Stress-tolerant non-conventional microbes enable next-generation chemical biosynthesis.

Microbial chemical production is a rapidly growing industry, with much of the growth fueled by advances in synthetic biology. New approaches have enabled rapid strain engineering for the production of various compounds; however, translation to industry is often problematic because native phenotypes of model hosts prevent the design of new low-cost bioprocesses. Here, we argue for a new approach that leverages the native stress-tolerant phenotypes of non-conventional microbes that directly address design challenges from the outset. Growth at high temperature, high salt and solvent concentrations, and low pH can enable cost savings by reducing the energy required for product separation, bioreactor cooling, and maintaining sterile conditions. These phenotypes have the added benefit of allowing for the use of low-cost sugar and water resources. Non-conventional hosts are needed because these phenotypes are polygenic and thus far have proven difficult to recapitulate in the common hosts Escherichia coli and Saccharomyces cerevisiae.

Genome-wide CRISPR-Cas9 screen reveals a persistent null-hyphal phenotype that maintains high carotenoid production in Yarrowia lipolytica.

Yarrowia lipolytica is a metabolic engineering host of growing industrial interest due to its ability to metabolize hydrocarbons, fatty acids, glycerol, and other renewable carbon sources. This dimorphic yeast undergoes a stress-induced transition to a multicellular hyphal state, which can negatively impact biosynthetic activity, reduce oxygen and nutrient mass transfer in cell cultures, and increase culture viscosity. Identifying mutations that prevent the formation of hyphae would help alleviate the bioprocess challenges that they create. To this end, we conducted a genome-wide CRISPR screen to identify genetic knockouts that prevent the transition to hyphal morphology. The screen identified five mutants with a null-hyphal phenotype-ΔRAS2, ΔRHO5, ΔSFL1, ΔSNF2, and ΔPAXIP1. Of these hits, only ΔRAS2 suppressed hyphal formation in an engineered lycopene production strain over a multiday culture. The RAS2 knockout was also the only genetic disruption characterized that did not affect lycopene production, producing more than 5 mg L-1 OD-1 from a heterologous pathway with enhanced carbon flux through the mevalonate pathway. These data suggest that a ΔRAS2 mutant of Y. lipolytica could prove useful in engineering a metabolic engineering host of the production of carotenoids and other biochemicals.

Validating genome-wide CRISPR-Cas9 function improves screening in the oleaginous yeast Yarrowia lipolytica.

Genome-wide mutational screens are central to understanding the genetic underpinnings of evolved and engineered phenotypes. The widespread adoption of CRISPR-Cas9 genome editing has enabled such screens in many organisms, but identifying functional sgRNAs still remains a challenge. Here, we developed a methodology to quantify the cutting efficiency of each sgRNA in a genome-scale library, and in doing so improve screens in the biotechnologically important yeast Yarrowia lipolytica. Screening in the presence and absence of native DNA repair enabled high-throughput quantification of sgRNA function leading to the identification of high efficiency sgRNAs that cover 94% of genes. Library validation enhanced the classification of essential genes by identifying inactive guides that create false negatives and mask the effects of successful disruptions. Quantification of guide effectiveness also creates a dataset from which determinants of CRISPR-Cas9 can be identified. Finally, application of the library identified novel mutations for metabolic engineering of high lipid accumulation.

CRISPRi repression of nonhomologous end-joining for enhanced genome engineering via homologous recombination in Yarrowia lipolytica.

In many organisms of biotechnological importance precise genome editing is limited by inherently low homologous recombination (HR) efficiencies. A number of strategies exist to increase the effectiveness of this native DNA repair pathway; however, most strategies rely on permanently disabling competing repair pathways, thus reducing an organism's capacity to repair naturally occurring double strand breaks. Here, we describe a CRISPR interference (CRISPRi) system for gene repression in the oleochemical-producing yeast Yarrowia lipolytica. By using a multiplexed sgRNA targeting strategy, we demonstrate efficient repression of eight out of nine targeted genes to enhance HR. Strains with nonhomologous end-joining repressed were shown to have increased rates of HR when transformed with a linear DNA fragment with homology to a genomic locus. With multiplexed targeting of KU70 and KU80, and enhanced repression with Mxi1 fused to deactivated Cas9 (dCas9), rates of HR as high as 90% were achieved. The developed CRISPRi system enables enhanced HR in Y. lipolytica without permanent genetic knockouts and promises to be a potent tool for other metabolic engineering, synthetic biology, and functional genomics studies.

acCRISPR: an activity-correction method for improving the accuracy of CRISPR screens.

High throughput CRISPR screens are revolutionizing the way scientists unravel the genetic underpinnings of engineered and evolved phenotypes. One of the critical challenges in accurately assessing screening outcomes is accounting for the variability in sgRNA cutting efficiency. Poorly active guides targeting genes essential to screening conditions obscure the growth defects that are expected from disrupting them. Here, we develop acCRISPR, an end-to-end pipeline that identifies essential genes in pooled CRISPR screens using sgRNA read counts obtained from next-generation sequencing. acCRISPR uses experimentally determined cutting efficiencies for each guide in the library to provide an activity correction to the screening outcomes via calculation of an optimization metric, thus determining the fitness effect of disrupted genes. CRISPR-Cas9 and -Cas12a screens were carried out in the non-conventional oleaginous yeast Yarrowia lipolytica and acCRISPR was used to determine a high-confidence set of essential genes for growth under glucose, a common carbon source used for the industrial production of oleochemicals. acCRISPR was also used in screens quantifying relative cellular fitness under high salt conditions to identify genes that were related to salt tolerance. Collectively, this work presents an experimental-computational framework for CRISPR-based functional genomics studies that may be expanded to other non-conventional organisms of interest.

Genome-wide functional screens enable the prediction of high activity CRISPR-Cas9 and -Cas12a guides in Yarrowia lipolytica.

Genome-wide functional genetic screens have been successful in discovering genotype-phenotype relationships and in engineering new phenotypes. While broadly applied in mammalian cell lines and in E. coli, use in non-conventional microorganisms has been limited, in part, due to the inability to accurately design high activity CRISPR guides in such species. Here, we develop an experimental-computational approach to sgRNA design that is specific to an organism of choice, in this case the oleaginous yeast Yarrowia lipolytica. A negative selection screen in the absence of non-homologous end-joining, the dominant DNA repair mechanism, was used to generate single guide RNA (sgRNA) activity profiles for both SpCas9 and LbCas12a. This genome-wide data served as input to a deep learning algorithm, DeepGuide, that is able to accurately predict guide activity. DeepGuide uses unsupervised learning to obtain a compressed representation of the genome, followed by supervised learning to map sgRNA sequence, genomic context, and epigenetic features with guide activity. Experimental validation, both genome-wide and with a subset of selected genes, confirms DeepGuide's ability to accurately predict high activity sgRNAs. DeepGuide provides an organism specific predictor of CRISPR guide activity that with retraining could be applied to other fungal species, prokaryotes, and other non-conventional organisms.

CRISPR Interference and Activation to Modulate Transcription in Yarrowia lipolytica.

Recent developments in RNA-guided nuclease technologies have advanced the engineering of a wide range of organisms, including the nonconventional yeast Yarrowia lipolytica. Y. lipolytica has been the focus of a range of synthetic biology and metabolic engineering studies due to its high capacity to synthesize and accumulate intracellular lipids. The CRISPR-Cas9 system from Streptococcus pyogenes has been successfully adapted and used for genome editing in Y. lipolytica. However, as engineered strains are moved closer to industrialization, the need for finer control of transcription is still present. To overcome this challenge, we have developed CRISPR interference (CRISPRi) and CRISPR activation (CRISPRa) systems to allow modulating the transcription of endogenous genes. We begin this protocol chapter by describing how to use the CRISPRi system to repress expression of any gene in Y. lipolytica. A second method describes how to use the CRISPRa system to increase expression of native Y. lipolytica genes. Finally, we describe how CRISPRi or CRISPRa vectors can be combined to enable multiplexed activation or repression of more than one gene. The implementation of CRISPRi and CRISPRa systems improves our ability to control gene expression in Y. lipolytica and promises to enable more advanced synthetic biology and metabolic engineering studies in this host.

Controlled intracellular trafficking alleviates an expression bottleneck in S. cerevisiae ester biosynthesis.

In metabolic engineering, most available pathway engineering strategies aim to control enzyme expression by making changes at the transcriptional level with an underlying assumption that translation and functional expression follow suit. In this work, we engineer expression of a key reaction step in medium chain ester biosynthesis that does not follow this common assumption. The native Saccharomyces cerevisiae alcohol acyltransferses Eeb1 and Eht1 condense acyl-CoAs with ethanol to produce the corresponding ester, a reaction that is rate limiting in engineering ester biosynthesis pathways. By changing the N- and C-termini of Eeb1 to those of Eht1, Eeb1 localization is changed from the mitochondria to lipid droplets. The change has no significant effect on transcription, but increases protein expression by 23-fold thus enabling a 3-fold increase in enzyme activity. This system demonstrates one example of the impact of protein trafficking on functional pathway expression, and will guide future metabolic engineering of ester biosynthesis and, potentially, other pathways with critical membrane-bound enzymes.

CRISPR-Cas9-Mediated Genome Editing and Transcriptional Control in Yarrowia lipolytica.

The discovery and adaptation of RNA-guided nucleases has resulted in the rapid development of efficient, scalable, and easily accessible synthetic biology tools for targeted genome editing and transcriptional control. In these systems, for example CRISPR-Cas9 from Streptococcus pyogenes, a protein with nuclease activity is targeted to a specific nucleotide sequence by a short RNA molecule, whereupon binding it cleaves the targeted nucleotide strand. To extend this genome-editing ability to the industrially important oleaginous yeast Yarrowia lipolytica, we developed a set of easily usable and effective CRISPR-Cas9 episomal vectors. In this protocols chapter, we first present a method by which arbitrary protein-coding genes can be disrupted via indel formation after CRISPR-Cas9 targeting. A second method demonstrates how the same CRISPR-Cas9 system can be used to induce markerless gene cassette integration into the genome by inducing homologous recombination after DNA cleavage by Cas9. Finally, we describe how a catalytically inactive form of Cas9 fused to a transcriptional repressor can be used to control transcription of native genes in Y. lipolytica. The CRISPR-Cas9 tools and strategies described here greatly increase the types of genome editing and transcriptional control that can be achieved in Y. lipolytica, and promise to facilitate more advanced engineering of this important oleaginous host.

Design of Hybrid RNA Polymerase III Promoters for Efficient CRISPR-Cas9 Function.

The discovery of the CRISPR-Cas9 system from Streptococcus pyogenes has allowed the development of genome engineering tools in a variety of organisms. A frequent limitation in CRISPR-Cas9 function is adequate expression levels of sgRNA. This protocol provides a strategy to construct hybrid RNA polymerase III (Pol III) promoters that facilitate high expression of sgRNA and improved CRISPR-Cas9 function. We provide selection criteria of Pol III promoters, efficient promoter construction methods, and a sample screening technique to test the efficiency of the hybrid promoters. A hybrid promoter system developed for Yarrowia lipolytica will serve as a model.

Highly Multiplexed CRISPRi Repression of Respiratory Functions Enhances Mitochondrial Localized Ethyl Acetate Biosynthesis in Kluyveromyces marxianus.

The emergence of CRISPR-Cas9 for targeted genome editing and regulation has enabled the manipulation of desired traits and enhanced strain development of nonmodel microorganisms. The natural capacity of the yeast Kluyveromyces marxianus to produce volatile esters at high rate and at elevated temperatures make it a potentially valuable production platform for industrial biotechnology. Here, we identify the native localization of ethyl acetate biosynthesis in K. marxianus and use this information to develop a multiplexed CRISPRi system for redirecting carbon flux along central metabolic pathways, increasing ethyl acetate productivity. First, we identified the primary pathways of precursor and acetate ester biosynthesis. A genetic knockout screen revealed that the alcohol acetyltransferase Eat1 is the critical enzyme for ethyl, isoamyl, and phenylethyl acetate production. Truncation studies revealed that high ester biosynthesis is contingent on Eat1 mitochondrial localization. As ethyl acetate is formed from the condensation of ethanol and acetyl-CoA, we modulated expression of the TCA cycle and electron transport chain genes using a highly multiplexed CRISPRi approach. The simultaneous knockdown of ACO2b, SDH2, RIP1, and MSS51 resulted in a 3.8-fold increase in ethyl acetate productivity over the already high natural capacity. This work demonstrates that multiplexed CRISPRi regulation of central carbon flux, supported by a fundamental understanding of pathway biochemistry, is a potent strategy for metabolic engineering in nonconventional microorganisms.

Multiplexed CRISPR Activation of Cryptic Sugar Metabolism Enables Yarrowia Lipolytica Growth on Cellobiose.

The yeast Yarrowia lipolytica has been widely studied for its ability to synthesize and accumulate intracellular lipids to high levels. Recent studies have identified native genes that enable growth on biomass-derived sugars, but these genes are not sufficiently expressed to facilitate robust metabolism. In this work, a CRISPR-dCas9 activation (CRISPRa) system in Y. lipolytica is developed and is used it to activate native ß-glucosidase expression to support growth on cellobiose. A series of different transcriptional activators are compared for their effectiveness in Y. lipolytica, with the synthetic tripartite activator VPR yielding the highest activation. A VPR-dCas9 fusion is then targeted to various locations in a synthetic promoter driving hrGFP expression, and activation is achieved. Subsequently, the CRISPRa system is used to activate transcription of two different native ß-glucosidase genes, facilitating enhanced growth on cellobiose as the sole carbon source. This work expands the synthetic biology toolbox for metabolic engineering in Y. lipolytica and demonstrates how the programmability of the CRISPR-Cas9 system can enable facile investigation of transcriptionally silent regions of the genome.

Genome and metabolic engineering in non-conventional yeasts: Current advances and applications.

Microbial production of chemicals and proteins from biomass-derived and waste sugar streams is a rapidly growing area of research and development. While the model yeast Saccharomyces cerevisiae is an excellent host for the conversion of glucose to ethanol, production of other chemicals from alternative substrates often requires extensive strain engineering. To avoid complex and intensive engineering of S. cerevisiae, other yeasts are often selected as hosts for bioprocessing based on their natural capacity to produce a desired product: for example, the efficient production and secretion of proteins, lipids, and primary metabolites that have value as commodity chemicals. Even when using yeasts with beneficial native phenotypes, metabolic engineering to increase yield, titer, and production rate is essential. The non-conventional yeasts Kluyveromyces lactis, K. marxianus, Scheffersomyces stipitis, Yarrowia lipolytica, Hansenula polymorpha and Pichia pastoris have been developed as eukaryotic hosts because of their desirable phenotypes, including thermotolerance, assimilation of diverse carbon sources, and high protein secretion. However, advanced metabolic engineering in these yeasts has been limited. This review outlines the challenges of using non-conventional yeasts for strain and pathway engineering, and discusses the developed solutions to these problems and the resulting applications in industrial biotechnology.

Host and Pathway Engineering for Enhanced Lycopene Biosynthesis in Yarrowia lipolytica.

Carotenoids are a class of molecules with commercial value as food and feed additives with nutraceutical properties. Shifting carotenoid synthesis from petrochemical-based precursors to bioproduction from sugars and other biorenewable carbon sources promises to improve process sustainability and economics. In this work, we engineered the oleaginous yeast Yarrowia lipolytica to produce the carotenoid lycopene. To enhance lycopene production, we tested a series of strategies to modify host cell physiology and metabolism, the most successful of which were mevalonate pathway overexpression and alleviating auxotrophies previously engineered into the PO1f strain of Y. lipolytica. The beneficial engineering strategies were combined into a single strain, which was then cultured in a 1-L bioreactor to produce 21.1 mg/g DCW. The optimized strain overexpressed a total of eight genes including two copies of HMG1, two copies of CrtI, and single copies of MVD1, EGR8, CrtB, and CrtE. Recovering leucine and uracil biosynthetic capacity also produced significant enhancement in lycopene titer. The successful engineering strategies characterized in this work represent a significant increase in understanding carotenoid biosynthesis in Y. lipolytica, not only increasing lycopene titer but also informing future studies on carotenoid biosynthesis.

Standardized Markerless Gene Integration for Pathway Engineering in Yarrowia lipolytica.

The yeast Yarrowia lipolytica is a promising microbial host due to its native capacity to produce lipid-based chemicals. Engineering stable production strains requires genomic integration of modified genes, avoiding episomal expression that requires specialized media to maintain selective pressures. Here, we develop a CRISPR-Cas9-based tool for targeted, markerless gene integration into the Y. lipolytica genome. A set of genomic loci was screened to identify sites that were accepting of gene integrations without impacting cell growth. Five sites were found to meet these criteria. Expression levels from a GFP expression cassette were consistent when inserted into AXP, XPR2, A08, and D17, with reduced expression from MFE1. The standardized tool is comprised of five pairs of plasmids (one homologous donor plasmid and a CRISPR-Cas9 expression plasmid), with each pair targeting gene integration into one of the characterized sites. To demonstrate the utility of the tool we rapidly engineered a semisynthetic lycopene biosynthesis pathway by integrating four different genes at different loci. The capability to integrate multiple genes without the need for marker recovery and into sites with known expression levels will enable more rapid and reliable pathway engineering in Y. lipolytica.

CRISPR-Cas9-enabled genetic disruptions for understanding ethanol and ethyl acetate biosynthesis in Kluyveromyces marxianus.

BACKGROUND: The thermotolerant yeast Kluyveromyces marxianus shows promise as an industrial host for the biochemical production of fuels and chemicals. Wild-type strains are known to ferment high titers of ethanol and can effectively convert a wide range of C5, C6, and C12 sugars into the volatile short-chain ester ethyl acetate. Strain engineering, however, has been limited due to a lack of advanced genome-editing tools and an incomplete understanding of ester and ethanol biosynthesis. RESULTS: Enabled by the design of hybrid RNA polymerase III promoters, this work adapts the CRISPR-Cas9 system from Streptococcus pyogenes for use in K. marxianus. The system was used to rapidly create functional disruptions to alcohol dehydrogenase (ADH) and alcohol-O-acetyltransferase (ATF) genes with putative function in ethyl acetate and ethanol biosynthesis. Screening of the KmATF disrupted strain revealed that Atf activity contributes to ethyl acetate biosynthesis, but the knockout reduced ethyl acetate titers by only ~15%. Overexpression experiments revealed that KmAdh7 can catalyze the oxidation of hemiacetal to ethyl acetate. Finally, analysis of the KmADH2 disrupted strain showed that the knockout almost completely eliminated ethanol production and resulted in the accumulation of acetaldehyde. CONCLUSIONS: Newly designed RNA polymerase III promoters for sgRNA expression in K. marxianus enable a CRISPR-Cas9 genome-editing system for the thermotolerant yeast. This system was used to disrupt genes involved in ethyl acetate biosynthesis, specifically KmADH1-7 and KmATF. KmAdh2 was found to be critical for aerobic and anaerobic ethanol production. Aerobically produced ethanol supplies the biosynthesis of ethyl acetate catalyzed by KmAtf. KmAdh7 was found to exhibit activity toward the oxidation of hemiacetal, a possible alternative route for the synthesis of ethyl acetate.

Synthetic RNA Polymerase III Promoters Facilitate High-Efficiency CRISPR-Cas9-Mediated Genome Editing in Yarrowia lipolytica.

The oleaginous yeast Yarrowia lipolytica is a valuable microbial host for chemical production because it has a high capacity to synthesize, modify, and store intracellular lipids; however, rapid strain development has been hampered by the limited availability of genome engineering tools. We address this limitation by adapting the CRISPR-Cas9 system from Streptococcus pyogenes for markerless gene disruption and integration in Y. lipolytica. Single gene disruption efficiencies of 92% and higher were achieved when single guide RNAs (sgRNA) were transcribed with synthetic hybrid promoters that combine native RNA polymerase III (Pol III) promoters with tRNA. The Pol III-tRNA hybrid promoters exploit endogenous tRNA processing to produce mature sgRNA for Cas9 targeting. The highest efficiencies were achieved with a SCR1'-tRNA(Gly) promoter and Y. lipolytica codon-optimized Cas9 expressed from a UAS1B8-TEF promoter. Cotransformation of the Cas9 and sgRNA expressing plasmid with a homologous recombination donor plasmid resulted in markerless homologous recombination efficiency of over 64%. Homologous recombination was observed in 100% of transformants when nonhomologous end joining was disrupted. The end result of these studies was the development of pCRISPRyl, a modular tool for markerless gene disruption and integration in Y. lipolytica.