Crabtree/Warburg-like aerobic xylose fermentation by engineered Saccharomyces cerevisiae

Bottlenecks in the efficient conversion of xylose into cost-effective biofuels have limited the widespread use of plant lignocellulose as a renewable feedstock. The yeast Saccharomyces cerevisiae ferments glucose into ethanol with such high metabolic flux that it ferments high concentrations of glucose aerobically, a trait called the Crabtree/Warburg Effect. In contrast to glucose, most engineered S. cerevisiae strains do not ferment xylose at economically viable rates and yields, and they require respiration to achieve sufficient xylose metabolic flux and energy return for growth aerobically. Here, we evolved respiration-deficient S. cerevisiae strains that can grow on and ferment xylose to ethanol aerobically, a trait analogous to the Crabtree/Warburg Effect for glucose. Through genome sequence comparisons and directed engineering, we determined that duplications of genes encoding engineered xylose metabolism enzymes, as well as TKL1, a gene encoding a transketolase in the pentose phosphate pathway, were the causative genetic changes for the evolved phenotype. Reengineered duplications of these enzymes, in combination with deletion mutations in HOG1, ISU1, GRE3, and IRA2, increased the rates of aerobic and anaerobic xylose fermentation. Importantly, we found that these genetic modifications function in another genetic background and increase the rate and yield of xylose-to-ethanol conversion in industrially relevant switchgrass hydrolysate, indicating that these specific genetic modifications may enable the sustainable production of industrial biofuels from yeast. We propose a model for how key regulatory mutations prime yeast for aerobic xylose fermentation by lowering the threshold for overflow metabolism, allowing mutations to increase xylose flux and to redirect it into fermentation products.     

Combinatorial metabolic pathway assembly in the yeast genome with RNA-guided Cas9

The yeast Saccharomyces cerevisiae is an important industrial platform for the production of grain and cellulosic ethanol, isobutanol, butanediol, isoprenoids, and other chemicals. The construction of a successful production strain usually involves multiple gene knockouts and chromosomal integration of expression cassettes to redirect the metabolic fluxes for the conversion of sugars and other feed stocks into the desired product. RNA-guided Cas9 based genome editing has been demonstrated in many prokaryotic and eukaryotic hosts including S. cerevisiae, in which it has been additionally exploited as a tool for metabolic engineering. To extend the utilization of RNA-guided Cas9 as a metabolic pathway building tool, we demonstrated the direct assembly and chromosomal integration of up to 17 overlapping DNA fragments encoding the beta-carotene biosynthetic pathway. Furthermore, we generated a combinatorial strain library for the beta-carotene biosynthetic pathway, directly integrated into the yeast genome to create a diverse library of strains. This enabled the screening of combinatorial libraries in stable chromosomally integrated strains for rapid improvements of product titers. This combinatorial approach for pathway assembly will significantly accelerate the current speed of metabolic engineering for S. cerevisiae as an industrial platform, and increase the number of strains that can be simultaneously evaluated for enzyme screening, expression optimization and protein engineering to achieve the titer, rate and yield necessary for the commercialization of new industrial fermentation products.     

Alcoholic fermentation of carbon sources in biomass hydrolysates by Saccharomyces cerevisiae: current status

Fuel ethanol production from plant biomass hydrolysates by Saccharomyces cerevisiae is of great economic and environmental significance. This paper reviews the current status with respect to alcoholic fermentation of the main plant biomass-derived monosaccharides by this yeast. Wild-type S. cerevisiae strains readily ferment glucose, mannose and fructose via the Embden–Meyerhof pathway of glycolysis, while galactose is fermented via the Leloir pathway. Construction of yeast strains that efficiently convert other potentially fermentable substrates in plant biomass hydrolysates into ethanol is a major challenge in metabolic engineering. The most abundant of these compounds is xylose. Recent metabolic and evolutionary engineering studies on S. cerevisiae strains that express a fungal xylose isomerase have enabled the rapid and efficient␣anaerobic fermentation of this pentose. l-Arabinose fermentation, based on the expression of a prokaryotic pathway in S. cerevisiae, has also been established, but needs further optimization before it can be considered for industrial implementation. In addition to these already investigated strategies, possible approaches for metabolic engineering of galacturonic acid and rhamnose fermentation by S. cerevisiae are discussed. An emerging and major challenge is to achieve the rapid transition from proof-of-principle experiments under ‘academic’ conditions (synthetic media, single substrates or simple substrate mixtures, absence of toxic inhibitors) towards efficient conversion of complex industrial substrate mixtures that contain synergistically acting inhibitors.     

Role of Hexose Transport in Control of Glycolytic Flux in Saccharomyces cerevisiae

The yeast Saccharomyces cerevisiae predominantly ferments glucose to ethanol at high external glucose concentrations, irrespective of the presence of oxygen. In contrast, at low external glucose concentrations and in the presence of oxygen, as in a glucose-limited chemostat, no ethanol is produced. The importance of the external glucose concentration suggests a central role for the affinity and maximal transport rates of yeast's glucose transporters in the control of ethanol production. Here we present a series of strains producing functional chimeras between the hexose transporters Hxt1 and Hxt7, each of which has distinct glucose transport characteristics. The strains display a range of decreasing glycolytic rates resulting in a proportional decrease in ethanol production. Using these strains, we show for the first time that at high glucose levels, the glucose uptake capacity of wild-type S. cerevisiae does not control glycolytic flux during exponential batch growth. In contrast, our chimeric Hxt transporters control the rate of glycolysis to a high degree. Strains whose glucose uptake is mediated by these chimeric transporters will undoubtedly provide a powerful tool with which to examine in detail the mechanism underlying the switch between fermentation and respiration in S. cerevisiae and will provide new tools for the control of industrial fermentations.     

Evaluation of industrial Saccharomyces cerevisiae strains as the chassis cell for second-generation bioethanol production

To develop a suitable Saccharomyces cerevisiae industrial strain as a chassis cell for ethanol production using lignocellulosic materials, 32 wild-type strains were evaluated for their glucose fermenting ability, their tolerance to the stresses they might encounter in lignocellulosic hydrolysate fermentation and their genetic background for pentose metabolism. The strain BSIF, isolated from tropical fruit in Thailand, was selected out of the distinctly different strains studied for its promising characteristics. The maximal specific growth rate of BSIF was as high as 0.65 h⁻¹ in yeast extract peptone dextrose medium, and the ethanol yield was 0.45 g g⁻¹ consumed glucose. Furthermore, compared with other strains, this strain exhibited superior tolerance to high temperature, hyperosmotic stress and oxidative stress; better growth performance in lignocellulosic hydrolysate; and better xylose utilization capacity when an initial xylose metabolic pathway was introduced. All of these results indicate that this strain is an excellent chassis strain for lignocellulosic ethanol production.     

Fermentation characteristics of Dekkera bruxellensis strains

The influence of pH, temperature and carbon source (glucose and maltose) on growth rate and ethanol yield of Dekkera bruxellensis was investigated using a full-factorial design. Growth rate and ethanol yield were lower on maltose than on glucose. In controlled oxygen-limited batch cultivations, the ethanol yield of the different combinations varied from 0.42 to 0.45 g (g glucose)⁻¹ and growth rates varied from 0.037 to 0.050 h⁻¹. The effect of temperature on growth rate and ethanol yield was negligible. It was not possible to model neither growth rate nor ethanol yield from the full-factorial design, as only marginal differences were observed in the conditions tested. When comparing three D. bruxellensis strains and two industrial isolates of Saccharomyces cerevisiae, S. cerevisiae grew five times faster, but the ethanol yields were 0–13% lower. The glycerol yields of S. cerevisiae strains were up to six-fold higher compared to D. bruxellensis, and the biomass yields reached only 72–84% of D. bruxellensis. Our results demonstrate that D. bruxellensis is robust to large changes in pH and temperature and may have a more energy-efficient metabolism under oxygen limitation than S. cerevisiae.     

Utilization of nitrate abolishes the “Custers effect” in Dekkera bruxellensis and determines a different pattern of fermentation products

Nitrate is one of the most abundant nitrogen sources in nature. Several yeast species have been shown to be able to assimilate nitrate and nitrite, but the metabolic pathway has been studied in very few of them. Dekkera bruxellensis can use nitrate as sole nitrogen source and this metabolic characteristic can render D. bruxellensis able to overcome S. cerevisiae populations in industrial bioethanol fermentations. In order to better characterize how nitrate utilization affects carbon metabolism and the yields of the fermentation products, we investigated this trait in defined media under well-controlled aerobic and anaerobic conditions. Our experiments showed that in D. bruxellensis, utilization of nitrate determines a different pattern of fermentation products. Acetic acid, instead of ethanol, became in fact the main product of glucose metabolism under aerobic conditions. We have also demonstrated that under anaerobic conditions, nitrate assimilation abolishes the “Custers effect”, in this way improving its fermentative metabolism. This can offer a new strategy, besides aeration, to sustain growth and ethanol production for the employment of this yeast in industrial processes.     

Fermentation performance and intracellular metabolite patterns in laboratory and industrial xylose-fermenting Saccharomyces cerevisiae

Heterologous genes for xylose utilization were introduced into an industrial Saccharomyces cerevisiae, strain A, with the aim of producing fuel ethanol from lignocellulosic feedstocks. Two transformants, A4 and A6, were evaluated by comparing the performance in 4-l anaerobic batch cultivations to both the parent strain and a laboratory xylose-utilizing strain: S. cerevisiae TMB 3001. During growth in a minimal medium containing a mixture of glucose and xylose (50 g/l each), glucose was preferentially consumed. During the first growth phase on glucose, the specific growth rates were 0.26, 0.32, 0.27 and 0.30 h^–1 for strains TMB 3001, A (parental strain), A4, and A6, respectively. The specific ethanol productivities were 0.04, 0.13, 0.04 and 0.03 g/g^.per hour, for TMB 3001, A, A4 and A6, respectively. The specific xylose consumption rates were 0.06, 0.21 and 0.14 g/g^.per hour, respectively for strains TMB 3001, A4 and A6. Xylose consumption resulted mainly in the formation of xylitol, with biomass and ethanol being minor products. The metabolite profile of intermediates in the pentose phosphate pathway and key glycolytic intermediates were determined during growth on glucose and xylose, respectively. The metabolite pattern differed depending on whether glucose or xylose was utilized. The levels of intracellular metabolites were higher in the industrial strains than in the laboratory strain during growth on xylose. Electronic Publication     

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.     

Metabolic engineering of Saccharomyces cerevisiae for the biotechnological production of succinic acid

The production of bio-based succinic acid is receiving great attention, and several predominantly prokaryotic organisms have been evaluated for this purpose. In this study we report on the suitability of the highly acid- and osmotolerant yeast Saccharomyces cerevisiae as a succinic acid production host. We implemented a metabolic engineering strategy for the oxidative production of succinic acid in yeast by deletion of the genes SDH1, SDH2, IDH1 and IDP1. The engineered strains harbor a TCA cycle that is completely interrupted after the intermediates isocitrate and succinate. The strains show no serious growth constraints on glucose. In glucose-grown shake flask cultures, the quadruple deletion strain Δsdh1Δsdh2Δidh1Δidp1 produces succinic acid at a titer of 3.62 g L^?1 (factor 4.8 compared to wild-type) at a yield of 0.11 mol (mol glucose)^?1. Succinic acid is not accumulated intracellularly. This makes the yeast S. cerevisiae a suitable and promising candidate for the biotechnological production of succinic acid on an industrial scale.     

Genotypic and phenotypic characterization of industrial autochthonous Saccharomyces cerevisiae for the selection of well-adapted bioethanol-producing strains

In northwestern Argentina, sugarcane-derived industrial fermentation is being extensively used for bioethanol production, where highly adaptive native strains compete with the baker's yeast Saccharomyces cerevisiae traditionally used as starter culture. Yeast populations of 10 distilleries from Tucumán (Argentina) were genotypic and phenotypic characterized to select well-adapted bioethanol-producing autochthonous strains to be used as starter cultures for the industrial production of bioethanol fuel. From the 192 isolates, 69.8% were identified as S. cerevisiae, 25.5% as non-Saccharomyces, and 4.7% as Saccharomyces sp. wild yeasts. The majority of S. cerevisiae isolates (68.5%) were non-flocculating yeasts, while the flocculating strains were all obtained from the only continuous fermentation process included in the study. Simple Sequence Repeat analysis revealed a high genetic diversity among S. cerevisiae genotypes, where all of them were very different from the original baker's strain used as starter. Among these, 38 strains multi-tolerant to stress by ethanol (8%), temperature (42.5 °C) and pH (2.0) were obtained. No major differences were found among these strains in terms of ethanol production and residual sugars in batch fermentation experiments with cell recycling. However, only 10 autochthonous strains maintained their viability (more than 80%) throughout five consecutive cycles of sugarcane-based fermentations. In summary, 10 autochthonous isolates were found to be superior to baker's yeast used as starter culture (S. cerevisiae Calsa) in terms of optimal technological, physiological and ecological properties. The knowledge generated on the indigenous yeast populations in industrial fermentation processes of bioethanol-producing distilleries allowed the selection of well-adapted bioethanol-producing strains.     

Mathematical model for aerobic culture of a recombinant yeast

A mathematical model was formulated to simulate cell growth, plasmid loss and recombinant protein production during the aerobic culture of a recombinant yeast S. cerevisiae. Model development was based on three simplified metabolic events in the yeast: glucose fermentation, glucose oxidation and ethanol oxidation. Cell growth was expressed as a composite of these metabolic events. Their contributions to the total specific growth rate depended on the activities of the pacemaker enzyme pools of the individual pathways. The pacemaker enzyme pools were regulated by the specific glucose uptake rate. The effect of substrate concentrations on the specific growth rate was described by a modified Monod equation. It was assumed that recombinant protein formation is only associated with oxidative pathways. Plasmid loss kinetics was formulated based on segregational instability during cell division by assuming constant probability of plasmid loss. Experiments on batch fermentation of recombinant S. cerevisiae C468/pGAC9 (ATCC 20690), which expresses Aspergillus awamori glucoamylase gene and secretes glucoamylase into the extracellular medium, were carried out in an airlift bioreactor in order to evaluate the proposed model. The model successfully predicted the dynamics of cell growth, glucose consumption, ethanol metabolism, glucoamylase production and plasmid instability. Excellent agreement between model simulations and our experimental data was achieved. Using published experimental data, model agreement was also found for other recombinant yeast strains. In general, the proposed model appears to be useful for the design, scale-up, control and optimization of recombinant yeast bioprocesses.     

Xylose assimilation enhances the production of isobutanol in engineered Saccharomyces cerevisiae

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

Alcoholic fermentation by wild-type <Emphasis Type="Italic">Hansenula polymorpha</Emphasis> and <Emphasis Type="Italic">Saccharomyces cerevisiae</Emphasis> versus recombinant strains with an elevated level of intracellular glutathione

The ability of baker’s yeast Saccharomyces cerevisiae and of the thermotolerant methylotrophic yeast Hansenula polymorpha to produce ethanol during alcoholic fermentation of glucose was compared between wild-type strains and recombinant strains possessing an elevated level of intracellular glutathione (GSH) due to overexpression of the first gene of GSH biosynthesis, gamma-glutamylcysteine synthetase, or of the central regulatory gene of sulfur metabolism, MET4. The analyzed strains of H. polymorpha with an elevated pool of intracellular GSH were found to accumulate almost twice as much ethanol as the wild-type strain during glucose fermentation, in contrast to GSH1-overexpressing S. cerevisiae strains, which also possessed an elevated pool of GSH. The ethanol tolerance of the GSH-overproducing strains was also determined. For this, the wild-type strain and transformants with an elevated GSH pool were compared for their viability upon exposure to exogenous ethanol. Unexpectedly, both S. cerevisiae and H. polymorpha transformants with a high GSH pool proved more sensitive to exogenous ethanol than the corresponding wild-type strains.     

Impact of overexpressing NADH kinase on glucose and xylose metabolism in recombinant xylose-utilizing <Emphasis Type="Italic">Saccharomyces cerevisiae</Emphasis>

During growth of Saccharomyces cerevisiae on glucose, the redox cofactors NADH and NADPH are predominantly involved in catabolism and biosynthesis, respectively. A deviation from the optimal level of these cofactors often results in major changes in the substrate uptake and biomass formation. However, the metabolism of xylose by recombinant S. cerevisiae carrying xylose reductase and xylitol dehydrogenase from the fungal pathway requires both NADH and NADPH and creates cofactor imbalance during growth on xylose. As one possible solution to overcoming this imbalance, the effect of overexpressing the native NADH kinase (encoded by the POS5 gene) in xylose-consuming recombinant S. cerevisiae directed either into the cytosol or to the mitochondria was evaluated. The physiology of the NADH kinase containing strains was also evaluated during growth on glucose. Overexpressing NADH kinase in the cytosol redirected carbon flow from CO₂ to ethanol during aerobic growth on glucose and to ethanol and acetate during anaerobic growth on glucose. However, cytosolic NADH kinase has an opposite effect during anaerobic metabolism of xylose consumption by channeling carbon flow from ethanol to xylitol. In contrast, overexpressing NADH kinase in the mitochondria did not affect the physiology to a large extent. Overall, although NADH kinase did not increase the rate of xylose consumption, we believe that it can provide an important source of NADPH in yeast, which can be useful for metabolic engineering strategies where the redox fluxes are manipulated.     

Comparison of the resistance of industrial and laboratory strains of Saccharomyces and Zygosaccharomyces to lignocellulose-derived fermentation inhibitors

Low-molecular weight aliphatic acids, furaldehydes and a broad range of different aromatic compounds are known to inhibit the fermentation of lignocellulose hydrolysates by yeasts. In this work, a cocktail of different lignocellulose-derived inhibitors was used to compare the inhibitor resistance of eleven different industrial and laboratory Saccharomyces cerevisiae strains and two Zygosaccharomyces strains. The inhibitor cocktail was composed of two aliphatic acids, formic and acetic acid, two furaldehydes, furfural and 5-hydroxymethylfurfural (HMF), and two aromatic compounds, cinnamic acid and coniferyl aldehyde. Fermentations were performed under oxygen-limited conditions and with different levels (100, 75, 50, 25 and 0%) of the inhibitor cocktail present. The ethanol yield on initial glucose, the volumetric and specific ethanol productivity, the biomass yield and the glucose consumption rates were used as criteria for the performance of the strains. The results revealed major differences in inhibitor resistance between yeast strains within the same species. The ethanol yield of the S. cerevisiae strain that was least affected decreased only with 10% at an inhibitor cocktail concentration of 100%, while the decrease in ethanol yield for the most sensitive S. cerevisiae strain was more than 50% already at an inhibitor cocktail concentration of 25%. Ethanol formation was generally less affected than growth and ethanol yield less than ethanol productivity. The two most resistant strains were an S. cerevisiae strain isolated from a spent sulphite liquor plant and one of the laboratory S. cerevisiae strains. Additional fermentations with either HMF or coniferyl aldehyde revealed that the degree of resistance of different yeast strains was highly dependent on the inhibitor used. A mutant strain of S. cerevisiae displaying enhanced resistance against coniferyl aldehyde compared with the parental strains was identified.     

Xylose isomerase overexpression along with engineering of the pentose phosphate pathway and evolutionary engineering enable rapid xylose utilization and ethanol production by Saccharomyces cerevisiae

Xylose is the main pentose and second most abundant sugar in lignocellulosic feedstocks. To improve xylose utilization, necessary for the cost-effective bioconversion of lignocellulose, several metabolic engineering approaches have been employed in the yeast Saccharomyces cerevisiae. In this study, we describe the rational metabolic engineering of a S. cerevisiae strain, including overexpression of the Piromyces xylose isomerase gene (XYLA), Pichia stipitis xylulose kinase (XYL3) and genes of the non-oxidative pentose phosphate pathway (PPP). This engineered strain (H131-A3) was used to initialize a three-stage process of evolutionary engineering, through first aerobic and anaerobic sequential batch cultivation followed by growth in a xylose-limited chemostat. The evolved strain H131-A3-AL^CS displayed significantly increased anaerobic growth rate (0.203±0.006 h^?1) and xylose consumption rate (1.866 g g^?1 h^?1) along with high ethanol conversion yield (0.41 g/g). These figures exceed by a significant margin any other performance metrics on xylose utilization and ethanol production by S. cerevisiae reported to-date. Further inverse metabolic engineering based on functional complementation suggested that efficient xylose assimilation is attributed, in part, to the elevated expression level of xylose isomerase, which was accomplished through the multiple-copy integration of XYLA in the chromosome of the evolved strain.