CoA synthase ACS1 and ACS2 [29]. When glucose is employed as the carbon source for S. cerevisiae, the glycolytic flux is channeled toward ethanol synthesis. Herein, the metabolic flux is redirected from ethanol synthesis to that of acetyl CoA via the overexpression of ALD6, SeACS derived from Salmonella enterica [30], and the alcohol dehydrogenase ADH2. Furthermore, an additional GGPP synthase PaCrtE was also overexpressed herein.

The constructed expression cassette containing the genes encoding ALD6, SeACS, ADH2, and PaCrtE was integrated into the YPL062W locus of strain T03; the resulting strain was verified upon sequencing and named T04. After 96 h of fermentation in 50 mL YPD liquid medium supplemented with 0.1 % (W/V) tyrosine, the intracellular product of T04 was extracted for HPLC analysis. The yield of δ-tocotrienol was found to be 679.5 μg/L, which is 1.4 times higher than that from strain T03 (Fig. 4a).

Fig. 4.

Fig. 4

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Enhancing precursor supply by application of metabolic engineering strategies to increase δ-tocotrienol production. (a) Overexpression of genes ALD6, SeACS, ADH2, and PaCrtE to increase the supply of acetyl CoA and GGPP leads to an increase in the production of δ-tocotrienol. (b) Increasing the supply of MGGBQ by constructing a fusion protein of PaCrtE and SyHPT leads to an increase in the production of δ-tocotrienol.

The overall efficiency of metabolic pathways may be improved by assembling the target enzymes using enzyme complexes, which can reduce the diffusion-mediated loss of metabolic intermediates, thereby increasing product yield. An effective strategy involves the use of fusion linkers, which not only prevent protein misfolding but also connect two enzymes involved in successive catalytic reactions, thereby accelerating the reaction between intermediates and the successive enzymes in the pathway [31,32]. For example, Liu et al. confirmed that the use of a flexible linker (GGGS) and optimization of the protein order of neryl pyrophosphate synthase (NPPS) and limonene synthase (LS) to NPPS-LS increased the limonene titer by 5.5 times compared to that obtained with the LS-NPPS group [33]. Additionally, achieving enzyme fusion via linker fusion may enhance enzyme stability, thereby increasing product yield. For example, Cheah et al. fused FPP synthase (FPPS) and nerolidol synthase (NES) to increase nerolidol yield by about 110 times owing to the enhancement in NES stability [34]. In the current study, the GGPP synthase PaCrtE was fused with SyHPT using a linker to achieve a potential increase in the production of δ-tocotrienol. The linkers used for protein fusion can be divided into rigid and flexible types, and the type of linker and order of enzyme in the fusion protein are crucial for product yield. Shu et al. showed that the effect of reverse fusion of ERG20F96W/N127W with tMS is better than that of the sequential fusion [35]. Ren et al. screened the rigidity/flexibility and length of the linker and found that the flexible linker (G)8 supported the best enzyme performance [36]. Using the strain L01 constructed herein by transforming the strain T03 with the coexpression plasmid pESC-URA-PaCrtE-SyHPT expressing both PaCrtE and SyHPT as a control, the fusion sequence of PaCrtE and SyHPT was first determined by selecting a short flexible linker GSG. The constructed plasmids pESC-URA-PaCrtE(GSG)SyHPT and pESC-URA-SyHPT(GSG)PaCrtE carrying the fusion enzymes were employed for the transformation of the strain T03 to obtain strains L02 and L03, respectively. After 96 h of shake-flask fermentation in 50 mL SD-URA medium supplemented with 0.1 % (W/V) tyrosine, the intracellular products were extracted for HPLC analysis. The results showed that strain L02 with the fusion enzyme PaCrtE(GSG)SyHPT exhibited a 1.4-fold increase in the production of δ-tocotrienol compared to that of the control strain L01 (Fig. 4b). By contrast, the production of δ-tocotrienol decreased in the strain L03 with the fusion enzyme SyHPT(GSG)PaCrtE, indicating that the sequential fusion of PaCrtE and SyHPT had a positive effect on δ-tocotrienol production.

After determining the fusion sequence of the enzymes PaCrtE and SyHPT, the length and type of fusion linker were screened. The flexible linkers GGGGS, (GGGGS)2, and (GGGGS)3 as well as the rigid connectors EAAAK, (EAAAK)2 [37], and (PT)4P [38] were selected for the subsequent fusion of PaCrtE and SyHPT. The constructed plasmids pESC-URA-PaCrtE(linker)SyHPT were employed for the transformation of the strain T03, thereby generating the strains L02-1 to L02-6, respectively. The strains so obtained were cultured and employed for fermentation in 50 mL SD-URA medium supplemented with 0.1 % (W/V) tyrosine for 96 h. Subsequently, the intracellular products were extracted and subjected to HPLC analysis. In general, the flexible linkers GGGGS and (GGGGS)2 were found to be superior to the rigid linkers; however, the long distance between the fused proteins arising from the use of an excessively long flexible linker (GGGGS)3 may result in little improvement in catalytic efficiency, which is consistent with the observation herein that short linkers are generally superior to the long linkers (Fig. 4b). Taken together, the results suggest that GSG is the most effective linker (Fig. 4b).

3.3. Enhanced efflux of δ-tocotrienol increases production

Vitamin E consists of a hydrophilic head and a hydrophobic tail, and previous studies have demonstrated its localization on the cell membranes [39]. Herein, δ-tocotrienol was only detectable following the collection and fragmentation of cells and not in the fermentation broth. This observation is in alignment with the results of previous studies. For instance, Jiao et al. revealed that the overexpression of the endogenous adenosine triphosphate (ATP)–binding cassette (ABC) transporter PDR1 in yeast in conjunction with biphasic fermentation and the addition of 2-hydroxypropyl-beta-cyclodextrin promoted the efflux of δ-tocotrienol out of the cell [14]. Similarly, Han et al. achieved the efflux of δ-tocotrienol via the overexpression of the transporter protein PDR11 in conjunction with biphasic fermentation [15]. Biphasic fermentation is commonly used for promoting product efflux. Dodecane (5 mL) was therefore added herein to the fermentation medium (50 mL YPD medium supplemented with 0.1 % [W/V] tyrosine) after 30 h of fermentation using strain T03 to promote the efflux of δ-tocotrienol. After 96 h of fermentation, the extracted intracellular products and dodecane phase were subjected to HPLC analysis. The results showed that the addition of dodecane promoted the efflux of δ-tocotrienol, with the amount of δ-tocotrienol secreted outside the cell accounting for 26.13 % of the total production (Fig. 5a). In addition, the efflux of the product drives the metabolic flux toward the synthesis of δ-tocotrienol. As a result, the total production of δ-tocotrienol using this approach was found to be 1.69 times that obtained using nonbiphasic fermentation (Fig. 5a).

Fig. 5.

Fig. 5

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The impact of the passive efflux engineering with dodecane added to the culture medium and the active efflux engineering with overexpression of transport protein PDR1 to promote product efflux on the production of δ-tocotrienol. (a) The effects of adding dodecane and overexpression of transporter protein PDR1 by different types of promoters on the production and efflux of δ-tocotrienol. (b) δ-tocotrienol production ability of engineered strains integrated engineering strategies of precursor supply enhancing and efflux enhancing.

Previous studies have shown that the endogenous ABC transporters in S. cerevisiae can promote the efflux of hydrophobic products [40]. A screening of the ABC transporters by Jiao et al. revealed that the transporter protein PDR1 had the maximum effect on the efflux of δ-tocotrienol [14]. Therefore, PDR1 was considered herein as the transporter protein responsible for enhancing the efflux of δ-tocotrienol. Considering that the ABC transporters are dependent on ATP hydrolysis as a source of energy for transport of molecules against a concentration gradient, their overexpression may result in competition for ATP with other cellular activities that require energy supply. Therefore, the promoters with relatively low (PTEF2), medium (PTEF1), high (PCCW12) [41], and inducible-high (PGAL10) strength were employed herein for PDR1 overexpression. The insertion of PDR1 expression cassettes of different promoter strengths into the DPP1 locus of the T03 genome yielded the strains D01–D04, respectively. Fermentation was carried out with the strains D01–D04 in 50 mL YPD liquid medium supplemented with 0.1 % (W/V) tyrosine. After 96 h of fermentation, the intracellular products were extracted and analyzed using HPLC. However, δ-tocotrienol was not detectable in the fermentation broth, which may be attributable to the low efflux of δ-tocotrienol (resulting in levels that were below the detection limit of HPLC) obtained upon the overexpression of the transporter protein alone. Subsequently, PDR1 overexpression was carried out in conjunction with biphasic fermentation and the addition of dodecane to the fermentation broth after 30 h of fermentation. The results showed that the strain D04 expressing PDR1 from the promoter PGAL10 exhibited a significant increase in δ-tocotrienol production compared to that in the strain T03 undergoing biphasic fermentation without PDR1 overexpression; the yield obtained was 1.57 times that with strain T03 under conditions of biphasic fermentation. Extracellular secretion of δ-tocotrienol in strain D04 accounted for 25.18 % of the total δ-tocotrienol produced. However, compared to strain T03, only weak enhancement in the production of δ-tocotrienol was observed in strains with PDR1 overexpression from constitutive promoters; moreover, the production even decreased in the group harboring PTEF1-PDR1. This may be attributable to the burden on cell growth caused by the constitutive overexpression of the transporter protein PDR1, given the decrease in the biomass of all strains expressing PDR1 from the constitutive promoters.

Subsequently, the various optimization strategies that had a positive effect on δ-tocotrienol production were integrated herein. Specifically, the strain T04 with enhanced precursor supply was used as the starting strain. The expression cassette PGAL1-PDR1 was inserted into the DPP1 site of strain T04 to obtain strain T05. Furthermore, the expression cassette encoding PaCrtE(GSG)SyHPT was inserted at the δ site of strain T05 to obtain strain T06. Biphasic fermentation (with added dodecane) was carried out with strain T06 in YPD medium supplemented with 0.1 % (W/V) tyrosine. After fermentation, the intracellular products were extracted and analyzed using HPLC. The strain T06, in which all the aforementioned metabolic engineering strategies have been integrated, produced 2682.6 μg/L of δ-tocotrienol; this value is 3.3 times higher than that obtained with the control strain T03, in which only the basic δ-tocotrienol biosynthesis pathways have been integrated (Fig. 5b). This result proves that these strategies effectively increased the production of δ-tocotrienol.

3.4. Engineering of AtTC via evolution-guided mutagenesis

The previous results showed that the overexpression of AtTC from a single copy of the corresponding gene failed to support the production of sufficient quantities of δ-tocotrienol for detection using HPLC, which is indicative of the limited catalytic activity of AtTC in S. cerevisiae. Therefore, engineering modifications were carried out on AtTC subsequently to enhance its catalytic activity in yeast. The application of protein engineering strategies for modifying enzymes and improving their catalytic activities has been widely studied; directed evolution and semirational design represent two commonly used protein engineering strategies for enzyme modification. However, directed evolution requires the construction of a large number of mutants and offers rare positive results during screening for mutations. Herein, multiple sequence alignment with other tocopherol cyclase was carried out for facilitating a semirational design of AtTC to improve its catalytic activity. Homologous residues in an enzyme reflect a “stable evolutionary intermediate” with a sequence that is closer to the optimum enzyme sequence from a functional perspective compared to that obtained via random mutations, thereby reducing the number of steps required for directed protein evolution.

Amino acid sequences of 78 TCs were selected from the National Center for Biotechnology Information database for homologous sequence alignment. These 78 sequences were aligned using Clustal X, and the results were visualized using the online tool WebLogo 3 (Fig. S2); conserved amino acid residues were identified from the aligned sequences of multiple TCs. An evaluation of the amino acid sequence of AtTC compared to the conserved amino acid sequence allowed the preliminary selection of 83 candidate residues; these amino acid residues of AtTC were mutated to the conserved amino acid, which is expected to improve the catalytic activity of AtTC. Herein, primers were designed to allow the incorporation of two or three adjacent mutations into a single set of primers to facilitate preliminary screening; for example, serine at position 70 and glutamic acid at position 74, which are located close to each other, were incorporated into a single pair of primers for mutagenesis. The plasmids pESC-URA harboring wild-type AtTC and its 35 mutants were used for the transformation of the strain T01 to yield the strains T01-0 to T01-35. These strains were employed for fermentation for 96 h in 50 mL SD-URA medium supplemented with 0.1 % (W/V) tyrosine. After fermentation, the intracellular products were extracted and analyzed using HPLC. The production of δ-tocotrienol in the strains T01-2, T01-18, T01-22, T01-26, T01-27, and T01-33 were found to be approximately 1.19, 1.23, 1.91, 1.34, 1.2, and 1.33 times higher, respectively, than that in the control strain T01-0 (Fig. 6b).

Fig. 6.

Fig. 6

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Enzyme engineering of tocopherol cyclase AtTC. (a) Screening the conserved amino acid sites guided by multiple sequence alignment with other tocopherol cyclase. (b) The identification of individual mutations responsible for the increase in δ-tocotrienol production. (c) The evaluation of dual mutations combined from four individual mutations T87S, R407K, S409T, and Q468N. (d) Screening of strains integrated with SyHPT-PGal10,Gal1-AtTCT87S spontaneous replication expression cassette. (e) Scale-up fermentation of δ-tocotrienol producing engineered strain T08 in 5 L bioreactors.

The mutant AtTCs employed in the six strains identified above harbored a total of 15-point mutations, including T87S, P88T, K90R, R284E, D285N, R360S, M363I, T398S, V400A, T404P, R407K, S409T, K464R, Q465R, and Q468N. To identify the individual mutations among these that were responsible for the increase in δ-tocotrienol production, AtTC mutants with single mutations were cloned into plasmid pESC-URA which were then used for the transformation of strain T01. The strains so obtained were used for fermentation, and the products were extracted as above. Compared with that of the control strain T01-0, the strains harboring the mutants T87S, R407K, S409T, and Q468N exhibited significantly increased production of δ-tocotrienol (approximately 2.33, 1.83, 2.17, and 2.30 times that of strain T01-0, respectively, Fig. 6c). Molecular docking analysis of the AtTC mutants with MGGBQ (Fig. S3) and the calculation of binding affinities revealed a more stable binding of these four mutants with the substrate MGGBQ compared to that of the wild-type AtTC (Table S4), which is consistent with their ability to support increased production of δ-tocotrienol. Additionally, δ-tocotrienol production was not detectable in the yeast strains harboring the mutants T398S, V400A, and T404P; this may be attributable to the location of these three sites near the active pocket of AtTC during the catalysis of MGGBQ cyclization, such that the mutations resulted in loss of enzyme activity.

The four mutations T87S, R407K, S409T, and Q468N, which supported significantly increased production of δ-tocotrienol, were then combined in pairs to evaluate the effects of the dual mutations on δ-tocotrienol production. The six dual-site mutants of AtTC were cloned into plasmids and used for the transformation of the strain T01 to obtain the corresponding strains. Compared with that of the control strain T01-0, the strains harboring the dual-site AtTC mutants T87S/S409T, R407K/Q468N, and S409T/Q468N showed an increase in δ-tocotrienol production (approximately 1.19, 1.29, and 1.09 times that of T01-0, respectively, Fig. 6d), albeit the levels were lower than those obtained with the corresponding single-site mutants.

To obtain a genetically stable strain containing the AtTCT87S mutant, a strain supporting a high yield of δ-tocotrienol with the integration of all the aforementioned metabolic engineering strategies was reconstructed herein. The precursor supply enhancement module comprising the genes encoding ALD6, SeACS, ADH2, and PaCrtE, the overexpression cassette PGAL1-PDR1 for facilitating protein efflux, and the expression cassette for the fusion enzyme PaCrtE(GSG)SyHPT were all integrated into strain T01 according to the method described earlier, generating strain T07. Finally, the expression cassette SyHPT-PGal10, PGal1-AtTCT87S was inserted into the RPL25 site of strain T07 according to the method reported in the previous study to generate clones DT01–DT06. Following fermentation using the strains and detection of the products using HPLC, the highest production of δ-tocotrienol (4337.3 μg/L) was obtained with the clone DT02, which was subsequently named as strain T08. The yield of δ-tocotrienol obtained with strain T08 was 1.6 times higher than that obtained with strain T06 and 8.9 times higher than that obtained with strain T03, in which only the complete tocotrienol synthesis pathway was integrated.

The fermentation processes using strain T08 were scaled up to a 5-L bioreactor. A two-stage characteristic was exhibited during the fermentation process. The first stage mainly involved rapid cell growth, which was accompanied by a continuous increase in biomass such that an OD600 of 48 was attained by 30 h. Galactose was then added at this juncture to induce the expression of genes associated with terpenoid production. An OD600 of 60 was attained after 48 h of fermentation; at this juncture, the feed was replaced with ethanol, allowing the initiation of the second stage of fermentation. During this stage, the growth rate of the strain is reduced, and the metabolic flux is increasingly directed toward the production of δ-tocotrienol, allowing its obvious accumulation. The peak yield of δ-tocotrienol (16.9 mg/L) was attained after 128 h of fermentation, followed by a reduction in product yield and the subsequent cessation of the fermentation processes at 144 h.

4. Conclusions

In the current study, the metabolic pathways of S. cerevisiae Sc027 were modified in multiple ways to enhance its ability to produce δ-tocotrienol. The following strategies were employed: 1) Overexpression of the genes associated with acetyl CoA production to increase the supply of the terpenoid precursor acetyl CoA. 2) Construction of the fusion protein PaCrtE(GSG)SyHPT to partially resolve the metabolic bottleneck. 3) Overexpression of proteins facilitating product efflux and the application of biphasic fermentation to enhance the efflux of the product δ-tocotrienol, resulting in redirection of the metabolic flux towards the synthesis of δ-tocotrienol. 4) Systematic modification of AtTC to improve its catalytic performance and further enhance the production of δ-tocotrienol by the engineered strain. Among these various approaches, the construction of a substrate channel by the fusion of the GGPP synthase PaCrtE and SyHPT using a linker to improve the efficiency of the reaction and the use of the mutant AtTCT87S to significantly improve the catalytic performance of AtTC are unique innovations employed in the current study and as such, have been reported for the first time herein. The approach of generating a fusion protein of CrtE and HPT can be extended as a universal synthetic biology strategy in biosynthesis of δ-tocotrienol by S. cerevisiae, while the AtTCT87S mutant can serve as a universal biopart for future research on δ-tocotrienol production.

CRediT authorship contribution statement

Ziming Liu: Writing – original draft, Validation, Supervision, Project administration, Investigation, Funding acquisition, Conceptualization. Min Tang: Writing – original draft, Visualization, Validation, Methodology, Investigation, Data curation, Conceptualization. Wanze Zhang: Writing – original draft, Visualization, Validation, Supervision, Software, Project administration, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. Yanjie Tian: Validation, Software, Methodology, Investigation. Jianjun Qiao: Writing – review & editing, Validation, Resources, Project administration, Methodology, Investigation, Funding acquisition, Conceptualization. Mingzhang Wen: Writing – review & editing, Software, Project administration, Methodology, Funding acquisition, Formal analysis, Conceptualization. Weiguo Li: Writing – review & editing, Validation, Supervision, Resources, Project administration, Funding acquisition. Qinggele Caiyin: Writing – review & editing, Validation, Supervision, Resources, Project administration, Methodology, Funding acquisition, Conceptualization.

Funding

This work was supported by Zhejiang Provincial Natural Science Foundation of China (No. LQN25C010006) and the National Key Research and Development Program of China (No. 2020YFA0907900).

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Footnotes

Peer review under the responsibility of Editorial Board of Synthetic and Systems Biotechnology.

Appendix A

Supplementary data to this article can be found online at https://doi.org/10.1016/j.synbio.2025.11.013.

Contributor Information

Weiguo Li, Email: liweiguo@tju.edu.cn.

Qinggele Caiyin, Email: qinggele@tju.edu.cn.

Appendix A. Supplementary data

The following is the Supplementary data to this article:

Multimedia component 1
mmc1.docx (10.2MB, docx)

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