FEMS yeast research最新文献

Due to its wide availability, glycerol is considered as a promising alternative feedstock for microbial fermentation. As a model eukaryote, Saccharomyces cerevisiae is commonly adopted for bioproduction of various bulk and value-added chemicals, but it does not efficiently utilize glycerol. In this review, the metabolic pathway of glycerol and its regulation in S. cerevisiae are first introduced. Then, strategies, including metabolic engineering of the endogenous pathway, introduction of exogenous pathways, adaptive evolution, and reverse metabolic engineering, are summarized for improving the glycerol utilization in S. cerevisiae. Finally, methods for further improving glycerol utilization by S. cerevisiae are proposed. This review provides insights for designing engineered S. cerevisiae for efficient utilization of glycerol.

My career developed very differently from those of most academic researchers. After school, I worked for 6 years in industries that employed yeast to manufacture ethanol and beer. At university, I was trained as a microbiologist with very little training in molecular biology. I retrained in 1987 in molecular yeast genetics and focused on genetic engineering of industrial yeasts to minimize the production of spoilage compounds in wine and ethyl carbamate, a carcinogen, in wine. The malolactic yeast ML01 and the urea-degrading yeast were the first genetically enhanced yeasts that obtained US FDA approval for commercial applications. Apart from applied research, I was fascinated by classic molecular yeast genetic studies using sophisticated techniques such as transcriptomics, proteomics, and metabolomics. Doing research at the University of British Columbia was stimulating and exciting, we established a core microarray and metabolomics facility that was used by many scientists at UBC and hospitals in Vancouver. I also established a state-of-the-art Wine Library that was used to study aging of wines produced in British Columbia. Finally, I have been fortunate to know and collaborate with leading yeast scientists who motivated me.

CRISPR-Cas9 technology is widely used for precise and specific editing of Saccharomyces cerevisiae genome to obtain marker-free engineered hosts. Targeted double-strand breaks are controlled by a guide RNA (gRNA), a chimeric RNA containing a structural segment for Cas9 binding and a 20-mer guide sequence that hybridises to the genomic DNA target. Introducing the 20-mer guide sequence into gRNA expression vectors often requires complex, time-consuming, and/or expensive cloning procedures. We present a new plasmid for CRISPR-Cas9 genome editing in S. cerevisiae, pCEC-red. This tool allows to (i) transform yeast with both Cas9 and gRNA expression cassettes in a single plasmid and (ii) insert the 20-mer sequence in the plasmid with high efficiency, thanks to Golden Gate Assembly and (iii) a red chromoprotein-based screening to speed up the selection of correct plasmids. We tested genome-editing efficiency of pCEC-red by targeting the ADE2 gene. We chose three different 20-mer targets and designed two types of repair fragments to test pCEC-red for precision editing and for large DNA region replacement procedures. We obtained high efficiencies (∼90%) for both engineering procedures, suggesting that the pCEC system can be used for fast and reliable marker-free genome editing.

Lager yeasts are hybrids between Saccharomyces cerevisiae and S. eubayanus. Wine yeast biodiversity, however, has only recently been discovered to include besides pure S. cerevisiae strains also hybrids between different Saccharomyces yeasts as well as introgressions from non-Saccharomyces species. Here, we analysed the genome of the Champagne Epernay Geisenheim (CEG) wine yeast. This yeast is an allotetraploid (4n - 1) hybrid of S. cerevisiae harbouring a substantially reduced S. kudriavzevii genome contributing only 1/3 of a full genome equivalent. We identified a novel oligopeptide transporter gene, FOT4, in CEG located on chromosome XVI. FOT genes were originally derived from Torulaspora microellipsoides and FOT4 arose by non-allelic recombination between adjacent FOT1 and FOT2 genes. Fermentations of CEG in Riesling and Müller-Thurgau musts were compared with the S. cerevisiae Geisenheim wine yeast GHM, which does not carry FOT genes. At low temperature (10°C), CEG completed fermentations faster and produced increased levels of higher alcohols (e.g. isoamyl alcohol). At higher temperature (18°C), CEG produced higher amounts of the pineapple-like alkyl esters i-butyric and propionic acid ethyl esters compared to GHM. The hybrid nature of CEG thus provides advantages in grape must fermentations over S. cerevisiae wine yeasts, especially with regard to aroma production.

There is a logic to doing successful research, but graduate students and indeed postdoctoral fellows and young independent investigators often learn it apprentice style, by experience. The purpose of this essay is to provide the product of that experience and advice that I have found useful to young researchers as they begin their training and careers.

Genome-editing toolboxes are essential for the exploration and exploitation of nonconventional yeast species as cell factories, as they facilitate both genome studies and metabolic engineering. The nonconventional yeast Candida intermedia is a biotechnologically interesting species due to its capacity to convert a wide range of carbon sources, including xylose and lactose found in forestry and dairy industry waste and side-streams, into added-value products. However, possibilities of genetic manipulation have so far been limited due to lack of molecular tools for this species. We describe here the development of a genome editing method for C. intermedia, based on electroporation and gene deletion cassettes containing the Candida albicans NAT1 dominant selection marker flanked by 1000 base pair sequences homologous to the target loci. Linear deletion cassettes targeting the ADE2 gene originally resulted in <1% targeting efficiencies, suggesting that C. intermedia mainly uses nonhomologous end joining for integration of foreign DNA fragments. By developing a split-marker based deletion technique for C. intermedia, we successfully improved the homologous recombination rates, achieving targeting efficiencies up to 70%. For marker-less deletions, we also employed the split-marker cassette in combination with a recombinase system, which enabled the construction of double deletion mutants via marker recycling. Overall, the split-marker technique proved to be a quick and reliable method for generating gene deletions in C. intermedia, which opens the possibility to uncover and enhance its cell factory potential.

Hanseniaspora guilliermondii is a well-recognized producer of acetate esters associated with fruity and floral aromas. The molecular mechanisms underneath this production or the environmental factors modulating it remain unknown. Herein, we found that, unlike Saccharomyces cerevisiae, H. guilliermondii over-produces acetate esters and higher alcohols at low carbon-to-assimilable nitrogen (C:N) ratios, with the highest titers being obtained in the amino acid-enriched medium YPD. The evidences gathered support a model in which the strict preference of H. guilliermondii for amino acids as nitrogen sources results in a channeling of keto-acids obtained after transamination to higher alcohols and acetate esters. This higher production was accompanied by higher expression of the four HgAATs, genes, recently proposed to encode alcohol acetyl transferases. In silico analyses of these HgAat's reveal that they harbor conserved AATs motifs, albeit radical substitutions were identified that might result in different kinetic properties. Close homologues of HgAat2, HgAat3, and HgAat4 were only found in members of Hanseniaspora genus and phylogenetic reconstruction shows that these constitute a distinct family of Aat's. These results advance the exploration of H. guilliermondii as a bio-flavoring agent providing important insights to guide future strategies for strain engineering and media manipulation that can enhance production of aromatic volatiles.

For decades, the industrial vitamin B12 (cobalamin) production has been based on bacterial producer strains. Due to limited methods for strain optimization and difficult strain handling, the desire for new vitamin B12-producing hosts has risen. As a vitamin B12-independent organism with a big toolbox for genomic engineering and easy-to-handle cultivation conditions, Saccharomyces cerevisiae has high potential for heterologous vitamin B12 production. However, the B12 synthesis pathway is long and complex. To be able to easily engineer and evolve B12-producing recombinant yeast cells, we have developed an S. cerevisiae strain whose growth is dependent on vitamin B12. For this, the B12-independent methionine synthase Met6 of yeast was replaced by a B12-dependent methionine synthase MetH from Escherichia coli. Adaptive laboratory evolution, RT-qPCR, and overexpression experiments show that additional high-level expression of a bacterial flavodoxin/ferredoxin-NADP+ reductase (Fpr-FldA) system is essential for in vivo reactivation of MetH activity and growth. Growth of MetH-containing yeast cells on methionine-free media is only possible with the addition of adenosylcobalamin or methylcobalamin. A heterologous vitamin B12 transport system turned out to be not necessary for the uptake of cobalamins. This strain should be a powerful chassis to engineer B12-producing yeast cells.

ErCas12a is a class 2 type V CRISPR-Cas nuclease isolated from Eubacterium rectale with attractive fundamental characteristics, such as RNA self-processing capability, and lacks reach-through royalties typical for Cas nucleases. This study aims to develop a ErCas12a-mediated genome editing tool applicable in the model yeast Saccharomyces cerevisiae. The optimal design parameters for ErCas12a editing in S. cerevisiae were defined as a 21-nt spacer flanked by 19 nt direct repeats expressed from either RNApolII or III promoters, achieving near 100% editing efficiencies in commonly targeted genomic locations. To be able to transfer the ErCas12a genome editing tool to different strain lineages, a transportable platform plasmid was constructed and evaluated for its genome editing efficiency. Using an identical crRNA expression design, the transportable ErCas12a genome editing tool showed lower efficiency when targeting the ADE2 gene. In contrast to genomic Ercas12a expression, episomal expression of Ercas12a decreases maximum specific growth rate on glucose, indicating ErCas12a toxicity at high expression levels. Moreover, ErCas12a processed a multispacer crRNA array using the RNA self-processing capability, which allowed for simultaneous editing of multiple chromosomal locations. ErCas12a is established as a valuable addition to the genetic toolbox for S. cerevisiae.