Reductive amination is one of the most synthetically direct routes to access chiral amines. Several Imine Reductases (IREDs) have been discovered to catalyze reductive amination (Reductive Aminases or RedAms), yet they are dependent on the expensive phosphorylated nicotinamide adenine dinucleotide cofactor NADPH and usually more active at basic pH. Here, we describe the discovery and synthetic potential of an IRED from Rhodococcus erythropolis (RytRedAm) that catalyzes reductive amination between a series of medium to large carbonyl and amine compounds with conversions of up to >99% and 99% enantiomeric excess at neutral pH. RytRedAm catalyzes the formation of a substituted γ-lactam and N-methyl-1-phenylethanamine with stereochemistry opposite to that of fungal RedAms, giving the (S)-enantiomer. This enzyme remarkably uses both NADPH and NADH cofactors with K_M values of 15 and 247 μM and turnover numbers k_cat of 3.6 and 9.0 s^-1, respectively, for the reductive amination of hexanal with allylamine. The crystal structure obtained provides insights into the flexibility to also accept NADH, with residues R35 and I69 diverging from that of other IREDs/RedAms in the otherwise conserved Rossmann fold. RytRedAm thus represents a subfamily of enzymes that enable synthetic applications using NADH-dependent reductive amination to access complementary chiral amine products.

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Chemically defined media for cultivation of Saccharomyces cerevisiae strains are commonly supplemented with a mixture of multiple Class-B vitamins, whose omission leads to strongly reduced growth rates. Fast growth without vitamin supplementation is interesting for industrial applications, as it reduces costs and complexity of medium preparation and may decrease susceptibility to contamination by auxotrophic microbes. In this study, suboptimal growth rates of S. cerevisiae CEN.PK113-7D in the absence of pantothenic acid, para-aminobenzoic acid (pABA), pyridoxine, inositol and/or biotin were corrected by single or combined overexpression of ScFMS1, ScABZ1/ScABZ2, ScSNZ1/ScSNO1, ScINO1 and Cyberlindnera fabianii BIO1, respectively. Several strategies were explored to improve growth of S. cerevisiae CEN.PK113-7D in thiamine-free medium. Overexpression of ScTHI4 and/or ScTHI5 enabled thiamine-independent growth at 83% of the maximum specific growth rate of the reference strain in vitamin-supplemented medium. Combined overexpression of seven native S. cerevisiae genes and CfBIO1 enabled a maximum specific growth rate of 0.33 ± 0.01 h⁻¹ in vitamin-free synthetic medium. This growth rate was only 17 % lower than that of a congenic reference strain in vitamin-supplemented medium. Physiological parameters of the engineered vitamin-independent strain in aerobic glucose-limited chemostat cultures (dilution rate 0.10 h⁻¹) grown on vitamin-free synthetic medium were similar to those of similar cultures of the parental strain grown on vitamin-supplemented medium. Transcriptome analysis revealed only few differences in gene expression between these cultures, which primarily involved genes with roles in Class-B vitamin metabolism. These results pave the way for development of fast-growing vitamin-independent industrial strains of S. cerevisiae.

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Emerging low-emission production technologies make ethanol an interesting substrate for yeast biotechnology, but information on growth rates and biomass yields of yeasts on ethanol is scarce. Strains of 52 Saccharomycotina yeasts were screened for growth on ethanol. The 21 fastest strains, among which representatives of the Phaffomycetales order were overrepresented, showed specific growth rates in ethanol-grown shake-flask cultures between 0.12 and 0.46 h⁻¹. Seven strains were studied in aerobic, ethanol-limited chemostats (dilution rate 0.10 h⁻¹). Saccharomyces cerevisiae and Kluyveromyces lactis, whose genomes do not encode Complex-I-type NADH dehydrogenases, showed biomass yields of 0.59 and 0.56 gbiomass g_ethanol⁻¹, respectively. Different biomass yields were observed among species whose genomes do harbour Complex-I-encoding genes: Phaffomyces thermotolerans (0.58 g g⁻¹), Pichia ethanolica (0.59 g g⁻¹), Saturnispora dispora (0.66 g g⁻¹), Ogataea parapolymorpha (0.67 g g⁻¹), and Cyberlindnera jadinii (0.73 g g⁻¹). Cyberlindnera jadinii biomass showed the highest protein content (59 ± 2%) of these yeasts. Its biomass yield corresponded to 88% of the theoretical maximum that is reached when growth is limited by assimilation rather than by energy availability. This study suggests that energy coupling of mitochondrial respiration and its regulation will become key factors for selecting and improving yeast strains for ethanol-based processes.

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Despite being present in trace amounts, ethyl esters play a crucial role as flavour compounds in lager beer. In yeast, ethyl hexanoate, ethyl octanoate and ethyl decanoate, responsible for fruity and floral taste tones, are synthesized from the toxic medium chain acyl-CoA intermediates released by the fatty acid synthase complex during the fatty acid biosynthesis, as a protective mechanism. The aim of this study was to enhance the production of ethyl esters in the hybrid lager brewing yeast Saccharomyces pastorianus by improving the medium chain acyl-CoA precursor supply. Through CRISPR-Cas9-based genetic engineering, specific FAS1 and FAS2 genes harbouring mutations in domains of the fatty acid synthesis complex were overexpressed in a single and combinatorial approach. These mutations in the ScFAS genes led to specific overproduction of the respective ethyl esters: overexpression of ScFAS1^I306A and ScFAS2^G1250S significantly improved ethyl hexanoate production and ScFAS1^R1834K boosted the ethyl octanoate production. Combinations of ScFAS1 mutant genes with ScFAS2^G1250S greatly enhanced predictably the final ethyl ester concentrations in cultures grown on full malt wort, but also resulted in increased levels of free medium chain fatty acids causing alterations in flavour profiles. Finally, the elevated medium chain fatty acid pool was directed towards the ethyl esters by overexpressing the esterase ScEEB1. The genetically modified S. pastorianus strains were utilized in lager beer production, and the resulting beverage exhibited significantly altered flavour profiles, thereby greatly expanding the possibilities of the flavour palette of lager beers.

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Current microbial cell factory methods for producing chemicals from renewable resources primarily rely on (fed-)batch production systems, leading to the accumulation of the desired product. Industrially relevant chemicals like 2-phenylethanol (2PE), a flavor and fragrance compound, can exhibit toxicity at low concentrations, inhibit the host activity, and negatively impact titer, rate, and yield. Batch liquid-liquid (L-L) In Situ Product Removal (ISPR) was employed to mitigate inhibition effects, but was not found sufficient for industrial-scale application. Here, we demonstrated that continuous selective L-L ISPR provides the solution for maintaining the productivity of de novo produced 2PE at an industrial pilot scale. A unique bioreactor concept called “Fermentation Accelerated by Separation Technology” (FAST) utilizes hydrostatic pressure differences to separate aqueous- and extractant streams within one unit operation, where both production and product extraction take place - allowing for the control of the concentration of the inhibiting compound. Controlled aqueous 2PE levels (0.43 ± 0.02 g kg−1) and extended production times (>100 h) were obtained and co-inhibiting by-product formation was reduced, resulting in a twofold increase of the final product output of batch L-L ISPR approaches. This study establishes that continuous selective L-L ISPR, enabled by FAST, can be applied for more economically viable production of inhibiting products.@en

Background
Elimination of greenhouse gas emissions in industrial biotechnology requires replacement of carbohydrates by alternative carbon substrates, produced from CO2 and waste streams. Ethanol is already industrially produced from agricultural residues and waste gas and is miscible with water, self-sterilizing and energy-dense. The yeast C. jadinii can grow on ethanol and has a history in the production of single-cell protein (SCP) for feed and food applications. To address a knowledge gap in quantitative physiology of C. jadinii during growth on ethanol, this study investigates growth kinetics, growth energetics, nutritional requirements, and biomass composition of C. jadinii strains in batch, chemostat and fed-batch cultures.

Results
In aerobic, ethanol-limited chemostat cultures, C. jadinii CBS 621 exhibited a maximum biomass yield on ethanol (Y max X/S) of 0.83 gbiomass (gethanol)−1 and an estimated maintenance requirement for ATP (mATP) of 2.7 mmolATP (gbiomass)−1 h−1. Even at specific growth rates below 0.05 h−1, a stable protein content of approximately 0.54 gprotein (gbiomass)−1 was observed. At low specific growth rates, up to 17% of the proteome consisted of alcohol dehydrogenase proteins, followed by aldehyde dehydrogenases and acetyl-CoA synthetase. Of 13 C. jadinii strains evaluated, 11 displayed fast growth on ethanol (μmax > 0.4 h−1) in mineral medium without vitamins, and CBS 621 was found to be a thiamine auxotroph. The prototrophic strain C. jadinii CBS 5947 was grown on an inorganic salts medium in fed-batch cultures (10-L scale) fed with pure ethanol. Biomass concentrations in these cultures increased up to 100 gbiomass (kgbroth)−1, with a biomass yield of 0.65 gbiomass (gethanol)−1. Model-based simulation, based on quantitative parameters determined in chemostat cultures, adequately predicted biomass production. A different protein content of chemostat- and fed-batch-grown biomass (54 and 42%, respectively) may reflect the more dynamic conditions in fed-batch cultures.

Conclusions
Analysis of ethanol-grown batch, chemostat and fed-batch cultures provided a quantitative physiology baseline for fundamental and applied research on C. jadinii. Its high maximum growth rate, high energetic efficiency of ethanol dissimilation, simple nutritional requirements and high protein content, make C. jadinii a highly interesting platform for production of SCP and other products from ethanol.@en

The biobased-economy aims to create a circular biotechnology ecosystem to transition from a fossil fuel-based to a sustainable industry based on biomass. For this, new microbial cell-factories are essential. We present the draft genome of the CEN.PK-derived Saccharomyces cerevisiae SpyCas9 expressing strain (IMX2600), that serve as chassis of new cell-factories.

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Carbon-based products are essential to society, yet producing them from fossil fuels is unsustainable. Microorganisms have the ability to take up electrons from solid electrodes and convert carbon dioxide (CO₂) to valuable carbon-based chemicals. However, higher productivities and energy efficiencies are needed to reach a viability that can make the technology transformative. Here, we show how a biofilm-based microbial porous cathode in a directed flow-through electrochemical system can continuously reduce CO₂ to even-chain C2–C6 carboxylic acids over 248 days. We demonstrate a threefold higher biofilm concentration, volumetric current density, and productivity compared with the state of the art. Most notably, the volumetric productivity (VP) resembles those achieved in laboratory-scale and industrial syngas (CO-H₂-CO₂) fermentation and chain elongation fermentation. This work highlights key design parameters for efficient electricity-driven microbial CO₂ reduction. There is need and room to improve the rates of electrode colonization and microbe-specific kinetics to scale up the technology.

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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.

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Biological functions are orchestrated by intricate networks of interacting genetic elements. Predicting the interaction landscape remains a challenge for systems biology and new research tools allowing simple and rapid mapping of sequence to function are desirable. Here, we describe CRI-SPA, a method allowing the transfer of chromosomal genetic features from a CRI-SPA Donor strain to arrayed strains in large libraries of Saccharomyces cerevisiae. CRI-SPA is based on mating, CRISPR-Cas9-induced gene conversion, and Selective Ploidy Ablation. CRI-SPA can be massively parallelized with automation and can be executed within a week. We demonstrate the power of CRI-SPA by transferring four genes that enable betaxanthin production into each strain of the yeast knockout collection (≈4800 strains). Using this setup, we show that CRI-SPA is highly efficient and reproducible, and even allows marker-free transfer of genetic features. Moreover, we validate a set of CRI-SPA hits by showing that their phenotypes correlate strongly with the phenotypes of the corresponding mutant strains recreated by reverse genetic engineering. Hence, our results provide a genome-wide overview of the genetic requirements for betaxanthin production. We envision that the simplicity, speed, and reliability offered by CRI-SPA will make it a versatile tool to forward systems-level understanding of biological processes.

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Mitochondria fulfil many essential roles and have their own genome, which is expressed as polycistronic transcripts that undergo co- or posttranscriptional processing and splicing. Due to the inherent complexity and limited technical accessibility of the mitochondrial transcriptome, fundamental questions regarding mitochondrial gene expression and splicing remain unresolved, even in the model eukaryote Saccharomyces cerevisiae. Long-read sequencing could address these fundamental questions. Therefore, a method for the enrichment of mitochondrial RNA and sequencing using Nanopore technology was developed, enabling the resolution of splicing of polycistronic genes and the quantification of spliced RNA. This method successfully captured the full mitochondrial transcriptome and resolved RNA splicing patterns with single-base resolution and was applied to explore the transcriptome of S. cerevisiae grown with glucose or ethanol as the sole carbon source, revealing the impact of growth conditions on mitochondrial RNA expression and splicing. This study uncovered a remarkable difference in the turnover of Group II introns between yeast grown in either mostly fermentative or fully respiratory conditions. Whether this accumulation of introns in glucose medium has an impact on mitochondrial functions remains to be explored. Combined with the high tractability of the model yeast S. cerevisiae, the developed method enables to monitor mitochondrial transcriptome responses in a broad range of relevant contexts, including oxidative stress, apoptosis and mitochondrial diseases.

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Saccharomyces pastorianus is not a classical taxon, it is an interspecific hybrid resulting from the cross of Saccharomyces cerevisiae and Saccharomyces eubayanus. Exhibiting heterosis for phenotypic traits such as wort α-oligosaccharide consumption and fermentation at low temperature, it has been domesticated to become the main workhorse of the brewing industry. Although CRISPR-Cas9 has been shown to be functional in S. pastorianus, repair of CRISPR-induced double strand breaks is unpredictable and preferentially uses the homoeologous chromosome as template, preventing targeted introduction of the desired repair construct. Here, we demonstrate that lager hybrids can be edited with near 100% efficiency at carefully selected landing sites on the chimeric SeScCHRIII. The landing sites were systematically selected and evaluated for (i) absence of loss of heterozygosity upon CRISPR-editing, (ii) efficiency of the gRNA, and (iii) absence of effect on strain physiology. Successful examples of highly efficient single and double gene integration illustrated that genome editing can be applied in interspecies hybrids, paving the way to a new impulse to lager yeast strain development.

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The construction of powerful cell factories requires intensive genetic engineering for the addition of new functionalities and the remodeling of native pathways and processes. The present study demonstrates the feasibility of extensive genome reprogramming using modular, specialized de novo-assembled neochromosomes in yeast. The in vivo assembly of linear and circular neochromosomes, carrying 20 native and 21 heterologous genes, enabled the first de novo production in a microbial cell factory of anthocyanins, plant compounds with a broad range of pharmacological properties. Turned into exclusive expression platforms for heterologous and essential metabolic routes, the neochromosomes mimic native chromosomes regarding mitotic and genetic stability, copy number, harmlessness for the host and editability by CRISPR/Cas9. This study paves the way for future microbial cell factories with modular genomes in which core metabolic networks, localized on satellite, specialized neochromosomes can be swapped for alternative configurations and serve as landing pads for the addition of functionalities.

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Dimethyl sulfide (DMS) is a sulfur containing volatile that enhances general fruity aroma and imparts aromatic notes in wine. The most important precursor of DMS is S-methylmethionine (SMM), which is synthesized by grapes and can be metabolized by the yeast S. cerevisiae during wine fermentation. Precursor molecules left after fermentation are chemically converted to DMS during wine maturation, meaning that wine DMS levels are determined by the amount of remaining precursors at bottling. To elucidate SMM metabolism in yeast we performed quantitative trait locus (QTL) mapping using a population of 130 F2-segregants obtained from a cross between two wine yeast strains, and we detected one major QTL explaining almost 30% of trait variation. Within the QTL, gene YLL058W and SMM transporter gene MMP1 were found to influence SMM metabolism, from which MMP1 has the bigger impact. We identified and characterized a variant coding for a truncated transporter with superior SMM preserving attributes. A population analysis with 85 yeast strains from different origins revealed a significant association of the variant to flor strains and minor occurrence in cheese and wine strains. These results will help selecting and improving S. cerevisiae strains for the production of wine and other fermented foods containing DMS such as cheese or beer.

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Although transplantation of single genes in yeast plays a key role in elucidating gene functionality in metazoans, technical challenges hamper humanization of full pathways and processes. Empowered by advances in synthetic biology, this study demonstrates the feasibility and implementation of full humanization of glycolysis in yeast. Single gene and full pathway transplantation revealed the remarkable conservation of glycolytic and moonlighting functions and, combined with evolutionary strategies, brought to light context-dependent responses. Human hexokinase 1 and 2, but not 4, required mutations in their catalytic or allosteric sites for functionality in yeast, whereas hexokinase 3 was unable to complement its yeast ortholog. Comparison with human tissues cultures showed preservation of turnover numbers of human glycolytic enzymes in yeast and human cell cultures. This demonstration of transplantation of an entire essential pathway paves the way for establishment of species-, tissue-, and disease-specific metazoan models.

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Even for the genetically accessible yeast Saccharomyces cerevisiae, the CRISPR-Cas DNA editing technology has strongly accelerated and facilitated strain construction. Several methods have been validated for fast and highly efficient single editing events, and diverse approaches for multiplex genome editing have been described in the literature by means of SpCas9 or FnCas12a endonucleases and their associated guide RNAs (gRNAs). The gRNAs used to guide the Cas endonuclease to the editing site are typically expressed from plasmids using native Pol II or Pol III RNA polymerases. These gRNA expression plasmids require laborious, time-consuming cloning steps, which hampers their implementation for academic and applied purposes. In this study, we explore the potential of expressing gRNA from linear DNA fragments using the T7 RNA polymerase (T7RNAP) for single and multiplex genome editing in Saccharomyces cerevisiae. Using FnCas12a, this work demonstrates that transforming short, linear DNA fragments encoding gRNAs in yeast strains expressing T7RNAP promotes highly efficient single and duplex DNA editing. These DNA fragments can be custom ordered, which makes this approach highly suitable for high-Throughput strain construction. This work expands the CRISPR toolbox for large-scale strain construction programs in S. cerevisiae and promises to be relevant for other less genetically accessible yeast species.

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Neocallimastigomycetes are unique examples of strictly anaerobic eukaryotes. This study investigates how these anaerobic fungi bypass reactions involved in synthesis of pyridine nucleotide cofactors and coenzyme A that, in canonical fungal pathways, require molecular oxygen. Analysis of Neocallimastigomycetes proteomes identified a candidate L-aspartate-decarboxylase (AdcA) and L-aspartate oxidase (NadB) and quinolinate synthase (NadA), constituting putative oxygen-independent bypasses for coenzyme A synthesis and pyridine nucleotide cofactor synthesis. The corresponding gene sequences indicated acquisition by ancient horizontal gene transfer (HGT) events involving bacterial donors. To test whether these enzymes suffice to bypass corresponding oxygen-requiring reactions, they were introduced into fms1∆ and bna2∆ Saccharomyces cerevisiae strains. Expression of nadA and nadB from Piromyces finnis and adcA from Neocallimastix californiae conferred cofactor prototrophy under aerobic and anaerobic conditions. This study simulates how HGT can drive eukaryotic adaptation to anaerobiosis and provides a basis for elimination of auxotrophic requirements in anaerobic industrial applications of yeasts and fungi. IMPORTANCE NAD (NAD ⁺) and coenzyme A (CoA) are central metabolic cofactors whose canonical biosynthesis pathways in fungi require oxygen. Anaerobic gut fungi of the Neocallimastigomycota phylum are unique eukaryotic organisms that adapted to anoxic environments. Analysis of Neocallimastigomycota genomes revealed that these fungi might have developed oxygen-independent biosynthetic pathways for NAD ⁺ and CoA biosynthesis, likely acquired through horizontal gene transfer (HGT) from prokaryotic donors. We confirmed functionality of these putative pathways under anaerobic conditions by heterologous expression in the yeast Saccharomyces cerevisiae. This approach, combined with sequence comparison, offers experimental insight on whether HGT events were required and/or sufficient for acquiring new traits. Moreover, our results demonstrate an engineering strategy for enabling S. cerevisiae to grow anaerobically in the absence of the precursor molecules pantothenate and nicotinate, thereby contributing to alleviate oxygen requirements and to move closer to prototrophic anaerobic growth of this industrially relevant yeast.

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The construction of microbial cell factories for sustainable production of chemicals and pharmaceuticals requires extensive genome engineering. Using Saccharomyces cerevisiae, this study proposes synthetic neochromosomes as orthogonal expression platforms for rewiring native cellular processes and implementing new functionalities. Capitalizing the powerful homologous recombination capability of S. cerevisiae, modular neochromosomes of 50 and 100 kb were fully assembled de novo from up to 44 transcriptional-unit-sized fragments in a single transformation. These assemblies were remarkably efficient and faithful to their in silico design. Neochromosomes made of non-coding DNA were stably replicated and segregated irrespective of their size without affecting the physiology of their host. These non-coding neochromosomes were successfully used as landing pad and as exclusive expression platform for the essential glycolytic pathway. This work pushes the limit of DNA assembly in S. cerevisiae and paves the way for de novo designer chromosomes as modular genome engineering platforms in S. cerevisiae.

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Metabolic capabilities of cells are not only defined by their repertoire of enzymes and metabolites, but also by availability of enzyme cofactors. The molybdenum cofactor (Moco) is widespread among eukaryotes but absent from the industrial yeast Saccharomyces cerevisiae. No less than 50 Moco-dependent enzymes covering over 30 catalytic activities have been described to date, introduction of a functional Moco synthesis pathway offers interesting options to further broaden the biocatalytic repertoire of S. cerevisiae. In this study, we identified seven Moco biosynthesis genes in the non-conventional yeast Ogataea parapolymorpha by SpyCas9-mediated mutational analysis and expressed them in S. cerevisiae. Functionality of the heterologously expressed Moco biosynthesis pathway in S. cerevisiae was assessed by co-expressing O. parapolymorpha nitrate-assimilation enzymes, including the Moco-dependent nitrate reductase. Following two-weeks of incubation, growth of the engineered S. cerevisiae strain was observed on nitrate as sole nitrogen source. Relative to the rationally engineered strain, the evolved derivatives showed increased copy numbers of the heterologous genes, increased levels of the encoded proteins and a 5-fold higher nitrate-reductase activity in cell extracts. Growth at nM molybdate concentrations was enabled by co-expression of a Chlamydomonas reinhardtii high-affinity molybdate transporter. In serial batch cultures on nitrate-containing medium, a non-engineered S. cerevisiae strain was rapidly outcompeted by the spoilage yeast Brettanomyces bruxellensis. In contrast, an engineered and evolved nitrate-assimilating S. cerevisiae strain persisted during 35 generations of co-cultivation. This result indicates that the ability of engineered strains to use nitrate may be applicable to improve competitiveness of baker's yeast in industrial processes upon contamination with spoilage yeasts.

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An oxygen requirement for de novo biotin synthesis in Saccharomyces cerevisiae precludes the application of biotin-prototrophic strains in anoxic processes that use biotin-free media. To overcome this issue, this study explores introduction of the oxygen-independent Escherichia coli biotin-biosynthesis pathway in S. cerevisiae. Implementation of this pathway required expression of seven E. coli genes involved in fatty-acid synthesis and three E. coli genes essential for the formation of a pimelate thioester, key precursor of biotin synthesis. A yeast strain expressing these genes readily grew in biotin-free medium, irrespective of the presence of oxygen. However, the engineered strain exhibited specific growth rates 25% lower in biotin-free media than in biotin-supplemented media. Following adaptive laboratory evolution in anoxic cultures, evolved cell lines that no longer showed this growth difference in controlled bioreactors, were characterized by genome sequencing and proteome analyses. The evolved isolates exhibited a whole-genome duplication accompanied with an alteration in the relative gene dosages of biosynthetic pathway genes. These alterations resulted in a reduced abundance of the enzymes catalyzing the first three steps of the E. coli biotin pathway. The evolved pathway configuration was reverse engineered in the diploid industrial S. cerevisiae strain Ethanol Red. The resulting strain grew at nearly the same rate in biotin-supplemented and biotin-free media non-controlled batches performed in an anaerobic chamber. This study established an unique genetic engineering strategy to enable biotin-independent anoxic growth of S. cerevisiae and demonstrated its portability in industrial strain backgrounds.

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