Co-cultivation of Saccharomyces cerevisiae strains combines advantages of different metabolic engineering strategies for improved ethanol yield.

Glycerol is the major organic byproduct of industrial ethanol production with the yeast Saccharomyces cerevisiae. Improved ethanol yields have been achieved with engineered S. cerevisiae strains in which heterologous pathways replace glycerol formation as the predominant mechanism for anaerobic re-oxidation of surplus NADH generated in biosynthetic reactions. Functional expression of heterologous phosphoribulokinase (PRK) and ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCO) genes enables yeast cells to couple a net oxidation of NADH to the conversion of glucose to ethanol. In another strategy, NADH-dependent reduction of exogenous acetate to ethanol is enabled by introduction of a heterologous acetylating acetaldehyde dehydrogenase (A-ALD). This study explores potential advantages of co-cultivating engineered PRK-RuBisCO-based and A-ALD-based strains in anaerobic bioreactor batch cultures. Co-cultivation of these strains, which in monocultures showed reduced glycerol yields and improved ethanol yields, strongly reduced the formation of acetaldehyde and acetate, two byproducts that were formed in anaerobic monocultures of a PRK-RuBisCO-based strain. In addition, co-cultures on medium with low acetate-to-glucose ratios that mimicked those in industrial feedstocks completely removed acetate from the medium. Kinetics of co-cultivation processes and glycerol production could be optimized by tuning the relative inoculum sizes of the two strains. Co-cultivation of a PRK-RuBisCO strain with a Δgpd1 Δgpd2 A-ALD strain, which was unable to grow in the absence of acetate and evolved for faster anaerobic growth in acetate-supplemented batch cultures, further reduced glycerol formation but led to extended fermentation times. These results demonstrate the potential of using defined consortia of engineered S. cerevisiae strains for high-yield, minimal-waste ethanol production.

Optimizing the balance between heterologous acetate- and CO2-reduction pathways in anaerobic cultures of Saccharomyces cerevisiae strains engineered for low-glycerol production.

In anaerobic Saccharomyces cerevisiae cultures, NADH (reduced form of nicotinamide adenine dinucleotide)-cofactor balancing by glycerol formation constrains ethanol yields. Introduction of an acetate-to-ethanol reduction pathway based on heterologous acetylating acetaldehyde dehydrogenase (A-ALD) can replace glycerol formation as 'redox-sink' and improve ethanol yields in acetate-containing media. Acetate concentrations in feedstock for first-generation bioethanol production are, however, insufficient to completely replace glycerol formation. An alternative glycerol-reduction strategy bypasses the oxidative reaction in glycolysis by introducing phosphoribulokinase (PRK) and ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCO). For optimal performance in industrial settings, yeast strains should ideally first fully convert acetate and, subsequently, continue low-glycerol fermentation via the PRK-RuBisCO pathway. However, anaerobic batch cultures of a strain carrying both pathways showed inferior acetate reduction relative to a strain expressing only the A-ALD pathway. Complete A-ALD-mediated acetate reduction by a dual-pathway strain, grown anaerobically on 50 g L-1 glucose and 5 mmol L-1 acetate, was achieved upon reducing PRK abundance by a C-terminal extension of its amino acid sequence. Yields of glycerol and ethanol on glucose were 55% lower and 6% higher, respectively, than those of a nonengineered reference strain. The negative impact of the PRK-RuBisCO pathway on acetate reduction was attributed to sensitivity of the reversible A-ALD reaction to intracellular acetaldehyde concentrations.

Respiratory reoxidation of NADH is a key contributor to high oxygen requirements of oxygen-limited cultures of Ogataea parapolymorpha.

While thermotolerance is an attractive trait for yeasts used in industrial ethanol production, oxygen requirements of known thermotolerant species are incompatible with process requirements. Analysis of oxygen-sufficient and oxygen-limited chemostat cultures of the facultatively fermentative, thermotolerant species Ogataea parapolymorpha showed its minimum oxygen requirements to be an order of magnitude larger than those reported for the thermotolerant yeast Kluyveromyces marxianus. High oxygen requirements of O. parapolymorpha coincided with a near absence of glycerol, a key NADH/NAD+ redox-cofactor-balancing product in many other yeasts, in oxygen-limited cultures. Genome analysis indicated absence of orthologs of the Saccharomyces cerevisiae glycerol-3-phosphate-phosphatase genes GPP1 and GPP2. Co-feeding of acetoin, whose conversion to 2,3-butanediol enables reoxidation of cytosolic NADH, supported a 2.5-fold increase of the biomass concentration in oxygen-limited cultures. An O. parapolymorpha strain in which key genes involved in mitochondrial reoxidation of NADH were inactivated did produce glycerol, but transcriptome analysis did not reveal a clear candidate for a responsible phosphatase. Expression of S. cerevisiae GPD2, which encodes NAD+-dependent glycerol-3-phosphate dehydrogenase, and GPP1 supported increased glycerol production by oxygen-limited chemostat cultures of O. parapolymorpha. These results identify dependence on respiration for NADH reoxidation as a key contributor to unexpectedly high oxygen requirements of O. parapolymorpha.

Towards closed carbon loop fermentations: Cofeeding of Yarrowia lipolytica with glucose and formic acid.

A novel fermentation process was developed in which renewable electricity is indirectly used as an energy source in fermentation, synergistically decreasing both the consumption of sugar as a first generation carbon source and emission of the greenhouse gas CO2 . As an illustration, a glucose-based process is co-fed with formic acid, which can be generated by capturing CO2 from fermentation offgas followed by electrochemical reduction with renewable electricity. This "closed carbon loop" concept is demonstrated by a case study in which cofeeding formic acid is shown to significantly increase the yield of biomass on glucose of the industrially relevant yeast species Yarrowia lipolytica. First, the optimal feed ratio of formic acid to glucose is established using chemostat cultivations. Subsequently, guided by a dynamic fermentation process model, a fed-batch protocol is developed and demonstrated on laboratory scale. Finally, the developed fed-batch process is tested and proven to be scalable at pilot scale. Extensions of the concept are discussed to apply the concept to anaerobic fermentations, and to recycle the O2 that is co-generated with the formic acid to aerobic fermentation processes for intensification purposes.

Carbon dioxide fixation via production of succinic acid from glycerol in engineered Saccharomyces cerevisiae.

BACKGROUND: The microbial production of succinic acid (SA) from renewable carbon sources via the reverse TCA (rTCA) pathway is a process potentially accompanied by net-fixation of carbon dioxide (CO2). Among reduced carbon sources, glycerol is particularly attractive since it allows a nearly twofold higher CO2-fixation yield compared to sugars. Recently, we described an engineered Saccharomyces cerevisiae strain which allowed SA production in synthetic glycerol medium with a maximum yield of 0.23 Cmol Cmol-1. The results of that previous study suggested that the glyoxylate cycle considerably contributed to SA accumulation in the respective strain. The current study aimed at improving the flux into the rTCA pathway accompanied by a higher CO2-fixation and SA yield. RESULTS: By changing the design of the expression cassettes for the rTCA pathway, overexpressing PYC2, and adding CaCO3 to the batch fermentations, an SA yield on glycerol of 0.63 Cmol Cmol-1 was achieved (i.e. 47.1% of the theoretical maximum). The modifications in this 2nd-generation SA producer improved the maximum biomass-specific glycerol consumption rate by a factor of nearly four compared to the isogenic baseline strain solely equipped with the dihydroxyacetone (DHA) pathway for glycerol catabolism. The data also suggest that the glyoxylate cycle did not contribute to the SA production in the new strain. Cultivation conditions which directly or indirectly increased the concentration of bicarbonate, led to an accumulation of malate in addition to the predominant product SA (ca. 0.1 Cmol Cmol-1 at the time point when SA yield was highest). Off-gas analysis in controlled bioreactors with CO2-enriched gas-phase indicated that CO2 was fixed during the SA production phase. CONCLUSIONS: The data strongly suggest that a major part of dicarboxylic acids in our 2nd-generation SA-producer was formed via the rTCA pathway enabling a net fixation of CO2. The greatly increased capacity of the rTCA pathway obviously allowed successful competition with other pathways for the common precursor pyruvate. The overexpression of PYC2 and the increased availability of bicarbonate, the co-substrate for the PYC reaction, further strengthened this capacity. The achievements are encouraging to invest in future efforts establishing a process for SA production from (crude) glycerol and CO2.

Rewriting yeast central carbon metabolism for industrial isoprenoid production.

A bio-based economy has the potential to provide sustainable substitutes for petroleum-based products and new chemical building blocks for advanced materials. We previously engineered Saccharomyces cerevisiae for industrial production of the isoprenoid artemisinic acid for use in antimalarial treatments. Adapting these strains for biosynthesis of other isoprenoids such as ß-farnesene (C15H24), a plant sesquiterpene with versatile industrial applications, is straightforward. However, S. cerevisiae uses a chemically inefficient pathway for isoprenoid biosynthesis, resulting in yield and productivity limitations incompatible with commodity-scale production. Here we use four non-native metabolic reactions to rewire central carbon metabolism in S. cerevisiae, enabling biosynthesis of cytosolic acetyl coenzyme A (acetyl-CoA, the two-carbon isoprenoid precursor) with a reduced ATP requirement, reduced loss of carbon to CO2-emitting reactions, and improved pathway redox balance. We show that strains with rewired central metabolism can devote an identical quantity of sugar to farnesene production as control strains, yet produce 25% more farnesene with that sugar while requiring 75% less oxygen. These changes lower feedstock costs and dramatically increase productivity in industrial fermentations which are by necessity oxygen-constrained. Despite altering key regulatory nodes, engineered strains grow robustly under taxing industrial conditions, maintaining stable yield for two weeks in broth that reaches >15% farnesene by volume. This illustrates that rewiring yeast central metabolism is a viable strategy for cost-effective, large-scale production of acetyl-CoA-derived molecules.

Energy coupling of membrane transport and efficiency of sucrose dissimilation in yeast.

Proton coupled transport of α-glucosides via Mal11 into Saccharomyces cerevisiae costs one ATP per imported molecule. Targeted mutation of all three acidic residues in the active site resulted in sugar uniport, but expression of these mutant transporters in yeast did not enable growth on sucrose. We then isolated six unique transporter variants of these mutants by directed evolution of yeast for growth on sucrose. In three variants, new acidic residues emerged near the active site that restored proton-coupled sucrose transport, whereas the other evolved transporters still catalysed sucrose uniport. The localization of mutations and transport properties of the mutants enabled us to propose a mechanistic model of proton-coupled sugar transport by Mal11. Cultivation of yeast strains expressing one of the sucrose uniporters in anaerobic, sucrose-limited chemostat cultures indicated an increase in the efficiency of sucrose dissimilation by 21% when additional changes in strain physiology were taken into account. We thus show that a combination of directed and evolutionary engineering results in more energy efficient sucrose transport, as a starting point to engineer yeast strains with increased yields for industrially relevant products.

Evolutionary engineering reveals amino acid substitutions in Ato2 and Ato3 that allow improved growth of Saccharomyces cerevisiae on lactic acid.

In Saccharomyces cerevisiae, the complete set of proteins involved in transport of lactic acid across the cell membrane has not been determined. In this study, we aimed to identify transport proteins not previously described to be involved in lactic acid transport via a combination of directed evolution, whole-genome resequencing and reverse engineering. Evolution of a strain lacking all known lactic acid transporters on lactate led to the discovery of mutated Ato2 and Ato3 as two novel lactic acid transport proteins. When compared to previously identified S. cerevisiae genes involved in lactic acid transport, expression of ATO3T284C was able to facilitate the highest growth rate (0.15 ± 0.01 h-1) on this carbon source. A comparison between (evolved) sequences and 3D models of the transport proteins showed that most of the identified mutations resulted in a widening of the narrowest hydrophobic constriction of the anion channel. We hypothesize that this observation, sometimes in combination with an increased binding affinity of lactic acid to the sites adjacent to this constriction, are responsible for the improved lactic acid transport in the evolved proteins.

Weak Acid Permeation in Synthetic Lipid Vesicles and Across the Yeast Plasma Membrane.

We present a fluorescence-based approach for determination of the permeability of small molecules across the membranes of lipid vesicles and living cells. With properly designed experiments, the method allows us to assess the membrane physical properties both in vitro and in vivo. We find that the permeability of weak acids increases in the order of benzoic > acetic > formic > lactic, both in synthetic lipid vesicles and the plasma membrane of Saccharomyces cerevisiae, but the permeability is much lower in yeast (one to two orders of magnitude). We observe a relation between the molecule permeability and the saturation of the lipid acyl chain (i.e., lipid packing) in the synthetic lipid vesicles. By analyzing wild-type yeast and a manifold knockout strain lacking all putative lactic acid transporters, we conclude that the yeast plasma membrane is impermeable to lactic acid on timescales up to ∼2.5 h.

Regulatory control circuits for stabilizing long-term anabolic product formation in yeast.

Engineering living cells for production of chemicals, enzymes and therapeutics can burden cells due to use of limited native co-factor availability and/or expression burdens, totalling a fitness deficit compared to parental cells encoded through long evolutionary trajectories to maximise fitness. Ultimately, this discrepancy puts a selective pressure against fitness-burdened engineered cells under prolonged bioprocesses, and potentially leads to complete eradication of high-performing engineered cells at the population level. Here we present the mutation landscapes of fitness-burdened yeast cells engineered for vanillin-ß-glucoside production. Next, we design synthetic control circuits based on transcriptome analysis and biosensors responsive to vanillin-ß-glucoside pathway intermediates in order to stabilize vanillin-ß-glucoside production over ~55 generations in sequential passage experiments. Furthermore, using biosensors with two different modes of action we identify control circuits linking vanillin-ß-glucoside pathway flux to various essential cellular functions, and demonstrate control circuits robustness and almost 2-fold higher vanillin-ß-glucoside production, including 5-fold increase in total vanillin-ß-glucoside pathway metabolite accumulation, in a fed-batch fermentation compared to vanillin-ß-glucoside producing cells without control circuits.

Contribution of Complex I NADH Dehydrogenase to Respiratory Energy Coupling in Glucose-Grown Cultures of Ogataea parapolymorpha.

The thermotolerant yeast Ogataea parapolymorpha (formerly Hansenula polymorpha) is an industrially relevant production host that exhibits a fully respiratory sugar metabolism in aerobic batch cultures. NADH-derived electrons can enter its mitochondrial respiratory chain either via a proton-translocating complex I NADH-dehydrogenase or via three putative alternative NADH dehydrogenases. This respiratory entry point affects the amount of ATP produced per NADH/O2 consumed and therefore impacts the maximum yield of biomass and/or cellular products from a given amount of substrate. To investigate the physiological importance of complex I, a wild-type O. parapolymorpha strain and a congenic complex I-deficient mutant were grown on glucose in aerobic batch, chemostat, and retentostat cultures in bioreactors. In batch cultures, the two strains exhibited a fully respiratory metabolism and showed the same growth rates and biomass yields, indicating that, under these conditions, the contribution of NADH oxidation via complex I was negligible. Both strains also exhibited a respiratory metabolism in glucose-limited chemostat cultures, but the complex I-deficient mutant showed considerably reduced biomass yields on substrate and oxygen, consistent with a lower efficiency of respiratory energy coupling. In glucose-limited retentostat cultures at specific growth rates down to ∼0.001 h-1, both O. parapolymorpha strains showed high viability. Maintenance energy requirements at these extremely low growth rates were approximately 3-fold lower than estimated from faster-growing chemostat cultures, indicating a stringent-response-like behavior. Quantitative transcriptome and proteome analyses indicated condition-dependent expression patterns of complex I subunits and of alternative NADH dehydrogenases that were consistent with physiological observations.IMPORTANCE Since popular microbial cell factories have typically not been selected for efficient respiratory energy coupling, their ATP yields from sugar catabolism are often suboptimal. In aerobic industrial processes, suboptimal energy coupling results in reduced product yields on sugar, increased process costs for oxygen transfer, and volumetric productivity limitations due to limitations in gas transfer and cooling. This study provides insights into the contribution of mechanisms of respiratory energy coupling in the yeast cell factory Ogataea parapolymorpha under different growth conditions and provides a basis for rational improvement of energy coupling in yeast cell factories. Analysis of energy metabolism of O. parapolymorpha at extremely low specific growth rates indicated that this yeast reduces its energy requirements for cellular maintenance under extreme energy limitation. Exploration of the mechanisms for this increased energetic efficiency may contribute to an optimization of the performance of industrial processes with slow-growing eukaryotic cell factories.

Functional expression of a bacterial α-ketoglutarate dehydrogenase in the cytosol of Saccharomyces cerevisiae.

Efficient production of fuels and chemicals by metabolically engineered micro-organisms requires availability of precursor molecules for product pathways. In eukaryotic cell factories, heterologous product pathways are usually expressed in the cytosol, which may limit availability of precursors that are generated in other cellular compartments. In Saccharomyces cerevisiae, synthesis of the precursor molecule succinyl-Coenzyme A is confined to the mitochondrial matrix. To enable cytosolic synthesis of succinyl-CoA, we expressed the structural genes for all three subunits of the Escherichia coli α-ketoglutarate dehydrogenase (αKGDH) complex in S. cerevisiae. The E. coli lipoic-acid scavenging enzyme was co-expressed to enable cytosolic lipoylation of the αKGDH complex, which is required for its enzymatic activity. Size-exclusion chromatography and mass spectrometry indicated that the heterologously expressed αKGDH complex contained all subunits and that its size was the same as in E. coli. Functional expression of the heterologous complex was evident from increased αKGDH activity in the cytosolic fraction of yeast cell homogenates. In vivo cytosolic activity of the αKGDH complex was tested by constructing a reporter strain in which the essential metabolite 5-aminolevulinic acid could only be synthetized from cytosolic, and not mitochondrial, succinyl-CoA. To this end HEM1, which encodes the succinyl-CoA-converting mitochondrial enzyme 5-aminolevulinic acid (ALA) synthase, was deleted and a bacterial ALA synthase was expressed in the cytosol. In the resulting strain, complementation of ALA auxotrophy depended on activation of the αKGDH complex by lipoic acid addition. Functional expression of a bacterial αKGDH complex in yeast represents a vital step towards efficient yeast-based production of compounds such as 1,4-butanediol and 4-aminobutyrate, whose product pathways use succinyl-CoA as a precursor.

A toolkit for rapid CRISPR-SpCas9 assisted construction of hexose-transport-deficient Saccharomyces cerevisiae strains.

Hexose transporter-deficient yeast strains are valuable testbeds for the study of sugar transport by native and heterologous transporters. In the popular Saccharomyces cerevisiae strain EBY.VW4000, deletion of 21 transporters completely abolished hexose transport. However, repeated use of the LoxP/Cre system in successive deletion rounds also resulted in major chromosomal rearrangements, gene loss and phenotypic changes. In the present study, CRISPR/SpCas9 was used to delete the 21 hexose transporters in an S. cerevisiae strain from the CEN.PK family in only three deletion rounds, using 11 unique guide RNAs. Even upon prolonged cultivation, the resulting strain IMX1812 (CRISPR-Hxt0) was unable to consume glucose, while its growth rate on maltose was the same as that of a strain equipped with a full set of hexose transporters. Karyotyping and whole-genome sequencing of the CRISPR-Hxt0 strain with Illumina and Oxford Nanopore technologies did not reveal chromosomal rearrangements or other unintended mutations besides a few SNPs. This study provides a new, 'genetically unaltered' hexose transporter-deficient strain and supplies a CRISPR toolkit for removing all hexose transporter genes from most S. cerevisiae laboratory strains in only three transformation rounds.

Laboratory evolution and physiological analysis of Saccharomyces cerevisiae strains dependent on sucrose uptake via the Phaseolus vulgaris Suf1 transporter.

Knowledge on the genetic factors important for the efficient expression of plant transporters in yeast is still very limited. Phaseolus vulgaris sucrose facilitator 1 (PvSuf1), a presumable uniporter, was an essential component in a previously published strategy aimed at increasing ATP yield in Saccharomyces cerevisiae. However, attempts to construct yeast strains in which sucrose metabolism was dependent on PvSUF1 led to slow sucrose uptake. Here, PvSUF1-dependent S. cerevisiae strains were evolved for faster growth. Of five independently evolved strains, two showed an approximately twofold higher anaerobic growth rate on sucrose than the parental strain (µ = 0.19 h-1 and µ = 0.08 h-1 , respectively). All five mutants displayed sucrose-induced proton uptake (13-50 µmol H+ (g biomass)-1  min-1 ). Their ATP yield from sucrose dissimilation, as estimated from biomass yields in anaerobic chemostat cultures, was the same as that of a congenic strain expressing the native sucrose symporter Mal11p. Four out of six observed amino acid substitutions encoded by evolved PvSUF1 alleles removed or introduced a cysteine residue and may be involved in transporter folding and/or oligomerization. Expression of one of the evolved PvSUF1 alleles (PvSUF1I209F C265F G326C ) in an unevolved strain enabled it to grow on sucrose at the same rate (0.19 h-1 ) as the corresponding evolved strain. This study shows how laboratory evolution may improve sucrose uptake in yeast via heterologous plant transporters, highlights the importance of cysteine residues for their efficient expression, and warrants reinvestigation of PvSuf1's transport mechanism.

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.

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.

Genome editing in Kluyveromyces and Ogataea yeasts using a broad-host-range Cas9/gRNA co-expression plasmid.

While CRISPR-Cas9-mediated genome editing has transformed yeast research, current plasmids and cassettes for Cas9 and guide-RNA expression are species specific. CRISPR tools that function in multiple yeast species could contribute to the intensifying research on non-conventional yeasts. A plasmid carrying a pangenomic origin of replication and two constitutive expression cassettes for Cas9 and ribozyme-flanked gRNAs was constructed. Its functionality was tested by analyzing inactivation of the ADE2 gene in four yeast species. In two Kluyveromyces species, near-perfect targeting (≥96%) and homologous repair (HR) were observed in at least 24% of transformants. In two Ogataea species, Ade- mutants were not observed directly after transformation, but prolonged incubation of transformed cells resulted in targeting efficiencies of 9% to 63% mediated by non-homologous end joining (NHEJ). In an Ogataea parapolymorpha ku80 mutant, deletion of OpADE2 mediated by HR was achieved, albeit at low efficiencies (<1%). Furthermore the expression of a dual polycistronic gRNA array enabled simultaneous interruption of OpADE2 and OpYNR1 demonstrating flexibility of ribozyme-flanked gRNA design for multiplexing. While prevalence of NHEJ prevented HR-mediated editing in Ogataea, such targeted editing was possible in Kluyveromyces. This broad-host-range CRISPR/gRNA system may contribute to exploration of Cas9-mediated genome editing in other Saccharomycotina yeasts.

A CRISPR/Cas9-based exploration into the elusive mechanism for lactate export in Saccharomyces cerevisiae.

CRISPR/Cas9-based genome editing allows rapid, simultaneous modification of multiple genetic loci in Saccharomyces cerevisiae. Here, this technique was used in a functional analysis study aimed at identifying the hitherto unknown mechanism of lactate export in this yeast. First, an S. cerevisiae strain was constructed with deletions in 25 genes encoding transport proteins, including the complete aqua(glycero)porin family and all known carboxylic acid transporters. The 25-deletion strain was then transformed with an expression cassette for Lactobacillus casei lactate dehydrogenase (LcLDH). In anaerobic, glucose-grown batch cultures this strain exhibited a lower specific growth rate (0.15 vs. 0.25 h-1) and biomass-specific lactate production rate (0.7 vs. 2.4 mmol g biomass-1 h-1) than an LcLDH-expressing reference strain. However, a comparison of the two strains in anaerobic glucose-limited chemostat cultures (dilution rate 0.10 h-1) showed identical lactate production rates. These results indicate that, although deletion of the 25 transporter genes affected the maximum specific growth rate, it did not impact lactate export rates when analysed at a fixed specific growth rate. The 25-deletion strain provides a first step towards a 'minimal transportome' yeast platform, which can be applied for functional analysis of specific (heterologous) transport proteins as well as for evaluation of metabolic engineering strategies.

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.

CRISPR/Cas9: a molecular Swiss army knife for simultaneous introduction of multiple genetic modifications in Saccharomyces cerevisiae.

A variety of techniques for strain engineering in Saccharomyces cerevisiae have recently been developed. However, especially when multiple genetic manipulations are required, strain construction is still a time-consuming process. This study describes new CRISPR/Cas9-based approaches for easy, fast strain construction in yeast and explores their potential for simultaneous introduction of multiple genetic modifications. An open-source tool (http://yeastriction.tnw.tudelft.nl) is presented for identification of suitable Cas9 target sites in S. cerevisiae strains. A transformation strategy, using in vivo assembly of a guideRNA plasmid and subsequent genetic modification, was successfully implemented with high accuracies. An alternative strategy, using in vitro assembled plasmids containing two gRNAs, was used to simultaneously introduce up to six genetic modifications in a single transformation step with high efficiencies. Where previous studies mainly focused on the use of CRISPR/Cas9 for gene inactivation, we demonstrate the versatility of CRISPR/Cas9-based engineering of yeast by achieving simultaneous integration of a multigene construct combined with gene deletion and the simultaneous introduction of two single-nucleotide mutations at different loci. Sets of standardized plasmids, as well as the web-based Yeastriction target-sequence identifier and primer-design tool, are made available to the yeast research community to facilitate fast, standardized and efficient application of the CRISPR/Cas9 system.