amdSYM, a new dominant recyclable marker cassette for Saccharomyces cerevisiae

Construction of industrial Saccharomyces cerevisiae strains for the efficient consolidated bioprocessing of raw starch

BACKGROUND

Consolidated bioprocessing (CBP) combines enzyme production, saccharification and fermentation into a one-step process. This strategy represents a promising alternative for economic ethanol production from starchy biomass with the use of amylolytic industrial yeast strains.

RESULTS

Recombinant Saccharomyces cerevisiae Y294 laboratory strains simultaneously expressing an α-amylase and glucoamylase gene were screened to identify the best enzyme combination for raw starch hydrolysis. The codon optimised Talaromyces emersonii glucoamylase encoding gene (temG_Opt) and the native T. emersonii α-amylase encoding gene (temA) were selected for expression in two industrial S. cerevisiae yeast strains, namely Ethanol Red™ (hereafter referred to as the ER) and M2n. Two δ-integration gene cassettes were constructed to allow for the simultaneous multiple integrations of the temG_Opt and temA genes into the yeasts' genomes. During the fermentation of 200 g l^-1 raw corn starch, the amylolytic industrial strains were able to ferment raw corn starch to ethanol in a single step with high ethanol yields. After 192 h at 30 °C, the S. cerevisiae ER T12 and M2n T1 strains (containing integrated temA and temG_Opt gene cassettes) produced 89.35 and 98.13 g l^-1 ethanol, respectively, corresponding to estimated carbon conversions of 87 and 94%, respectively. The addition of a commercial granular starch enzyme cocktail in combination with the amylolytic yeast allowed for a 90% reduction in exogenous enzyme dosage, compared to the conventional simultaneous saccharification and fermentation (SSF) control experiment with the parental industrial host strains.

CONCLUSIONS

A novel amylolytic enzyme combination has been produced by two industrial S. cerevisiae strains. These recombinant strains represent potential drop-in CBP yeast substitutes for the existing conventional and raw starch fermentation processes.

Engineering the protein secretory pathway of Saccharomyces cerevisiae enables improved protein production

Baker's yeast Saccharomyces cerevisiae is one of the most important and widely used cell factories for recombinant protein production. Many strategies have been applied to engineer this yeast for improving its protein production capacity, but productivity is still relatively low, and with increasing market demand, it is important to identify new gene targets, especially targets that have synergistic effects with previously identified targets. Despite improved protein production, previous studies rarely focused on processes associated with intracellular protein retention. Here we identified genetic modifications involved in the secretory and trafficking pathways, the histone deacetylase complex, and carbohydrate metabolic processes as targets for improving protein secretion in yeast. Especially modifications on the endosome-to-Golgi trafficking was found to effectively reduce protein retention besides increasing protein secretion. Through combinatorial genetic manipulations of several of the newly identified gene targets, we enhanced the protein production capacity of yeast by more than fivefold, and the best engineered strains could produce 2.5 g/L of a fungal α-amylase with less than 10% of the recombinant protein retained within the cells, using fed-batch cultivation.

A protocol for introduction of multiple genetic modifications in Saccharomyces cerevisiae using CRISPR/Cas9

Here, two methods are described for efficient genetic modification of Saccharomyces cerevisiae using CRISPR/Cas9. The first method enables the modification of a single genetic locus using in vivo assembly of a guide RNA (gRNA) expression plasmid without the need for prior cloning. A second method using in vitro assembled plasmids that could contain up to two gRNAs was used to simultaneously introduce up to six genetic modifications (e.g. six gene deletions) in a single transformation step by transforming up to three gRNA expression plasmids simultaneously. The method is not only suitable for gene deletion but is also applicable for in vivo site-directed mutagenesis and integration of multiple DNA fragments in a single locus. In all cases, the strain transformed with the gRNA expression plasmids was equipped with a genomic integration of Spcas9, leading to strong and constitutive expression of SpCas9. The protocols detailed here have been streamlined to be executed by virtually any yeast molecular geneticist.

Laboratory evolution for forced glucose-xylose co-consumption enables identification of mutations that improve mixed-sugar fermentation by xylose-fermenting Saccharomyces cerevisiae

Simultaneous fermentation of glucose and xylose can contribute to improved productivity and robustness of yeast-based processes for bioethanol production from lignocellulosic hydrolysates. This study explores a novel laboratory evolution strategy for identifying mutations that contribute to simultaneous utilisation of these sugars in batch cultures of Saccharomyces cerevisiae. To force simultaneous utilisation of xylose and glucose, the genes encoding glucose-6-phosphate isomerase (PGI1) and ribulose-5-phosphate epimerase (RPE1) were deleted in a xylose-isomerase-based xylose-fermenting strain with a modified oxidative pentose-phosphate pathway. Laboratory evolution of this strain in serial batch cultures on glucose-xylose mixtures yielded mutants that rapidly co-consumed the two sugars. Whole-genome sequencing of evolved strains identified mutations in HXK2, RSP5 and GAL83, whose introduction into a non-evolved xylose-fermenting S. cerevisiae strain improved co-consumption of xylose and glucose under aerobic and anaerobic conditions. Combined deletion of HXK2 and introduction of a GAL83G673T allele yielded a strain with a 2.5-fold higher xylose and glucose co-consumption ratio than its xylose-fermenting parental strain. These two modifications decreased the time required for full sugar conversion in anaerobic bioreactor batch cultures, grown on 20 g L-1 glucose and 10 g L-1 xylose, by over 24 h. This study demonstrates that laboratory evolution and genome resequencing of microbial strains engineered for forced co-consumption is a powerful approach for studying and improving simultaneous conversion of mixed substrates.

Structural, Physiological and Regulatory Analysis of Maltose Transporter Genes in Saccharomyces eubayanus CBS 12357T

Saccharomyces pastorianus lager brewing yeasts are domesticated hybrids of Saccharomyces cerevisiae and cold-tolerant Saccharomyces eubayanus. To understand the contribution of both parental genomes to maltose metabolism in brewing wort, this study focuses on maltose transport in the S. eubayanus type strain CBS 12357^T/FM1318. To obtain complete sequences of the MAL loci of this strain, a near-complete genome assembly was generated using the Oxford Nanopore Technology MinION sequencing platform. Except for CHRXII, all sixteen chromosomes were assembled as single contigs. Four loci harboring putative maltose transporter genes (SeMALT1-4), located in subtelomeric regions of CHRII, CHRV, CHRXIII, and CHRXVI, were completely resolved. The near-identical loci on CHRV and CHRXVI strongly resembled canonical S. cerevisiae MAL loci, while those on CHRII and CHRXIII showed different structures suggestive of gene loss. Overexpression of SeMALT1-4 in a maltose-transport-deficient S. cerevisiae strain restored growth on maltose, but not on maltotriose, indicating maltose-specific transport functionality of all four transporters. Simultaneous CRISPR-Cas9-assisted deletion of only SeMALT2 and SeMALT4, which shared 99.7% sequence identity, eliminated growth of S. eubayanus CBS 12357^T on maltose. Transcriptome analysis of S. eubayanus CBS 12357^T established that SeMALT1 and SeMALT3, are poorly expressed in maltose-grown cultures, while SeMALT2 and SeMALT4 were expressed at much higher levels than SeMALT1 and SeMALT3, indicating that only SeMALT2/4 are responsible for maltose consumption in CBS 12357^T. These results represent a first genomic and physiological characterization of maltose transport in S. eubayanus CBS 12357^T and provides a valuable resource for further industrial exploitation of this yeast.

Selection of Pof-Saccharomyces eubayanus Variants for the Construction of S. cerevisiae × S. eubayanus Hybrids With Reduced 4-Vinyl Guaiacol Formation

Saccharomyces pastorianus is an interspecies hybrid between S. cerevisiae and S. eubayanus. The identification of the parental species of S. pastorianus enabled the de novo reconstruction of hybrids that could potentially combine a wide array of phenotypic traits. Lager yeasts are characterized by their inability to decarboxylate ferulic acid present in wort, a phenotype also known as Pof ^- (phenolic off-flavor). However, all known S. eubayanus strains characterized so far produce clove-like aroma specific of 4-vinyl guaiacol, a decarboxylated form of ferulic acid. This study explored a non-GMO approach to construct Pof ^-S. eubayanus variants derived from the parental strain S. eubayanus CBS 12357. To rapidly screen a population of UV-mutagenized cells two complementary assays were developed. The first assay was based on the difference of light absorption spectra of ferulic acid and 4-vinyl guaiacol, while the second was based on the difference of sensitivity of Pof ^- and Pof⁺ strains to cinnamic acid. The S. eubayanus variant HTSE042 was selected and was confirmed not to produce 4-vinyl guaiacol. Whole genome sequencing revealed that this variant lost the subtelomeric region of the CHRXIII right arm that carried the two clustered genes SePAD1- SeFDC1 whose deletion in a naïve S. eubayanus strain (CBS 12357/FM1318) resulted in an identical phenotype. Subsequently, the Pof ^- variant was crossed with a Pof^-S. cerevisiae partner. The resulting hybrid was not able to convert ferulic acid demonstrating the undisputable value of the mutagenized variant HTSE042 to eventually construct S. cerevisiae × S. eubayanus hybrids with phenotypic characteristics of S. pastorianus.

Design of Experiments Methodology to Build a Multifactorial Statistical Model Describing the Metabolic Interactions of Alcohol Dehydrogenase Isozymes in the Ethanol Biosynthetic Pathway of the Yeast Saccharomyces cerevisiae

Multifactorial approaches can quickly and efficiently model complex, interacting natural or engineered biological systems in a way that traditional one-factor-at-a-time experimentation can fail to do. We applied a Design of Experiments (DOE) approach to model ethanol biosynthesis in yeast, which is well-understood and genetically tractable, yet complex. Six alcohol dehydrogenase (ADH) isozymes catalyze ethanol synthesis, differing in their transcriptional and post-translational regulation, subcellular localization, and enzyme kinetics. We generated a combinatorial library of all ADH gene deletions and measured the impact of gene deletion(s) and environmental context on ethanol production of a subset of this library. The data were used to build a statistical model that described known behaviors of ADH isozymes and identified novel interactions. Importantly, the model described features of ADH metabolic behavior without explicit a priori knowledge. The method is therefore highly suited to understanding and optimizing metabolic pathways in less well-understood systems.

Implementing CRISPR-Cas technologies in conventional and non-conventional yeasts: Current state and future prospects

Within five years, the CRISPR-Cas system has emerged as the dominating tool for genome engineering, while also changing the speed and efficiency of metabolic engineering in conventional (Saccharomyces cerevisiae and Schizosaccharomyces pombe) and non-conventional (Yarrowia lipolytica, Pichia pastoris syn. Komagataella phaffii, Kluyveromyces lactis, Candida albicans and C. glabrata) yeasts. Especially in S. cerevisiae, an extensive toolbox of advanced CRISPR-related applications has been established, including crisprTFs and gene drives. The comparison of innovative CRISPR-Cas expression strategies in yeasts presented here may also serve as guideline to implement and refine CRISPR-Cas systems for highly efficient genome editing in other eukaryotic organisms.

Low affinity uniporter carrier proteins can increase net substrate uptake rate by reducing efflux

Many organisms have several similar transporters with different affinities for the same substrate. Typically, high-affinity transporters are expressed when substrate is scarce and low-affinity ones when it is abundant. The benefit of using low instead of high-affinity transporters remains unclear, especially when additional nutrient sensors are present. Here, we investigate two hypotheses. It was previously hypothesized that there is a trade-off between the affinity and the catalytic efficiency of transporters, and we find some but no definitive support for it. Additionally, we propose that for uptake by facilitated diffusion, at saturating substrate concentrations, lowering the affinity enhances the net uptake rate by reducing substrate efflux. As a consequence, there exists an optimal, external-substrate-concentration dependent transporter affinity. A computational model of Saccharomyces cerevisiae glycolysis shows that using the low affinity HXT3 transporter instead of the high affinity HXT6 enhances the steady-state flux by 36%. We tried to test this hypothesis with yeast strains expressing a single glucose transporter modified to have either a high or a low affinity. However, due to the intimate link between glucose perception and metabolism, direct experimental proof for this hypothesis remained inconclusive. Still, our theoretical results provide a novel reason for the presence of low-affinity transport systems.

Balanced trafficking between the ER and the Golgi apparatus increases protein secretion in yeast

The yeast Saccharomyces cerevisiae is widely used as a cell factory to produce recombinant proteins. However, S. cerevisiae naturally secretes only a few proteins, such as invertase and the mating alpha factor, and its secretory capacity is limited. It has been reported that engineering protein anterograde trafficking from the endoplasmic reticulum to the Golgi apparatus by the moderate overexpression of SEC16 could increase recombinant protein secretion in S. cerevisiae. In this study, the retrograde trafficking in a strain with moderate overexpression of SEC16 was engineered by overexpression of ADP-ribosylation factor GTP activating proteins, Gcs1p and Glo3p, which are involved in the process of COPI-coated vesicle formation. Engineering the retrograde trafficking increased the secretion of α-amylase but did not induce production of reactive oxygen species. An expanded ER membrane was detected in both the GCS1 and GLO3 overexpression strains. Physiological characterizations during batch fermentation showed that GLO3 overexpression had better effect on recombinant protein secretion than GCS1 overexpression. Additionally, the GLO3 overexpression strain had higher secretion of two other recombinant proteins, endoglucanase I from Trichoderma reesei and glucan-1,4-α-glucosidase from Rhizopus oryzae, indicating overexpression of GLO3 in a SEC16 moderate overexpression strain might be a general strategy for improving production of secreted proteins by yeast.

Exploring D-xylose oxidation in Saccharomyces cerevisiae through the Weimberg pathway

Engineering of the yeast Saccharomyces cerevisiae towards efficient D-xylose assimilation has been a major focus over the last decades since D-xylose is the second most abundant sugar in nature, and its conversion into products could significantly improve process economy in biomass-based processes. Up to now, two different metabolic routes have been introduced via genetic engineering, consisting of either the isomerization or the oxido-reduction of D-xylose to D-xylulose that is further connected to the pentose phosphate pathway and glycolysis. In the present study, cytosolic D-xylose oxidation was investigated instead, through the introduction of the Weimberg pathway from Caulobacter crescentus in S. cerevisiae. This pathway consists of five reaction steps that connect D-xylose to the TCA cycle intermediate α-ketoglutarate. The corresponding genes could be expressed in S. cerevisiae, but no growth was observed on D-xylose indicating that not all the enzymes were functionally active. The accumulation of the Weimberg intermediate D-xylonate suggested that the dehydration step(s) might be limiting, blocking further conversion into α-ketoglutarate. Although four alternative dehydratases both of bacterial and archaeon origins were evaluated, D-xylonate accumulation still occurred. A better understanding of the mechanisms associated with the activity of dehydratases, both at a bacterial and yeast level, appears essential to obtain a fully functional Weimberg pathway in S. cerevisiae.

Engineering 1-Alkene Biosynthesis and Secretion by Dynamic Regulation in Yeast

Microbial production of fatty acid-derived hydrocarbons offers a great opportunity to sustainably supply biofuels and oleochemicals. One challenge is to achieve a high production rate. Besides, low efficiency in secretion will cause high separation costs, and it is therefore desirable to have product secretion. Here, we engineered the budding yeast Saccharomyces cerevisiae to produce and secrete 1-alkenes by manipulation of the fatty acid metabolism, enzyme selection, engineering the electron transfer system and expressing a transporter. Furthermore, we implemented a dynamic regulation strategy to control the expression of membrane enzyme and transporter, which improved 1-alkene production and cell growth by relieving the possible toxicity of overexpressed membrane proteins. With these efforts, the engineered yeast cell factory produced 35.3 mg/L 1-alkenes with more than 80% being secreted. This represents a 10-fold improvement compared with earlier reported hydrocarbon production by S. cerevisiae.

Acetamidase as a dominant recyclable marker for Komagataella phaffii strain engineering

We have investigated the use of the gene coding for acetamidase (amdS) as a recyclable dominant marker for the methylotrophic yeast Komagataella phaffii in order to broaden its genetic toolbox. First, the endogenous constitutive AMD2 gene (a putative acetamidase) was deleted generating strain LA1. A cassette (amdSloxP) was constructed bearing a codon-optimized version of the Aspergillus nidulans amdS gene flanked by loxP sites for marker excision with Cre recombinase. This cassette was successfully tested as a dominant selection marker for transformation of the LA1 strain after selection on plates containing acetamide as a sole nitrogen source. Finally, amdSloxP was used to sequentially disrupt the K. phaffii ADE2 and URA5 genes. After each disruption event, a Cre-mediated marker recycling step was performed by plating cells on medium containing fluoroacetamide. In conclusion, amdS proved to be a suitable tool for K. phaffii transformation and marker recycling thus providing a new antibiotic-free system for genetic manipulation of this yeast.

Optimizing anaerobic growth rate and fermentation kinetics in Saccharomyces cerevisiae strains expressing Calvin-cycle enzymes for improved ethanol yield

BACKGROUND

Reduction or elimination of by-product formation is of immediate economic relevance in fermentation processes for industrial bioethanol production with the yeast Saccharomyces cerevisiae. Anaerobic cultures of wild-type S. cerevisiae require formation of glycerol to maintain the intracellular NADH/NAD+ balance. Previously, functional expression of the Calvin-cycle enzymes ribulose-1,5-bisphosphate carboxylase (RuBisCO) and phosphoribulokinase (PRK) in S. cerevisiae was shown to enable reoxidation of NADH with CO2 as electron acceptor. In slow-growing cultures, this engineering strategy strongly decreased the glycerol yield, while increasing the ethanol yield on sugar. The present study explores engineering strategies to improve rates of growth and alcoholic fermentation in yeast strains that functionally express RuBisCO and PRK, while maximizing the positive impact on the ethanol yield.

RESULTS

Multi-copy integration of a bacterial-RuBisCO expression cassette was combined with expression of the Escherichia coli GroEL/GroES chaperones and expression of PRK from the anaerobically inducible DAN1 promoter. In anaerobic, glucose-grown bioreactor batch cultures, the resulting S. cerevisiae strain showed a 31% lower glycerol yield and a 31% lower specific growth rate than a non-engineered reference strain. Growth of the engineered strain in anaerobic, glucose-limited chemostat cultures revealed a negative correlation between its specific growth rate and the contribution of the Calvin-cycle enzymes to redox homeostasis. Additional deletion of GPD2, which encodes an isoenzyme of NAD+-dependent glycerol-3-phosphate dehydrogenase, combined with overexpression of the structural genes for enzymes of the non-oxidative pentose-phosphate pathway, yielded a CO2-reducing strain that grew at the same rate as a non-engineered reference strain in anaerobic bioreactor batch cultures, while exhibiting a 86% lower glycerol yield and a 15% higher ethanol yield.

CONCLUSIONS

The metabolic engineering strategy presented here enables an almost complete elimination of glycerol production in anaerobic, glucose-grown batch cultures of S. cerevisiae, with an associated increase in ethanol yield, while retaining near wild-type growth rates and a capacity for glycerol formation under osmotic stress. Using current genome-editing techniques, the required genetic modifications can be introduced in one or a few transformations. Evaluation of this concept in industrial strains and conditions is therefore a realistic next step towards its implementation for improving the efficiency of first- and second-generation bioethanol production.

The Penicillium chrysogenum transporter PcAraT enables high-affinity, glucose-insensitive l-arabinose transport in Saccharomyces cerevisiae

BACKGROUND

l-Arabinose occurs at economically relevant levels in lignocellulosic hydrolysates. Its low-affinity uptake via the Saccharomyces cerevisiae Gal2 galactose transporter is inhibited by d-glucose. Especially at low concentrations of l-arabinose, uptake is an important rate-controlling step in the complete conversion of these feedstocks by engineered pentose-metabolizing S. cerevisiae strains.

RESULTS

Chemostat-based transcriptome analysis yielded 16 putative sugar transporter genes in the filamentous fungus Penicillium chrysogenum whose transcript levels were at least threefold higher in l-arabinose-limited cultures than in d-glucose-limited and ethanol-limited cultures. Of five genes, that encoded putative transport proteins and showed an over 30-fold higher transcript level in l-arabinose-grown cultures compared to d-glucose-grown cultures, only one (Pc20g01790) restored growth on l-arabinose upon expression in an engineered l-arabinose-fermenting S. cerevisiae strain in which the endogenous l-arabinose transporter, GAL2, had been deleted. Sugar transport assays indicated that this fungal transporter, designated as PcAraT, is a high-affinity (K_m = 0.13 mM), high-specificity l-arabinose-proton symporter that does not transport d-xylose or d-glucose. An l-arabinose-metabolizing S. cerevisiae strain in which GAL2 was replaced by PcaraT showed 450-fold lower residual substrate concentrations in l-arabinose-limited chemostat cultures than a congenic strain in which l-arabinose import depended on Gal2 (4.2 × 10^-3 and 1.8 g L^-1, respectively). Inhibition of l-arabinose transport by the most abundant sugars in hydrolysates, d-glucose and d-xylose was far less pronounced than observed with Gal2. Expression of PcAraT in a hexose-phosphorylation-deficient, l-arabinose-metabolizing S. cerevisiae strain enabled growth in media supplemented with both 20 g L^-1 l-arabinose and 20 g L^-1 d-glucose, which completely inhibited growth of a congenic strain in the same condition that depended on l-arabinose transport via Gal2.

CONCLUSION

Its high affinity and specificity for l-arabinose, combined with limited sensitivity to inhibition by d-glucose and d-xylose, make PcAraT a valuable transporter for application in metabolic engineering strategies aimed at engineering S. cerevisiae strains for efficient conversion of lignocellulosic hydrolysates.

CRISPR-Cas9 mediated gene deletions in lager yeast Saccharomyces pastorianus

BACKGROUND

The ease of use of CRISPR-Cas9 reprogramming, its high efficacy, and its multiplexing capabilities have brought this technology at the forefront of genome editing techniques. Saccharomyces pastorianus is an aneuploid interspecific hybrid of Saccharomyces cerevisiae and Saccharomyces eubayanus that has been domesticated for centuries and is used for the industrial fermentation of lager beer. For yet uncharacterised reasons, this hybrid yeast is far more resilient to genetic alteration than its ancestor S. cerevisiae.

RESULTS

This study reports a new CRISPR-Cas9 method for accurate gene deletion in S. pastorianus. This method combined the Streptococcus pyogenes cas9 gene expressed from either a chromosomal locus or from a mobile genetic element in combination with a plasmid-borne gRNA expression cassette. While the well-established gRNA expression system using the RNA polymerase III dependent SNR52 promoter failed, expression of a gRNA flanked with Hammerhead and Hepatitis Delta Virus ribozymes using the RNA polymerase II dependent TDH3 promoter successfully led to accurate deletion of all four alleles of the SeILV6 gene in strain CBS1483. Furthermore the expression of two ribozyme-flanked gRNAs separated by a 10-bp linker in a polycistronic array successfully led to the simultaneous deletion of SeATF1 and SeATF2, genes located on two separate chromosomes. The expression of this array resulted in the precise deletion of all five and four alleles mediated by homologous recombination in the strains CBS1483 and Weihenstephan 34/70 respectively, demonstrating the multiplexing abilities of this gRNA expression design.

CONCLUSIONS

These results firmly established that CRISPR-Cas9 significantly facilitates and accelerates genome editing in S. pastorianus.

Combined engineering of disaccharide transport and phosphorolysis for enhanced ATP yield from sucrose fermentation in Saccharomyces cerevisiae

Anaerobic industrial fermentation processes do not require aeration and intensive mixing and the accompanying cost savings are beneficial for production of chemicals and fuels. However, the free-energy conservation of fermentative pathways is often insufficient for the production and export of the desired compounds and/or for cellular growth and maintenance. To increase free-energy conservation during fermentation of the industrially relevant disaccharide sucrose by Saccharomyces cerevisiae, we first replaced the native yeast α-glucosidases by an intracellular sucrose phosphorylase from Leuconostoc mesenteroides (LmSPase). Subsequently, we replaced the native proton-coupled sucrose uptake system by a putative sucrose facilitator from Phaseolus vulgaris (PvSUF1). The resulting strains grew anaerobically on sucrose at specific growth rates of 0.09 ± 0.02h^-1 (LmSPase) and 0.06 ± 0.01h^-1 (PvSUF1, LmSPase). Overexpression of the yeast PGM2 gene, which encodes phosphoglucomutase, increased anaerobic growth rates on sucrose of these strains to 0.23 ± 0.01h^-1 and 0.08 ± 0.00h^-1, respectively. Determination of the biomass yield in anaerobic sucrose-limited chemostat cultures was used to assess the free-energy conservation of the engineered strains. Replacement of intracellular hydrolase with a phosphorylase increased the biomass yield on sucrose by 31%. Additional replacement of the native proton-coupled sucrose uptake system by PvSUF1 increased the anaerobic biomass yield by a further 8%, resulting in an overall increase of 41%. By experimentally demonstrating an energetic benefit of the combined engineering of disaccharide uptake and cleavage, this study represents a first step towards anaerobic production of compounds whose metabolic pathways currently do not conserve sufficient free-energy.

Efficient protein production by yeast requires global tuning of metabolism

The biotech industry relies on cell factories for production of pharmaceutical proteins, of which several are among the top-selling medicines. There is, therefore, considerable interest in improving the efficiency of protein production by cell factories. Protein secretion involves numerous intracellular processes with many underlying mechanisms still remaining unclear. Here, we use RNA-seq to study the genome-wide transcriptional response to protein secretion in mutant yeast strains. We find that many cellular processes have to be attuned to support efficient protein secretion. In particular, altered energy metabolism resulting in reduced respiration and increased fermentation, as well as balancing of amino-acid biosynthesis and reduced thiamine biosynthesis seem to be particularly important. We confirm our findings by inverse engineering and physiological characterization and show that by tuning metabolism cells are able to efficiently secrete recombinant proteins. Our findings provide increased understanding of which cellular regulations and pathways are associated with efficient protein secretion.

The contribution of metabolic pathways to protein secretion is largely unknown. Here, the authors find conserved metabolic patterns in yeast by examining genome-wide transcriptional responses in high protein secretion mutants and reveal critical factors that can be tuned for efficient protein secretion.

Elimination of sucrose transport and hydrolysis in Saccharomyces cerevisiae: a platform strain for engineering sucrose metabolism

Many relevant options to improve efficacy and kinetics of sucrose metabolism in Saccharomyces cerevisiae and, thereby, the economics of sucrose-based processes remain to be investigated. An essential first step is to identify all native sucrose-hydrolysing enzymes and sucrose transporters in this yeast, including those that can be activated by suppressor mutations in sucrose-negative strains. A strain in which all known sucrose-transporter genes (MAL11, MAL21, MAL31, MPH2, MPH3) were deleted did not grow on sucrose after 2 months of incubation. In contrast, a strain with deletions in genes encoding sucrose-hydrolysing enzymes (SUC2, MAL12, MAL22, MAL32) still grew on sucrose. Its specific growth rate increased from 0.08 to 0.25 h⁻¹ after sequential batch cultivation. This increase was accompanied by a 3-fold increase of in vitro sucrose-hydrolysis and isomaltase activities, as well as by a 3- to 5-fold upregulation of the isomaltase-encoding genes IMA1 and IMA5. One-step Cas9-mediated deletion of all isomaltase-encoding genes (IMA1-5) completely abolished sucrose hydrolysis. Even after 2 months of incubation, the resulting strain did not grow on sucrose. This sucrose-negative strain can be used as a platform to test metabolic engineering strategies and for fundamental studies into sucrose hydrolysis or transport.

Laboratory Evolution of a Biotin-Requiring Saccharomyces cerevisiae Strain for Full Biotin Prototrophy and Identification of Causal Mutations

Biotin prototrophy is a rare, incompletely understood, and industrially relevant characteristic of Saccharomyces cerevisiae strains. The genome of the haploid laboratory strain CEN.PK113-7D contains a full complement of biotin biosynthesis genes, but its growth in biotin-free synthetic medium is extremely slow (specific growth rate [μ] ≈ 0.01 h^-1). Four independent evolution experiments in repeated batch cultures and accelerostats yielded strains whose growth rates (μ ≤ 0.36 h^-1) in biotin-free and biotin-supplemented media were similar. Whole-genome resequencing of these evolved strains revealed up to 40-fold amplification of BIO1, which encodes pimeloyl-coenzyme A (CoA) synthetase. The additional copies of BIO1 were found on different chromosomes, and its amplification coincided with substantial chromosomal rearrangements. A key role of this gene amplification was confirmed by overexpression of BIO1 in strain CEN.PK113-7D, which enabled growth in biotin-free medium (μ = 0.15 h^-1). Mutations in the membrane transporter genes TPO1 and/or PDR12 were found in several of the evolved strains. Deletion of TPO1 and PDR12 in a BIO1-overexpressing strain increased its specific growth rate to 0.25 h^-1 The effects of null mutations in these genes, which have not been previously associated with biotin metabolism, were nonadditive. This study demonstrates that S. cerevisiae strains that carry the basic genetic information for biotin synthesis can be evolved for full biotin prototrophy and identifies new targets for engineering biotin prototrophy into laboratory and industrial strains of this yeast.IMPORTANCE Although biotin (vitamin H) plays essential roles in all organisms, not all organisms can synthesize this vitamin. Many strains of baker's yeast, an important microorganism in industrial biotechnology, contain at least some of the genes required for biotin synthesis. However, most of these strains cannot synthesize biotin at all or do so at rates that are insufficient to sustain fast growth and product formation. Consequently, this expensive vitamin is routinely added to baker's yeast cultures. In this study, laboratory evolution in biotin-free growth medium yielded new strains that grew as fast in the absence of biotin as in its presence. By analyzing the DNA sequences of evolved biotin-independent strains, mutations were identified that contributed to this ability. This work demonstrates full biotin independence of an industrially relevant yeast and identifies mutations whose introduction into other yeast strains may reduce or eliminate their biotin requirements.

A method for labeling proteins with tags at the native genomic loci in budding yeast

Fluorescent proteins and epitope tags are often used as protein fusion tags to study target proteins. One prevailing technique in the budding yeast Saccharomyces cerevisiae is to fuse these tags to a target gene at the precise chromosomal location via homologous recombination. However, several limitations hamper the application of this technique, such as the selectable markers not being reusable, tagging of only the C-terminal being possible, and a "scar" sequence being left in the genome. Here, we describe a strategy to solve these problems by tagging target genes based on a pop-in/pop-out and counter-selection system. Three fluorescent protein tag (mCherry, sfGFP, and mKikGR) and two epitope tag (HA and 3×FLAG) constructs were developed and utilized to tag HHT1, UBC13 or RAD5 at the chromosomal locus as proof-of-concept.

Metabolic engineering strategies for optimizing acetate reduction, ethanol yield and osmotolerance in Saccharomyces cerevisiae

BACKGROUND

Glycerol, whose formation contributes to cellular redox balancing and osmoregulation in Saccharomyces cerevisiae, is an important by-product of yeast-based bioethanol production. Replacing the glycerol pathway by an engineered pathway for NAD⁺-dependent acetate reduction has been shown to improve ethanol yields and contribute to detoxification of acetate-containing media. However, the osmosensitivity of glycerol non-producing strains limits their applicability in high-osmolarity industrial processes. This study explores engineering strategies for minimizing glycerol production by acetate-reducing strains, while retaining osmotolerance.

RESULTS

GPD2 encodes one of two S. cerevisiae isoenzymes of NAD⁺-dependent glycerol-3-phosphate dehydrogenase (G3PDH). Its deletion in an acetate-reducing strain yielded a fourfold lower glycerol production in anaerobic, low-osmolarity cultures but hardly affected glycerol production at high osmolarity. Replacement of both native G3PDHs by an archaeal NADP⁺-preferring enzyme, combined with deletion of ALD6, yielded an acetate-reducing strain the phenotype of which resembled that of a glycerol-negative gpd1Δ gpd2Δ strain in low-osmolarity cultures. This strain grew anaerobically at high osmolarity (1 mol L^-1 glucose), while consuming acetate and producing virtually no extracellular glycerol. Its ethanol yield in high-osmolarity cultures was 13% higher than that of an acetate-reducing strain expressing the native glycerol pathway.

CONCLUSIONS

Deletion of GPD2 provides an attractive strategy for improving product yields of acetate-reducing S. cerevisiae strains in low, but not in high-osmolarity media. Replacement of the native yeast G3PDHs by a heterologous NADP⁺-preferring enzyme, combined with deletion of ALD6, virtually eliminated glycerol production in high-osmolarity cultures while enabling efficient reduction of acetate to ethanol. After further optimization of growth kinetics, this strategy for uncoupling the roles of glycerol formation in redox homeostasis and osmotolerance can be applicable for improving performance of industrial strains in high-gravity acetate-containing processes.

Mutations in PMR1 stimulate xylose isomerase activity and anaerobic growth on xylose of engineered Saccharomyces cerevisiae by influencing manganese homeostasis

Combined overexpression of xylulokinase, pentose-phosphate-pathway enzymes and a heterologous xylose isomerase (XI) is required but insufficient for anaerobic growth of Saccharomyces cerevisiae on d-xylose. Single-step Cas9-assisted implementation of these modifications yielded a yeast strain expressing Piromyces XI that showed fast aerobic growth on d-xylose. However, anaerobic growth required a 12-day adaptation period. Xylose-adapted cultures carried mutations in PMR1, encoding a Golgi Ca2+/Mn2+ ATPase. Deleting PMR1 in the parental XI-expressing strain enabled instantaneous anaerobic growth on d-xylose. In pmr1 strains, intracellular Mn2+ concentrations were much higher than in the parental strain. XI activity assays in cell extracts and reconstitution experiments with purified XI apoenzyme showed superior enzyme kinetics with Mn2+ relative to other divalent metal ions. This study indicates engineering of metal homeostasis as a relevant approach for optimization of metabolic pathways involving metal-dependent enzymes. Specifically, it identifies metal interactions of heterologous XIs as an underexplored aspect of engineering xylose metabolism in yeast.

Introducing MX Cassettes into Saccharomyces cerevisiae

The Saccharomyces cerevisiae genome can be readily and precisely modified with the use of knock out (KO) marker cassettes to delete genes. The most frequently used family of KO cassettes is the MX cassettes. This protocol describes how to use the different types of MX cassettes by selecting for prototrophy, utilization of cytosine or acetamide as a sole nitrogen source, or resistance to one of six different drugs.

Popping Out MX Cassettes from Saccharomyces cerevisiae

MX cassettes are frequently used to generate knockout (KO) mutations in Saccharomyces cerevisiae. The recycling or "popping out" of an MX cassette flanked by direct repeats allows the same cassette to be reused in a strain to generate additional KO mutations. Popping out MX cassettes also eliminates MX homology in a strain, which facilitates the subsequent generation of additional KO mutations with other MX cassettes. MX cassettes can be recycled or "popped out" of the genome by spontaneous recombination between large, cassette-borne MX3 or PR direct repeats and by Cre-mediated, site-specific recombination between small, cassette-borne loxP direct repeats. Both of these techniques leave a mutation with a cassette-encoded "scar." For the URA3MX, LYS5MX, FCA1/FCY1MX, and amdSYM cassettes, there are counterselections. Counterselections are extremely useful as they allow for positive selection for plasmid shuffling, transplacement of mutant alleles into the genome, and recycling or popping out cassettes flanked by cassette-encoded direct repeats to yield mutations with a cassette-encoded scar. Finally, after amplifying with the appropriately designed primers, integrated counterselectable MX cassettes can be popped out to generate seamless or "scar-free" deletion mutations, as well as indel and point mutations.

MX Cassettes for Knocking Out Genes in Yeast

Precise modifications of the Saccharomyces cerevisiae genome use marker cassettes, most often in the form of "knockout" (KO) marker cassettes, to delete genes. Many different KO marker cassettes exist, some of which require strains with specific genotypes, such as auxotrophic mutations, and others that have no strain genotype requirements, such as selections for drug resistance and one of two selections for nitrogen source utilization. This introduction focuses on the most frequently used family of KO cassettes-the MX cassettes. In particular, we focus on and describe the different types of MX cassettes and selections; specifically, selections for prototrophy; selections for utilization of cytosine or acetamide as sole nitrogen sources; and selections for resistance to six different drugs. The use of cassettes to place genes under regulated control is briefly discussed. Also discussed are strain genotype requirements (where applicable); media requirements; how to "recycle" or "pop out" cassettes; and counterselections against specific KO cassettes.

Quantitative Trait Loci (QTL)-Guided Metabolic Engineering of a Complex Trait

Engineering complex phenotypes for industrial and synthetic biology applications is difficult and often confounds rational design. Bioethanol production from lignocellulosic feedstocks is a complex trait that requires multiple host systems to utilize, detoxify, and metabolize a mixture of sugars and inhibitors present in plant hydrolysates. Here, we demonstrate an integrated approach to discovering and optimizing host factors that impact fitness of Saccharomyces cerevisiae during fermentation of a Miscanthus x giganteus plant hydrolysate. We first used high-resolution Quantitative Trait Loci (QTL) mapping and systematic bulk Reciprocal Hemizygosity Analysis (bRHA) to discover 17 loci that differentiate hydrolysate tolerance between an industrially related (JAY291) and a laboratory (S288C) strain. We then used this data to identify a subset of favorable allelic loci that were most amenable for strain engineering. Guided by this "genetic blueprint", and using a dual-guide Cas9-based method to efficiently perform multikilobase locus replacements, we engineered an S288C-derived strain with superior hydrolysate tolerance than JAY291. Our methods should be generalizable to engineering any complex trait in S. cerevisiae, as well as other organisms.

Novel methods to optimize gene and statistic test for evaluation - an application for Escherichia coli

Background

Since the recombinant protein was discovered, it has become more popular in many aspects of life science. The value of global pharmaceutical market was $87 billion in 2008 and the sales for industrial enzyme exceeded $4 billion in 2012. This is strong evidence showing the great potential of recombinant protein. However, native genes introduced into a host can cause incompatibility of codon usage bias, GC content, repeat region, Shine-Dalgarno sequence with host’s expression system, so the yields can fall down significantly. Hence, we propose novel methods for gene optimization based on neural network, Bayesian theory, and Euclidian distance.

Result

The correlation coefficients of our neural network are 0.86, 0.73, and 0.90 in training, validation, and testing process. In addition, genes optimized by our methods seem to associate with highly expressed genes and give reasonable codon adaptation index values. Furthermore, genes optimized by the proposed methods are highly matched with the previous experimental data.

Conclusion

The proposed methods have high potential for gene optimization and further researches in gene expression. We built a demonstrative program using Matlab R2014a under Mac OS X. The program was published in both standalone executable program and Matlab function files. The developed program can be accessed from http://www.math.hcmus.edu.vn/~ptbao/paper_soft/GeneOptProg/.

Synthetic gene design-The rationale for codon optimization and implications for molecular pharming in plants

Degeneracy in the genetic code allows multiple codon sequences to encode the same protein. Codon usage bias in genes is the term given to the preferred use of particular synonymous codons. Synonymous codon substitutions had been regarded as "silent" as the primary structure of the protein was not affected; however, it is now accepted that synonymous substitutions can have a significant effect on heterologous protein expression. Codon optimization, the process of altering codons within the gene sequence to improve recombinant protein expression, has become widely practised. Multiple inter-linked factors affecting protein expression need to be taken into consideration when optimizing a gene sequence. Over the years, various computer programmes have been developed to aid in the gene sequence optimization process. However, as the rulebook for altering codon usage to affect protein expression is still not completely understood, it is difficult to predict which strategy, if any, will design the "optimal" gene sequence. In this review, codon usage bias and factors affecting codon selection will be discussed and the evidence for codon optimization impact will be reviewed for recombinant protein expression using plants as a case study. These developments will be relevant to all recombinant expression systems; however, molecular pharming in plants is an area which has consistently encountered difficulties with low levels of recombinant protein expression, and should benefit from an evidence based rational approach to synthetic gene design. Biotechnol. Bioeng. 2017;114: 492-502. © 2016 Wiley Periodicals, Inc.

Pathway swapping: Toward modular engineering of essential cellular processes

Recent developments in synthetic biology enable one-step implementation of entire metabolic pathways in industrial microorganisms. A similarly radical remodelling of central metabolism could greatly accelerate fundamental and applied research, but is impeded by the mosaic organization of microbial genomes. To eliminate this limitation, we propose and explore the concept of "pathway swapping," using yeast glycolysis as the experimental model. Construction of a "single-locus glycolysis" Saccharomyces cerevisiae platform enabled quick and easy replacement of this yeast's entire complement of 26 glycolytic isoenzymes by any alternative, functional glycolytic pathway configuration. The potential of this approach was demonstrated by the construction and characterization of S. cerevisiae strains whose growth depended on two nonnative glycolytic pathways: a complete glycolysis from the related yeast Saccharomyces kudriavzevii and a mosaic glycolysis consisting of yeast and human enzymes. This work demonstrates the feasibility and potential of modular, combinatorial approaches to engineering and analysis of core cellular processes.

Production of fatty acid-derived oleochemicals and biofuels by synthetic yeast cell factories

Sustainable production of oleochemicals requires establishment of cell factory platform strains. The yeast Saccharomyces cerevisiae is an attractive cell factory as new strains can be rapidly implemented into existing infrastructures such as bioethanol production plants. Here we show high-level production of free fatty acids (FFAs) in a yeast cell factory, and the production of alkanes and fatty alcohols from its descendants. The engineered strain produces up to 10.4 g l(-1) of FFAs, which is the highest reported titre to date. Furthermore, through screening of specific pathway enzymes, endogenous alcohol dehydrogenases and aldehyde reductases, we reconstruct efficient pathways for conversion of fatty acids to alkanes (0.8 mg l(-1)) and fatty alcohols (1.5 g l(-1)), to our knowledge the highest titres reported in S. cerevisiae. This should facilitate the construction of yeast cell factories for production of fatty acids derived products and even aldehyde-derived chemicals of high value.

Improving ethanol yield in acetate-reducing Saccharomyces cerevisiae by cofactor engineering of 6-phosphogluconate dehydrogenase and deletion of ALD6

BACKGROUND

Acetic acid, an inhibitor of sugar fermentation by yeast, is invariably present in lignocellulosic hydrolysates which are used or considered as feedstocks for yeast-based bioethanol production. Saccharomyces cerevisiae strains have been constructed, in which anaerobic reduction of acetic acid to ethanol replaces glycerol formation as a mechanism for reoxidizing NADH formed in biosynthesis. An increase in the amount of acetate that can be reduced to ethanol should further decrease acetic acid concentrations and enable higher ethanol yields in industrial processes based on lignocellulosic feedstocks. The stoichiometric requirement of acetate reduction for NADH implies that increased generation of NADH in cytosolic biosynthetic reactions should enhance acetate consumption.

RESULTS

Replacement of the native NADP(+)-dependent 6-phosphogluconate dehydrogenase in S. cerevisiae by a prokaryotic NAD(+)-dependent enzyme resulted in increased cytosolic NADH formation, as demonstrated by a ca. 15% increase in the glycerol yield on glucose in anaerobic cultures. Additional deletion of ALD6, which encodes an NADP(+)-dependent acetaldehyde dehydrogenase, led to a 39% increase in the glycerol yield compared to a non-engineered strain. Subsequent replacement of glycerol formation by an acetate reduction pathway resulted in a 44% increase of acetate consumption per amount of biomass formed, as compared to an engineered, acetate-reducing strain that expressed the native 6-phosphogluconate dehydrogenase and ALD6. Compared to a non-acetate reducing reference strain under the same conditions, this resulted in a ca. 13% increase in the ethanol yield on glucose.

CONCLUSIONS

The combination of NAD(+)-dependent 6-phosphogluconate dehydrogenase expression and deletion of ALD6 resulted in a marked increase in the amount of acetate that was consumed in these proof-of-principle experiments, and this concept is ready for further testing in industrial strains as well as in hydrolysates. Altering the cofactor specificity of the oxidative branch of the pentose-phosphate pathway in S. cerevisiae can also be used to increase glycerol production in wine fermentation and to improve NADH generation and/or generation of precursors derived from the pentose-phosphate pathway in other industrial applications of this yeast.

Development of a Candida glabrata dominant nutritional transformation marker utilizing the Aspergillus nidulans acetamidase gene (amdS)

The gene encoding Aspergillus nidulans acetamidase (amdS) was placed under control of Candida albicans ACT1 promoter and terminator sequences and then cloned into a plasmid containing C. glabrata ARS10,CEN8 or ARS10+CEN8 sequences. All plasmids transformed C. glabrata wild-type cells to acetamide+, with the ARS-only containing plasmid transforming cells at the highest frequencies (>1.0 × 10(4) transformants μg(-1)). Plasmids were rapidly lost under non-selective conditions with the frequency dependent on chromosomal element, thus recycling the acetamide- phenotype. The amdS plasmid was used to transform a set of clinical isolates resistant to a variety of antifungal drugs. All strains were successfully transformed to the acetamide+ phenotype at high frequency, confirming that this plasmid construct could be used as a simple dominant marker on virtually any strain. Gap repair experiments demonstrated that just as in Saccharomyces cerevisiae, gap repair functions efficiently inC. glabrata, suggesting that C. glabrata has numerous similarities toS. cerevisiae with regard to ease of molecular manipulation. The amdS system is inexpensive and efficient, and combined with existing C. glabrata plasmid elements, confers a high transformation frequency for C. glabrata with a phenotype that can be easily recycled.

Alternative reactions at the interface of glycolysis and citric acid cycle in Saccharomyces cerevisiae

Pyruvate and acetyl-coenzyme A, located at the interface between glycolysis and TCA cycle, are important intermediates in yeast metabolism and key precursors for industrially relevant products. Rational engineering of their supply requires knowledge of compensatory reactions that replace predominant pathways when these are inactivated. This study investigates effects of individual and combined mutations that inactivate the mitochondrial pyruvate-dehydrogenase (PDH) complex, extramitochondrial citrate synthase (Cit2) and mitochondrial CoA-transferase (Ach1) in Saccharomyces cerevisiae. Additionally, strains with a constitutively expressed carnitine shuttle were constructed and analyzed. A predominant role of the PDH complex in linking glycolysis and TCA cycle in glucose-grown batch cultures could be functionally replaced by the combined activity of the cytosolic PDH bypass and Cit2. Strongly impaired growth and a high incidence of respiratory deficiency in pda1Δ ach1Δ strains showed that synthesis of intramitochondrial acetyl-CoA as a metabolic precursor requires activity of either the PDH complex or Ach1. Constitutive overexpression of AGP2, HNM1, YAT2, YAT1, CRC1 and CAT2 enabled the carnitine shuttle to efficiently link glycolysis and TCA cycle in l-carnitine-supplemented, glucose-grown batch cultures. Strains in which all known reactions at the glycolysis-TCA cycle interface were inactivated still grew slowly on glucose, indicating additional flexibility at this key metabolic junction.

Replacement of the initial steps of ethanol metabolism in Saccharomyces cerevisiae by ATP-independent acetylating acetaldehyde dehydrogenase

In Saccharomyces cerevisiae ethanol dissimilation is initiated by its oxidation and activation to cytosolic acetyl-CoA. The associated consumption of ATP strongly limits yields of biomass and acetyl-CoA-derived products. Here, we explore the implementation of an ATP-independent pathway for acetyl-CoA synthesis from ethanol that, in theory, enables biomass yield on ethanol that is up to 40% higher. To this end, all native yeast acetaldehyde dehydrogenases (ALDs) were replaced by heterologous acetylating acetaldehyde dehydrogenase (A-ALD). Engineered Ald⁻ strains expressing different A-ALDs did not immediately grow on ethanol, but serial transfer in ethanol-grown batch cultures yielded growth rates of up to 70% of the wild-type value. Mutations in ACS1 were identified in all independently evolved strains and deletion of ACS1 enabled slow growth of non-evolved Ald⁻ A-ALD strains on ethanol. Acquired mutations in A-ALD genes improved affinity—V_max/K_m for acetaldehyde. One of five evolved strains showed a significant 5% increase of its biomass yield in ethanol-limited chemostat cultures. Increased production of acetaldehyde and other by-products was identified as possible cause for lower than theoretically predicted biomass yields. This study proves that the native yeast pathway for conversion of ethanol to acetyl-CoA can be replaced by an engineered pathway with the potential to improve biomass and product yields.

This manuscript investigates a metabolic engineering strategy to improve the use of ethanol as a feedstock for production of bio-based fuels and chemicals with yeast.

CRISPR/Cas9 advances engineering of microbial cell factories

One of the key drivers for successful metabolic engineering in microbes is the efficacy by which genomes can be edited. As such there are many methods to choose from when aiming to modify genomes, especially those of model organisms like yeast and bacteria. In recent years, clustered regularly interspaced palindromic repeats (CRISPR) and its associated proteins (Cas) have become the method of choice for precision genome engineering in many organisms due to their orthogonality, versatility and efficacy. Here we review the strategies adopted for implementation of RNA-guided CRISPR/Cas9 genome editing with special emphasis on their application for metabolic engineering of yeast and bacteria. Also, examples of how nuclease-deficient Cas9 has been applied for RNA-guided transcriptional regulation of target genes will be reviewed, as well as tools available for computer-aided design of guide-RNAs will be highlighted. Finally, this review will provide a perspective on the immediate challenges and opportunities foreseen by the use of CRISPR/Cas9 genome engineering and regulation in the context of metabolic engineering.

Comparative assessment of native and heterologous 2-oxo acid decarboxylases for application in isobutanol production by Saccharomyces cerevisiae

BACKGROUND

Decarboxylation of α-ketoisovalerate to isobutyraldehyde is a key reaction in metabolic engineering of Saccharomyces cerevisiae for isobutanol production with published studies relying on overexpression of either the native ARO10 gene or of the Lactococcus lactis kivD decarboxylase gene resulting in low enzymatic activities. Here, we compare relevant properties for isobutanol production of Aro10, KivD and an additional, less studied, L. lactis decarboxylase KdcA.

RESULTS

To eliminate interference by native decarboxylases, each 2-oxo acid decarboxylase was overexpressed in a 'decarboxylase-negative' (pdc1Δ pdc5Δ pdc6Δ aro10Δ) S. cerevisiae background. Kinetic analyses in cell extracts revealed a superior V max/K m ratio of KdcA for α-ketoisovalerate and a wide range of linear and branched-chain 2-oxo acids. However, KdcA also showed the highest activity with pyruvate which, in engineered strains, can contribute to formation of ethanol as a by-product. Removal of native decarboxylase genes eliminated growth on valine as sole nitrogen source and subsequent complementation of this growth impairment by expression of each decarboxylase indicated that based on the increased growth rate, the in vivo activity of KdcA with α-ketoisovalerate was higher than that of KivD and Aro10. Moreover, during oxygen-limited incubation in the presence of glucose, strains expressing kdcA or kivD showed a ca. twofold higher in vivo rate of conversion of α-ketoisovalerate into isobutanol than an ARO10-expressing strain. Finally, cell extracts from cultures grown on different nitrogen sources revealed increased activity of constitutively expressed KdcA after growth on both valine and phenylalanine, while KivD and Aro10 activity was only increased after growth on phenylalanine suggesting a difference in the regulation of these enzymes.

CONCLUSIONS

This study illustrates important differences in substrate specificity, enzyme kinetics and functional expression between different decarboxylases in the context of isobutanol production and identifies KdcA as a promising alternative decarboxylase not only for isobutanol production but also for other branched-chain and linear alcohols.

EasyClone 2.0: expanded toolkit of integrative vectors for stable gene expression in industrial Saccharomyces cerevisiae strains

Saccharomyces cerevisiae is one of the key cell factories for production of chemicals and active pharmaceuticals. For large-scale fermentations, particularly in biorefinery applications, it is desirable to use stress-tolerant industrial strains. However, such strains are less amenable for metabolic engineering than the standard laboratory strains. To enable easy delivery and overexpression of genes in a wide range of industrial S. cerevisiae strains, we constructed a set of integrative vectors with long homology arms and dominant selection markers. The vectors integrate into previously validated chromosomal locations via double cross-over and result in homogenous stable expression of the integrated genes, as shown for several unrelated industrial strains. Cre-mediated marker rescue is possible for removing markers positioned on different chromosomes. To demonstrate the applicability of the presented vector set for metabolic engineering of industrial yeast, we constructed xylose-utilizing strains overexpressing xylose isomerase, xylose transporter and five genes of the pentose phosphate pathway.

Flexible metabolic pathway construction using modular and divisible selection gene regulators

Genetic selections are important to biological engineering. Although selectable traits are limited, currently each trait only permits simultaneous introduction of a single DNA fragment. Complex pathway and strain construction however depends on rapid, combinatorial introduction of many genes that encode putative pathway candidates and homologs. To triple the utility of existing selection genes, we have developed divisible selection in Saccharomyces cerevisiae. Here, independent DNA fragments can be introduced and selected for simultaneously using a set of split hybrid transcription factors composed of parts from Escherichia coli LexA and Herpes simplex VP16 to regulate one single selectable phenotype of choice. Only when co-expressed, these split hybrid transcription factors promote transcription of a selection gene, causing tight selection of transformants containing all desired DNA fragments. Upon transformation, 94% of the selected colonies resulted strictly from transforming all three modules based on ARS/CEN plasmids. Similarly when used for chromosome integration, 95% of the transformants contained all three modules. The divisible selection system acts dominantly and thus expands selection gene utility from one to three without any genomic pre-modifications of the strain. We demonstrate the approach by introducing the fungal rubrofusarin polyketide pathway at a gene load of 11 kb distributed on three different plasmids, using a single selection trait and one yeast transformation step. By tripling the utility of existing selection genes, the employment of divisible selection improves flexibility and freedom in the strain engineering process.

Adaptive mutations in sugar metabolism restore growth on glucose in a pyruvate decarboxylase negative yeast strain

Background

A Saccharomyces cerevisiae strain carrying deletions in all three pyruvate decarboxylase (PDC) genes (also called Pdc negative yeast) represents a non-ethanol producing platform strain for the production of pyruvate derived biochemicals. However, it cannot grow on glucose as the sole carbon source, and requires supplementation of C₂ compounds to the medium in order to meet the requirement for cytosolic acetyl-CoA for biosynthesis of fatty acids and ergosterol.

Results

In this study, a Pdc negative strain was adaptively evolved for improved growth in glucose medium via serial transfer, resulting in three independently evolved strains, which were able to grow in minimal medium containing glucose as the sole carbon source at the maximum specific rates of 0.138, 0.148, 0.141 h⁻¹, respectively. Several genetic changes were identified in the evolved Pdc negative strains by genomic DNA sequencing. Among these genetic changes, 4 genes were found to carry point mutations in at least two of the evolved strains: MTH1 encoding a negative regulator of the glucose-sensing signal transduction pathway, HXT2 encoding a hexose transporter, CIT1 encoding a mitochondrial citrate synthase, and RPD3 encoding a histone deacetylase. Reverse engineering of the non-evolved Pdc negative strain through introduction of the MTH1^81D allele restored its growth on glucose at a maximum specific rate of 0.053 h⁻¹ in minimal medium with 2% glucose, and the CIT1 deletion in the reverse engineered strain further increased the maximum specific growth rate to 0.069 h⁻¹.

Conclusions

In this study, possible evolving mechanisms of Pdc negative strains on glucose were investigated by genome sequencing and reverse engineering. The non-synonymous mutations in MTH1 alleviated the glucose repression by repressing expression of several hexose transporter genes. The non-synonymous mutations in HXT2 and CIT1 may function in the presence of mutated MTH1 alleles and could be related to an altered central carbon metabolism in order to ensure production of cytosolic acetyl-CoA in the Pdc negative strain.

Electronic supplementary material

The online version of this article (doi:10.1186/s12934-015-0305-6) contains supplementary material, which is available to authorized users.

A Minimal Set of Glycolytic Genes Reveals Strong Redundancies in Saccharomyces cerevisiae Central Metabolism

As a result of ancestral whole-genome and small-scale duplication events, the genomes of Saccharomyces cerevisiae and many eukaryotes still contain a substantial fraction of duplicated genes. In all investigated organisms, metabolic pathways, and more particularly glycolysis, are specifically enriched for functionally redundant paralogs. In ancestors of the Saccharomyces lineage, the duplication of glycolytic genes is purported to have played an important role leading to S. cerevisiae's current lifestyle favoring fermentative metabolism even in the presence of oxygen and characterized by a high glycolytic capacity. In modern S. cerevisiae strains, the 12 glycolytic reactions leading to the biochemical conversion from glucose to ethanol are encoded by 27 paralogs. In order to experimentally explore the physiological role of this genetic redundancy, a yeast strain with a minimal set of 14 paralogs was constructed (the "minimal glycolysis" [MG] strain). Remarkably, a combination of a quantitative systems approach and semiquantitative analysis in a wide array of growth environments revealed the absence of a phenotypic response to the cumulative deletion of 13 glycolytic paralogs. This observation indicates that duplication of glycolytic genes is not a prerequisite for achieving the high glycolytic fluxes and fermentative capacities that are characteristic of S. cerevisiae and essential for many of its industrial applications and argues against gene dosage effects as a means of fixing minor glycolytic paralogs in the yeast genome. The MG strain was carefully designed and constructed to provide a robust prototrophic platform for quantitative studies and has been made available to the scientific community.

Chromosomal Copy Number Variation in Saccharomyces pastorianus Is Evidence for Extensive Genome Dynamics in Industrial Lager Brewing Strains

Lager brewing strains of Saccharomyces pastorianus are natural interspecific hybrids originating from the spontaneous hybridization of Saccharomyces cerevisiae and Saccharomyces eubayanus. Over the past 500 years, S. pastorianus has been domesticated to become one of the most important industrial microorganisms. Production of lager-type beers requires a set of essential phenotypes, including the ability to ferment maltose and maltotriose at low temperature, the production of flavors and aromas, and the ability to flocculate. Understanding of the molecular basis of complex brewing-related phenotypic traits is a prerequisite for rational strain improvement. While genome sequences have been reported, the variability and dynamics of S. pastorianus genomes have not been investigated in detail. Here, using deep sequencing and chromosome copy number analysis, we showed that S. pastorianus strain CBS1483 exhibited extensive aneuploidy. This was confirmed by quantitative PCR and by flow cytometry. As a direct consequence of this aneuploidy, a massive number of sequence variants was identified, leading to at least 1,800 additional protein variants in S. pastorianus CBS1483. Analysis of eight additional S. pastorianus strains revealed that the previously defined group I strains showed comparable karyotypes, while group II strains showed large interstrain karyotypic variability. Comparison of three strains with nearly identical genome sequences revealed substantial chromosome copy number variation, which may contribute to strain-specific phenotypic traits. The observed variability of lager yeast genomes demonstrates that systematic linking of genotype to phenotype requires a three-dimensional genome analysis encompassing physical chromosomal structures, the copy number of individual chromosomes or chromosomal regions, and the allelic variation of copies of individual genes.

CRISPR-Cas system enables fast and simple genome editing of industrial Saccharomyces cerevisiae strains

There is a demand to develop 3rd generation biorefineries that integrate energy production with the production of higher value chemicals from renewable feedstocks. Here, robust and stress-tolerant industrial strains of Saccharomyces cerevisiae will be suitable production organisms. However, their genetic manipulation is challenging, as they are usually diploid or polyploid. Therefore, there is a need to develop more efficient genetic engineering tools. We applied a CRISPR–Cas9 system for genome editing of different industrial strains, and show simultaneous disruption of two alleles of a gene in several unrelated strains with the efficiency ranging between 65% and 78%. We also achieved simultaneous disruption and knock-in of a reporter gene, and demonstrate the applicability of the method by designing lactic acid-producing strains in a single transformation event, where insertion of a heterologous gene and disruption of two endogenous genes occurred simultaneously. Our study provides a foundation for efficient engineering of industrial yeast cell factories.

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We developed CRISPR–Cas9-based system for gene disruptions in industrial yeast.

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We showed high rate of disruption efficiency in unrelated industrial strains.

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Gene knock-in may be performed simultaneously with gene disruption.

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Use of the described Cas9-based system results in marker-free stable genetic modifications.

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The method was applied for single-step construction of lactic acid-producing strains.

Evolutionary Engineering Improves Tolerance for Replacement Jet Fuels in Saccharomyces cerevisiae

Monoterpenes are liquid hydrocarbons with applications ranging from flavor and fragrance to replacement jet fuel. Their toxicity, however, presents a major challenge for microbial synthesis. Here we evolved limonene-tolerant Saccharomyces cerevisiae strains and sequenced six strains across the 200-generation evolutionary time course. Mutations were found in the tricalbin proteins Tcb2p and Tcb3p. Genomic reconstruction in the parent strain showed that truncation of a single protein (tTcb3p(1-989)), but not its complete deletion, was sufficient to recover the evolved phenotype improving limonene fitness 9-fold. tTcb3p(1-989) increased tolerance toward two other monoterpenes (β-pinene and myrcene) 11- and 8-fold, respectively, and tolerance toward the biojet fuel blend AMJ-700t (10% cymene, 50% limonene, 40% farnesene) 4-fold. tTcb3p(1-989) is the first example of successful engineering of phase tolerance and creates opportunities for production of the highly toxic C10 alkenes in yeast.