Characterization and designing of an SAM riboswitch to establish a high-throughput screening platform for SAM overproduction in Saccharomyces cerevisiae

S-adenosyl- l -methionine (SAM) is a high-value compound widely used in the treatment of various diseases. SAM can be produced through fermentation, but further enhancing the microbial production of SAM requires novel high-throughput screening methods for rapid detection and screening of mutant libraries. In this work, an SAM-OFF riboswitch capable of responding to the SAM concentration was obtained and a high-throughput platform for screening SAM overproducers was established. SAM synthase was engineered by semirational design and directed evolution, which resulted in the SAM2^{S203F,W164R,T251S,Y285F,S365R} mutant with almost twice higher catalytic activity than the parental enzyme. The best mutant was then introduced into Saccharomyces cerevisiae BY4741, and the resulting strain BSM8 produced a sevenfold higher SAM titer in shake-flask fermentation, reaching 1.25 g L⁻¹. This work provides a reference for designing biosensors to dynamically detect metabolite concentrations for high-throughput screening and the construction of effective microbial cell factories.     

Protease-Deficient Saccharomyces cerevisiae Strains for the Synthesis of Human-Compatible Glycoproteins

Saccharomyces cerevisiae strains engineered previously to produce proteins with mammalian high mannose structures showed severe growth defects and decreased protein productivity. In strain YAB101, derived from one of these strains by a mutagenesis technique based on the disparity theory of evolution, these undesirable phenotypes were alleviated. Here we describe further engineering of YAB101 with the aim of synthesizing heterologous glycoproteins with Man₅GlcNAc₂, an intermediate for the mammalian hybrid and complex type oligosaccharides. About 60% conversion of Man₈GlcNAc₂ to Man₅GlcNAc₂ was observed after integration of Aspergillus saitoi α-1,2-mannosidase fused to the transmembrane domain of S. cerevisiae Och1. To obtain a higher yield of the target protein, a protease-deficient version of this strain was generated by disruption of PEP4 and PRB1, resulting in YAB101-4. Inactivation of these vacuolar proteases enhanced the secretion of human interferon-β by approximately 10-fold.     

In vivo evolutionary engineering of riboswitch with high-threshold for N-acetylneuraminic acid production

Riboswitches with desired properties, such as sensitivity, threshold, dynamic range, is important for its application. However, the property change of a natural riboswitch is difficult due to the lack of the understanding of aptamer ligand binding properties and a proper screening method for both rational and irrational design. In this study, an effective method to change the threshold of riboswitch was established in vivo based on growth coupled screening by combining both positive and negative selections. The feasibility of the method was verified by the model library. Using this method, an N-acetylneuraminic acid (NeuAc) riboswitch was evolved and modified riboswitches with high threshold and large dynamic range were obtained. Then, using a new NeuAc riboswitch, both ribosome binding sites and key gene in NeuAc biosynthesis pathway were optimized. The highest NeuAc production of 14.32 g/l that has been reported using glucose as sole carbon source was obtained.     

Engineering Saccharomyces cerevisiae for isoprenol production

Isoprenol (3-methyl-3-butene-1-ol) is a valuable drop-in biofuel and an important precursor of several commodity chemicals. Synthetic microbial systems using the heterologous mevalonate pathway have recently been developed for the production of isoprenol in Escherichia coli, and a significant yield and titer improvement has been achieved through a decade of research. Saccharomyces cerevisiae has been widely used in the biotechnology industry for isoprenoid production, but there has been no good example of isoprenol production reported in this host. In this study, we engineered the budding yeast S. cerevisiae for improved biosynthesis of isoprenol. The strain engineered with the mevalonate pathway achieved isoprenol production at the titer of 36.02 ± 0.92 mg/L in the flask. The IPP (isopentenyl diphosphate)-bypass pathway, which has shown more efficient isoprenol production by avoiding the accumulation of the toxic intermediate in E. coli, was also constructed in S. cerevisiae and improved the isoprenol titer by 2-fold. We further engineered the strains by deleting a promiscuous endogenous kinase that could divert the pathway flux away from the isoprenol production and improved the titer to 130.52 ± 8.01 mg/L. Finally, we identified a pathway bottleneck using metabolomics analysis and overexpressed a promiscuous alkaline phosphatase to relieve this bottleneck. The combined efforts resulted in the titer improvement to 383.1 ± 31.62 mg/L in the flask. This is the highest isoprenol titer up to date in S. cerevisiae and this work provides the key strategies to engineer yeast as an industrial platform for isoprenol production.     

Advanced method for high-throughput expression of mutated eukaryotic membrane proteins in Saccharomyces cerevisiae

Crystallization of eukaryotic membrane proteins is a challenging, iterative process. The protein of interest is often modified in an attempt to improve crystallization and diffraction results. To accelerate this process, we took advantage of a GFP-fusion yeast expression system that uses PCR to direct homologous recombination and gene cloning. We explored the possibility of employing more than one PCR fragment to introduce various mutations in a single step, and found that when up to five PCR fragments were co-transformed into yeast, the recombination frequency was maintained as the number of fragments was increased. All transformants expressed the model membrane protein, while the resulting plasmid from each clone contained the designed mutations only. Thus, we have demonstrated a technique allowing the expression of mutant membrane proteins within 5 days, combining a GFP-fusion expression system and yeast homologous recombination.     

Metabolic engineering of Saccharomyces cerevisiae for production of ginsenosides

Ginsenosides are the primary bioactive components of ginseng, which is a popular medicinal herb and exhibits diverse pharmacological activities. Protopanaxadiol is the aglycon of several dammarane-type ginsenosides, which also has anticancer activity. For microbial production of protopanaxadiol, dammarenediol-II synthase and protopanaxadiol synthase genes of Panax ginseng, together with a NADPH-cytochrome P450 reductase gene of Arabidopsis thaliana, were introduced into Saccharomyces cerevisiae, resulting in production of 0.05 mg/g DCW protopanaxadiol. Increasing squalene and 2,3-oxidosqualene supplies through overexpressing truncated 3-hydroxyl-3-methylglutaryl-CoA reductase, farnesyl diphosphate synthase, squalene synthase and 2,3-oxidosqualene synthase genes, together with increasing protopanaxadiol synthase activity through codon optimization, led to 262-fold increase of protopanaxadiol production. Finally, using two-phase extractive fermentation resulted in production of 8.40 mg/g DCW protopanaxadiol (1189 mg/L), together with 10.94 mg/g DCW dammarenediol-II (1548 mg/L). The yeast strains engineered in this work can serve as the basis for creating an alternative way for production of ginsenosides in place of extraction from plant sources.     

Heterologous production of the epoxycarotenoid violaxanthin in Saccharomyces cerevisiae

Microbial production of carotenoids has mainly focused towards a few products, such as β-carotene, lycopene and astaxanthin. However, other less explored carotenoids, like violaxanthin, have also shown unique properties and promissory applications. Violaxanthin is a plant-derived epoxidated carotenoid with strong antioxidant activity and a key precursor of valuable compounds, such as fucoxanthin and β-damascenone. In this study, we report for the first time the heterologous production of epoxycarotenoids in yeast. We engineered the yeast Saccharomyces cerevisiae following multi-level strategies for the efficient accumulation of violaxanthin. Starting from a β-carotenogenic yeast strain, we first evaluated the performance of several β-carotene hydroxylases (CrtZ), and zeaxanthin epoxidases (ZEP) from different species, together with their respective N-terminal truncated variants. The combined expression of CrtZ from Pantoea ananatis and truncated ZEP of Haematococcus lacustris showed the best performance and led to a yield of 1.6 mg/g_DCW of violaxanthin. Further improvement of the epoxidase activity was achieved by promoting the transfer of reducing equivalents to ZEP by expressing several redox partner systems. The co-expression of the plant truncated ferredoxin-3, and truncated root ferredoxin oxidoreductase-1 resulted in a 2.2-fold increase in violaxanthin yield (3.2 mg/g_DCW). Finally, increasing gene copy number of carotenogenic genes enabled reaching a final production of 7.3 mg/g_DCW in shake flask cultures and batch bioreactors, which is the highest yield of microbially produced violaxanthin reported to date.     

A general method for determining the contribution of split pathways in metabolite production in the yeast Saccharomyces cerevisiae

We describe a simple method using (13)C labeling and NMR spectroscopy to determine the flux contributions of alternative pathways in Saccharomyces cerevisiae that produce the same metabolite with identical labeling patterns. Cells were incubated with a (13)C-labeled precursor for one of the branches, and the absolute enrichment of the product and its metabolic precursor(s) was quantified. The ratio of the absolute enrichment of the product to that of its precursor reflects the contribution of the pathway. The method was applied to the biosynthesis of glycine in yeast, which can occur from threonine via threonine aldolase or from serine via serine hydroxymethyltransferase. [2-(13)C]Aspartate and [2-(13)C]serine were used as labeled precursors for the threonine aldolase and serine hydroxymethyltransferase pathways, respectively. The data show that in cells possessing both pathways, the serine hydroxymethyltransferase pathway contributes 65-75% of the total glycine production. In comparison with other approaches, this method provides an inexpensive, flexible alternative to determining the flux contributions of split pathways under controlled conditions and should have wide applicability in the metabolic engineering of microorganisms.     

Construction of industrial xylose-fermenting Saccharomyces cerevisiae strains through combined approaches

Saccharomyces cerevisiae strain with excellent xylose-fermenting capacity and inhibitor tolerance is crucial for lignocellulosic ethanol production. In this study, a combined strategy including site-directed mutagenesis, mating, evolutionary engineering, and haploidization was applied to obtain strains with ideal xylose fermentabilities. Haploid industrial strain KFG4-6B was engineered to overexpress endogenous xylulokinase (XK) and heterologous native or mutated xylose reductase (XR) and xylitol dehydrogenase (XDH) from Scheffersomyces stipitis. The XR-mutated strain HX57D showed over 12% increase in both xylose consumption rate and ethanol yield compared with the XR-native strain. To improve the xylose uptake, the HX57D-derived diploids were subjected to evolutionary engineering. In comparison with HX57D, evolved diploid Z4X-21-18 achieved 4.5-fold increases in rates of xylose consumption and ethanol production when fermenting xylose. When fermenting mixed sugars, the glucose and xylose uptake rates were 1.4-fold and 8.3-fold, respectively, higher. H18s28, a haploid of Z4X-21-18, enabled a further 10% increase in xylose consumption rate when fermenting xylose only. However, it was inferior to its diploid parent when fermenting mixed sugars. In the presaccharification-simultaneous saccharification and fermentation (P-SSF) of the whole pretreated wheat straw slurry with high contents of multiple inhibitors, Z4X-21-18 produced approximately 42 g/L ethanol with a yield of 0.38 g/g total sugars.     

A micro-scale process for high-throughput expression of cDNAs in the yeast Saccharomyces cerevisiae

Methods have been developed aimed at applying at high-throughput technology for expression of cloned cDNAs in yeast. Yeast is a eukaryotic host, which produces soluble recombinant proteins and is capable of introducing post-translational modifications of protein. It is, thus, an appropriate expression system both for the routine expression of various cDNAs or protein domains and for the expression of proteins, which are not correctly expressed in Escherichia coli. Here, we describe a standard system in Saccharomyces cerevisiae, based on a vector for intracellular protein expression, where the gene products are fused to specific peptide sequences (tags). These epitope tags, the N-terminal His(6) tag and the C-terminal StrepII tag, allow subsequent immunological identification and purification of the gene products by a two-step affinity chromatography. This method of dual-tagged recombinant protein purification eliminates contamination by degraded protein products. A miniaturization of the procedures for cloning, expression, and detection was performed to allow all steps to be carried out in 96-well microtiter plates. The system is, thus, suitable for automation. We were able to analyze the simultaneous protein expression of a large number of cDNA clones due to the highly parallel approach of protein production and purification. The microtiter plate technology format was extended to quantitative analysis. An ELISA-based assay was developed that detects StrepII-tagged proteins. The application of this high-throughput expression system for protein production will be a useful tool for functional and structural analyses of novel genes, identified by the Human Genome Project and other large-scale sequencing projects.     

Utilizing an endogenous pathway for 1-butanol production in Saccharomyces cerevisiae

Microbial production of higher alcohols from renewable feedstock has attracted intensive attention thanks to its potential as a source for next-generation gasoline substitutes. Here we report the discovery, characterization and engineering of an endogenous 1-butanol pathway in Saccharomyces cerevisiae. Upon introduction of a single gene deletion adh1Δ, S. cerevisiae was able to accumulate more than 120 mg/L 1-butanol from glucose in rich medium. Precursor feeding, ¹³C-isotope labeling and gene deletion experiments demonstrated that the endogenous 1-butanol production was dependent on catabolism of threonine in a manner similar to fusel alcohol production by the Ehrlich pathway. Specifically, the leucine biosynthesis pathway was engaged in the conversion of key 2-keto acid intermediates. Overexpression of the pathway enzymes and elimination of competing pathways achieved the highest reported 1-butanol titer in S. cerevisiae (242.8 mg/L).     

Genetic analysis of the metabolic pathways responsible for aroma metabolite production by Saccharomyces cerevisiae

During alcoholic fermentation, higher alcohols, esters, and acids are formed from amino acids via the Ehrlich pathway by yeast, but many of the genes encoding the enzymes have not yet been identified. When the BAT1/2 genes, encoding transaminases that deaminate amino acids in the first step of the Ehrlich pathway are deleted, higher metabolite formation is significantly decreased. Screening yeast strains with deletions of genes encoding decarboxylases, dehydrogenases, and reductases revealed nine genes whose absence had the most significant impact on higher alcohol production. The seven most promising genes (AAD6, BAT2, HOM2, PAD1, PRO2, SPE1, and THI3) were further investigated by constructing double- and triple-deletion mutants. All double-deletion strains showed a greater decrease in isobutanol, isoamyl alcohol, isobutyric, and isovaleric acid production than the corresponding single deletion strains with the double-deletion strains in combination with ?bat2 and the ?hom2-?aad6 strain revealing the greatest impact. BAT2 is the dominant gene in these deletion strains and this suggests the initial transaminase step of the Ehrlich pathway is rate-limiting. The triple-deletion strains in combination with BAT2 (?bat2-?thi3-?aad6 and ?bat2-?thi3-?hom2) had the greatest impact on the end metabolite production with the exception of isoamyl alcohol and isovaleric acid. The strain deleted for two dehydrogenases and a reductase (?hom2-?pro2-?aad6) had a greater effect on the levels of these two compounds. This study contributes to the elucidation of the Ehrlich pathway and its significance for aroma production by fermenting yeast cells.     

A fluorescence-based glycosyltransferase assay for high-throughput screening

Glycosyltransferases catalyze the transfer of a monosaccharide unit from a nucleotide or lipid sugar donor to polysaccharides, lipids, and proteins in a stereospecific manner. Considerable effort has been invested in engineering glycosyltransferases to diversify sugar-containing drugs. An important requirement for glycosyltransferase engineering is the availability of a glycosyltransferase assay system for high-throughput screening of glycosyltransferase mutants. In this study, a general glycosyltransferase assay system was developed based on an ATP sensor. This system showed submicromolar sensitivity and compatibility with both purified enzymes and crude cell extracts. The assay system will be useful for glycosyltransferase engineering based on high-throughput screening, as well as for general glycosyltransferase assays and kinetics.     

A high-throughput pH indicator assay for screening glycosyltransferase saturation mutagenesis libraries

Protein engineering using directed evolution or saturation mutagenesis at hot spots is often used to improve enzyme properties such as their substrate selectivity or stability. This requires access to robust high-throughput assays to facilitate the analysis of enzyme libraries. However, relatively few studies on directed evolution or saturation mutagenesis of glycosyltransferases have been reported in part due to a lack of suitable screening methods. In the present study we report a general screening assay for glycosyltransferases that has been developed using the blood group α-(1→3)-galactosyltransferase (GTB) as a model. GTB utilizes UDP-Gal as a donor substrate and α-L-Fucp-(1→2)-β-D-Galp-O-R (H antigen) as an acceptor substrate and synthesizes the blood group B antigen α-D-Galp-(1→3)-[α-L-Fucp-(1→2)]-β-D-Galp-O-R. A closely related α-(1→3)-N-acetylgalactosaminyltransferase (GTA) uses UDP-GalNAc as a donor with the same H acceptor, yielding the A antigen α-D-Galp-NAc-(1→3)-[α-L-Fuc(1→2)]-β-D-Gal-O-R. GTA and GTB are highly homologous enzymes differing in only 4 of 354 amino acids, Arg/Gly-176, Gly/Ser-235, Leu/Met-266, and Gly/Ala-268. The screening assay is based on the color change of the pH indicator bromothymol blue when a proton is released during the transfer of Gal/GalNAc from UDP-Gal/UDP-GalNAc to the acceptor substrate. Saturation mutagenesis of GTB enzyme at M214, a hot spot adjacent to the ²¹¹DVD²¹³ metal binding motif, was performed and the resulting library was screened for increases in UDP-GalNAc transfer activity. Two novel mutants, M214G and M214S, identified by pH indicator screening, were purified and kinetically characterized. M214S and M214G both exhibited two-fold higher k_cat and specific activity than wild-type GTB for UDP-GalNAc. The results confirm the importance of residue M214 for donor enzyme specificity.     

代谢工程改造酿酒酵母合成葡萄糖二酸

葡萄糖二酸是一种高附加值的有机酸,广泛用于食品、医药和化工领域。为获得生产葡萄糖二酸的微生物细胞工厂,通过共表达小鼠来源的肌醇加氧酶(MIOX)及恶臭假单胞菌来源的醛酸脱氢酶(Udh),在酿酒酵母Saccharomyces cerevisiae CEN.PK2-1C中构建了葡萄糖二酸合成途径,产量为(28.28±3.15)mg/L。在此基础上,通过调控前体肌醇的合成途径,发现肌醇-1-磷酸合成酶(INO1)是葡萄糖二酸合成途径的限速酶,过量表达INO1,葡萄糖二酸产量达到(107.51±10.87)mg/L,提高了2.8倍。进一步弱化竞争支路中磷酸果糖激酶(PFK1)的表达,最终葡萄糖二酸的产量达到(230.22±10.75)mg/L,为进一步获得高产葡萄糖二酸细胞工厂提供基础。     

Xylose assimilation enhances the production of isobutanol in engineered Saccharomyces cerevisiae

Bioconversion of xylose—the second most abundant sugar in nature—into high-value fuels and chemicals by engineered Saccharomyces cerevisiae has been a long-term goal of the metabolic engineering community. Although most efforts have heavily focused on the production of ethanol by engineered S. cerevisiae, yields and productivities of ethanol produced from xylose have remained inferior as compared with ethanol produced from glucose. However, this entrenched focus on ethanol has concealed the fact that many aspects of xylose metabolism favor the production of nonethanol products. Through reduced overall metabolic flux, a more respiratory nature of consumption, and evading glucose signaling pathways, the bioconversion of xylose can be more amenable to redirecting flux away from ethanol towards the desired target product. In this report, we show that coupling xylose consumption via the oxidoreductive pathway with a mitochondrially-targeted isobutanol biosynthesis pathway leads to enhanced product yields and titers as compared to cultures utilizing glucose or galactose as a carbon source. Through the optimization of culture conditions, we achieve 2.6 g/L of isobutanol in the fed-batch flask and bioreactor fermentations. These results suggest that there may be synergistic benefits of coupling xylose assimilation with the production of nonethanol value-added products.     

Efficient production of glutathione in Saccharomyces cerevisiae via a synthetic isozyme system

Glutathione, a tripeptide consisting of cysteine, glutamic acid, and glycine, has multiple beneficial effects on human health. Previous studies have focused on producing glutathione in Saccharomyces cerevisiae by overexpressing γ-glutamylcysteine synthetase (GSH1) and glutathione synthetase (GSH2), which are the rate-limiting enzymes involved in the glutathione biosynthetic pathway. However, the production yield and titer of glutathione remain low due to the feedback inhibition on GSH1. To overcome this limitation, a synthetic isozyme system consisting of a novel bifunctional enzyme (GshF) from Gram-positive bacteria possessing both GSH1 and GSH2 activities, in addition to GSH1/GSH2, was introduced into S. cerevisiae, as GshF is insensitive to feedback inhibition. Given the HSP60 chaperonin system mismatch between bacteria and S. cerevisiae, co-expression of Group-I HSP60 chaperonins (GroEL and GroES) from Escherichia coli was required for functional expression of GshF. Among various strains constructed in this study, the SKSC222 strain capable of synthesizing glutathione with the synthetic isozyme system produced 240 mg L^-1 glutathione with glutathione content and yield of 4.3% and 25.6 mg_glutathione/g_glucose, respectively. These values were 6.6-, 4.9-, and 4.3-fold higher than the corresponding values of the wild-type strain. In a glucose-limited fed-batch fermentation, the SKSC222 strain produced 2.0 g L^-1 glutathione in 67 h. Therefore, this study highlights the benefits of the synthetic isozyme system in enhancing the production titer and yield of value-added chemicals by engineered strains of S. cerevisiae.     

Metabolic engineering of Saccharomyces cerevisiae for the production of triacetic acid lactone

Biobased chemicals have become attractive replacements for their fossil-fuel counterparts. Recent studies have shown triacetic acid lactone (TAL) to be a promising candidate, capable of undergoing chemical conversion to sorbic acid and other valuable intermediates. In this study, Saccharomyces cerevisiae was engineered for the high-level production of TAL by overexpression of the Gerbera hybrida 2-pyrone synthase (2-PS) and systematic engineering of the yeast metabolic pathways. Pathway analysis and a computational approach were employed to target increases in cofactor and precursor pools to improve TAL synthesis. The pathways engineered include those for energy storage and generation, pentose biosynthesis, gluconeogenesis, lipid biosynthesis and regulation, cofactor transport, and fermentative capacity. Seventeen genes were selected for disruption and independently screened for their effect on TAL production; combinations of knockouts were then evaluated. A combination of the pathway engineering and optimal culture parameters led to a 37-fold increase in titer to 2.2 g/L and a 50-fold increase in yield to 0.13 (g/g glucose). These values are the highest reported in the literature, and provide a 3-fold improvement in yield over previous reports using S. cerevisiae. Identification of these metabolic bottlenecks provides a strategy for overproduction of other acetyl-CoA-dependent products in yeast.     

Genetic studies of the PRP11 gene of Saccharomyces cerevisiae

Summary PRP11 is a gene that encodes an essential function for pre-messenger RNA (mRNA) processing in Saccharomyces cerevisiae. We have carried out a mutational study to locate essential and non-essential regions of the PRP11 protein. The existing temperature-sensitive (ts) mutation (prpl1-1) was isolated from the chromosome of the original mutant and its position in the gene was determined. When the prp11-1 gene was transcribed from the GAL1 promoter, the overproduced protein was able to reverse the is prp11-1 phenotype; this is compatible with the possibility that the defect in the prp11-1 gene product affects its binding to the spliceosome. Thirteen linker-insertion mutations were constructed. Only five (prp11-4, 11-6, 11-10, –13 and –14) resulted in a null phenotype. One of these became temperature-sensitive when the insertion was.reduced in size from four (prp11-10) to two (prp11-15) amino acids. A sequence of ten amino acids of which also occurs in the human U1 small nuclear ribonucleoprotein particle (snRNP) A protein and the U2 snRNP B protein, when deleted from PRP11, had no phenotype and thus appears to be nonessential for PRP11 function. However, a linker-insertion mutation (prp11–10) immediately adjacent to this region resulted in a null phenotype.