A highly efficient single-step,markerless strategy for multi-copy chromosomal integration of large biochemical pathways in Saccharomyces cerevisiae

Despite recent advances in genome editing capabilities for the model organism Saccharomyces cerevisiae, the chromosomal integration of large biochemical pathways for stable industrial production remains challenging. In this work, we developed a simple platform for high-efficiency, single-step, markerless, multi-copy chromosomal integration of full biochemical pathways in Saccharomyces cerevisiae. In this Di-CRISPR (delta integration CRISPR-Cas) platform based on the Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) and CRISPR-associated systems (Cas), we specifically designed guide RNA sequences to target multiple delta sites in the yeast genome. The generation of double stranded breaks at the delta sites allowed simultaneous integration of multiple copies of linearized donor DNA containing large biochemical pathways. With our newly developed Di-CRISPR platform, we were able to attain highly efficient and markerless integration of large biochemical pathways and achieve an unprecedented 18-copy genomic integration of a 24 kb combined xylose utilization and (R,R)-2,3-butanediol (BDO) production pathway in a single step, thus generating a strain that was able to produce BDO directly from xylose. The simplicity and high efficiency of the Di-CRISPR platform could provide a superior alternative to high copy plasmids and would render this platform an invaluable tool for genome editing and metabolic engineering in S. cerevisiae.     

Expanding the CRISPR/Cas9 toolkit for Pichia pastoris with efficient donor integration and alternative resistance markers

Komagataella phaffii (syn. Pichia pastoris) is one of the most commonly used host systems for recombinant protein expression. Achieving targeted genetic modifications had been hindered by low frequencies of homologous recombination (HR). Recently, a CRISPR/Cas9 genome editing system has been implemented for P. pastoris enabling gene knockouts based on indels (insertion, deletions) via non‐homologous end joining (NHEJ) at near 100% efficiency. However, specifically integrating homologous donor cassettes via HR for replacement studies had proven difficult resulting at most in ～20% correct integration using CRISPR/Cas9. Here, we demonstrate the CRISPR/Cas9 mediated integration of markerless donor cassettes at efficiencies approaching 100% using a ku70 deletion strain. The Ku70p is involved in NHEJ repair and lack of the protein appears to favor repair via HR near exclusively. While the absolute number of transformants in the Δku70 strain is reduced, virtually all surviving transformants showed correct integration. In the wildtype strain, markerless donor cassette integration was also improved up to 25‐fold by placing an autonomously replicating sequence (ARS) on the donor cassette. Alternative strategies for improving donor cassette integration using a Cas9 nickase variant or reducing off targeting associated toxicity using a high fidelity Cas9 variant were so far not successful in our hands in P. pastoris. Furthermore we provide Cas9/gRNA expression plasmids with a Geneticin resistance marker which proved to be versatile tools for marker recycling. The reported CRSIPR‐Cas9 tools can be applied for modifying existing production strains and also pave the way for markerless whole genome modification studies in P. pastoris.     

CRISPR-Cas9 for the genome engineering of cyanobacteria and succinate production

Cyanobacteria hold promise as a cell factory for producing biofuels and bio-derived chemicals, but genome engineering of cyanobacteria such as Synechococcus elongatus PCC 7942 poses challenges because of their oligoploidy nature and long-term instability of the introduced gene. CRISPR-Cas9 is a newly developed RNA-guided genome editing system, yet its application for cyanobacteria engineering has yet to be reported. Here we demonstrated that CRISPR-Cas9 system can effectively trigger programmable double strand break (DSB) at the chromosome of PCC 7942 and provoke cell death. With the co-transformation of template plasmid harboring the gene cassette and flanking homology arms, CRISPR-Cas9-mediated DSB enabled precise gene integration, ameliorated the homologous recombination efficiency and allowed the use of lower amount of template DNA and shorter homology arms. The CRISPR-Cas9-induced cell death imposed selective pressure and enhanced the chance of concomitant integration of gene cassettes into all chromosomes of PCC 7942, hence accelerating the process of obtaining homogeneous and stable recombinant strains. We further explored the feasibility of engineering cyanobacteria by CRISPR-Cas9-assisted simultaneous glgc knock-out and gltA/ppc knock-in, which improved the succinate titer to 435.0±35.0 μg/L, an ≈11-fold increase when compared with that of the wild-type cells. These data altogether justify the use of CRISPR-Cas9 for genome engineering and manipulation of metabolic pathways in cyanobacteria.     

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.     

Genome engineering using a synthetic gene circuit in Bacillus subtilis

Genome engineering without leaving foreign DNA behind requires an efficient counter-selectable marker system. Here, we developed a genome engineering method in Bacillus subtilis using a synthetic gene circuit as a counter-selectable marker system. The system contained two repressible promoters (B. subtilis xylA (P_xyl) and spac (P_spac)) and two repressor genes (lacI and xylR). P_xyl-lacI was integrated into the B. subtilis genome with a target gene containing a desired mutation. The xylR and P_spac–chloramphenicol resistant genes (cat) were located on a helper plasmid. In the presence of xylose, repression of XylR by xylose induced LacI expression, the LacIs repressed the P_spac promoter and the cells become chloramphenicol sensitive. Thus, to survive in the presence of chloramphenicol, the cell must delete P_xyl-lacI by recombination between the wild-type and mutated target genes. The recombination leads to mutation of the target gene. The remaining helper plasmid was removed easily under the chloramphenicol absent condition. In this study, we showed base insertion, deletion and point mutation of the B. subtilis genome without leaving any foreign DNA behind. Additionally, we successfully deleted a 2-kb gene (amyE) and a 38-kb operon (ppsABCDE). This method will be useful to construct designer Bacillus strains for various industrial applications.     

Engineering strategies for enhanced production of protein and bio-products in Pichia pastoris: A review

Pichia pastoris has been recognized as one of the most industrially important hosts for heterologous protein production. Despite its high protein productivity, the optimization of P. pastoris cultivation is still imperative due to strain- and product-specific challenges such as promoter strength, methanol utilization type and oxygen demand. To address the issues, strategies involving genetic and process engineering have been employed. Optimization of codon usage and gene dosage, as well as engineering of promoters, protein secretion pathways and methanol metabolic pathways have proved beneficial to innate protein expression levels. Large-scale production of proteins via high cell density fermentation additionally relies on the optimization of process parameters including methanol feed rate, induction temperature and specific growth rate. Recent progress related to the enhanced production of proteins in P. pastoris via various genetic engineering and cultivation strategies are reviewed. Insight into the regulation of the P. pastoris alcohol oxidase 1 (AOX1) promoter and the development of methanol-free systems are highlighted. Novel cultivation strategies such as mixed substrate feeding are discussed. Recent advances regarding substrate and product monitoring techniques are also summarized. Application of P. pastoris to the production of biodiesel and other value-added products via metabolic engineering are also reviewed. P. pastoris is becoming an indispensable platform through the use of these combined engineering strategies.     

Efficient genome editing in Caenorhabditis elegans by CRISPR-targeted homologous recombination

Cas9 is an RNA-guided double-stranded DNA nuclease that participates in clustered regularly interspaced short palindromic repeats (CRISPR)-mediated adaptive immunity in prokaryotes. CRISPR–Cas9 has recently been used to generate insertion and deletion mutations in Caenorhabditis elegans, but not to create tailored changes (knock-ins). We show that the CRISPR–CRISPR-associated (Cas) system can be adapted for efficient and precise editing of the C. elegans genome. The targeted double-strand breaks generated by CRISPR are substrates for transgene-instructed gene conversion. This allows customized changes in the C. elegans genome by homologous recombination: sequences contained in the repair template (the transgene) are copied by gene conversion into the genome. The possibility to edit the C. elegans genome at selected locations will facilitate the systematic study of gene function in this widely used model organism.     

Insertional mutagenesis based on illegitimate recombination in Schizosaccharomyces pombe

An efficient insertional mutagenesis system has been developed for Schizosaccharomyces pombe based on linear PCR-generated cassettes containing selectable markers. It depends upon illegitimate recombination for integration into the genome. Various selectable markers of different sizes can be used to obtain sufficiently high transformation and integration frequencies. Based on Southern blotting, a single insertion is found in each strain and integration sites are broadly distributed in the genome. Sequence analysis of the insert junctions frequently reveals small regions of homology (4–10 bp) between the ends of the integrated cassette and the disrupted gene. The system has been used for simple genetic screens of various types and as a promoter trap for in-frame GFP fusions.     

Rapid construction of insulated genetic circuits via synthetic sequence-guided isothermal assembly

In vitro recombination methods have enabled one-step construction of large DNA sequences from multiple parts. Although synthetic biological circuits can in principle be assembled in the same fashion, they typically contain repeated sequence elements such as standard promoters and terminators that interfere with homologous recombination. Here we use a computational approach to design synthetic, biologically inactive unique nucleotide sequences (UNSes) that facilitate accurate ordered assembly. Importantly, our designed UNSes make it possible to assemble parts with repeated terminator and insulator sequences, and thereby create insulated functional genetic circuits in bacteria and mammalian cells. Using UNS-guided assembly to construct repeating promoter-gene-terminator parts, we systematically varied gene expression to optimize production of a deoxychromoviridans biosynthetic pathway in Escherichia coli. We then used this system to construct complex eukaryotic AND-logic gates for genomic integration into embryonic stem cells. Construction was performed by using a standardized series of UNS-bearing BioBrick-compatible vectors, which enable modular assembly and facilitate reuse of individual parts. UNS-guided isothermal assembly is broadly applicable to the construction and optimization of genetic circuits and particularly those requiring tight insulation, such as complex biosynthetic pathways, sensors, counters and logic gates.     

Efficient Genome Editing in Caenorhabditis elegans with a Toolkit of Dual-Marker Selection Cassettes

Use of the CRISPR/Cas9 RNA-guided endonuclease complex has recently enabled the generation of double-strand breaks virtually anywhere in the C. elegans genome. Here, we present an improved strategy that makes all steps in the genome editing process more efficient. We have created a toolkit of template-mediated repair cassettes that contain an antibiotic resistance gene to select for worms carrying the repair template and a fluorescent visual marker that facilitates identification of bona fide recombinant animals. Homozygous animals can be identified as early as 4–5 days post-injection, and minimal genotyping by PCR is required. We demonstrate that our toolkit of dual-marker vectors can generate targeted disruptions, deletions, and endogenous tagging with fluorescent proteins and epitopes. This strategy should be useful for a wide variety of additional applications and will provide researchers with increased flexibility when designing genome editing experiments.     

Heritable Custom Genomic Modifications in Caenorhabditis elegans via a CRISPR–Cas9 System

We adapted the CRISPR–Cas9 system for template-mediated repair of targeted double-strand breaks via homologous recombination in Caenorhabditis elegans, enabling customized and efficient genome editing. This system can be used to create specific insertions, deletions, and base pair changes in the germline of C. elegans.     

Towards systems metabolic engineering in Pichia pastoris

The methylotrophic yeast Pichia pastoris is firmly established as a host for the production of recombinant proteins, frequently outperforming other heterologous hosts. Already, a sizeable amount of systems biology knowledge has been acquired for this non-conventional yeast. By applying various omics-technologies, productivity features have been thoroughly analyzed and optimized via genetic engineering. However, challenging clonal variability, limited vector repertoire and insufficient genome annotation have hampered further developments. Yet, in the last few years a reinvigorated effort to establish P. pastoris as a host for both protein and metabolite production is visible. A variety of compounds from terpenoids to polyketides have been synthesized, often exceeding the productivity of other microbial systems. The clonal variability was systematically investigated and strategies formulated to circumvent untargeted events, thereby streamlining the screening procedure. Promoters with novel regulatory properties were discovered or engineered from existing ones. The genetic tractability was increased via the transfer of popular manipulation and assembly techniques, as well as the creation of new ones. A second generation of sequencing projects culminated in the creation of the second best functionally annotated yeast genome. In combination with landmark physiological insights and increased output of omics-data, a good basis for the creation of refined genome-scale metabolic models was created. The first application of model-based metabolic engineering in P. pastoris showcased the potential of this approach. Recent efforts to establish yeast peroxisomes for compartmentalized metabolite synthesis appear to fit ideally with the well-studied high capacity peroxisomal machinery of P. pastoris. Here, these recent developments are collected and reviewed with the aim of supporting the establishment of systems metabolic engineering in P. pastoris.     

Engineering Corynebacterium glutamicum with a comprehensive genomic library and phage-based vectors

The Gram-positive bacterium Corynebacterium glutamicum sustains the industrial production of chiral molecules such as L-amino acids. Through heterologous gene expression, C. glutamicum is becoming a sustainable source of small organic molecules and added-value chemicals. The current methods to implement heterologous genes in C. glutamicum rely on replicative vectors requiring lasting selection or chromosomal integration using homologous recombination. Here, we present a set of dedicated and transversal tools for genome editing and gene delivery into C. glutamicum. We generated a cosmid-based library suitable for efficient double allelic exchange, covering more than 94% of the chromosome with an average 5.1x coverage. We employed the library and an iterative marker excision system to generate the carotenoid-free C. glutamicum BT1-C31-Albino (BCA) host, featuring the attachment sites for actinophages ϕC31 and ϕBT1 for one-step chromosomal integration. As a proof-of-principle, we employed a ϕC31-based integration and a Cre system for the markerless expression of the type III polyketide synthase RppA, and a ϕBT1-based integration system for the expression of the phosphopantetheinylation-dependent non-ribosomal peptide synthetase BpsA in the C. glutamicum BCA host. The developed genomic library and microbial host, and the characterized molecular tools will contribute to the study of the physiology and the rise of C. glutamicum as a leading host for drug discovery.     

Streamlined CRISPR genome engineering in wild-type bacteria using SIBR-Cas

CRISPR-Cas is a powerful tool for genome editing in bacteria. However, its efficacy is dependent on host factors (such as DNA repair pathways) and/or exogenous expression of recombinases. In this study, we mitigated these constraints by developing a simple and widely applicable genome engineering tool for bacteria which we termed SIBR-Cas (Self-splicing Intron-Based Riboswitch-Cas). SIBR-Cas was generated from a mutant library of the theophylline-dependent self-splicing T4 td intron that allows for tight and inducible control over CRISPR-Cas counter-selection. This control delays CRISPR-Cas counter-selection, granting more time for the editing event (e.g. by homologous recombination) to occur. Without the use of exogenous recombinases, SIBR-Cas was successfully applied to knock-out several genes in three wild-type bacteria species (Escherichia coli MG1655, Pseudomonas putida KT2440 and Flavobacterium IR1) with poor homologous recombination systems. Compared to other genome engineering tools, SIBR-Cas is simple, tightly regulated and widely applicable for most (non-model) bacteria. Furthermore, we propose that SIBR can have a wider application as a simple gene expression and gene regulation control mechanism for any gene or RNA of interest in bacteria.     

High efficiency, site-specific excision of a marker gene by the phage P1 cre–loxP system in the yellow fever mosquito, Aedes aegypti

The excision of specific DNA sequences from integrated transgenes in insects permits the dissection in situ of structural elements that may be important in controlling gene expression. Furthermore, manipulation of potential control elements in the context of a single integration site mitigates against insertion site influences of the surrounding genome. The cre–loxP site-specific recombination system has been used successfully to remove a marker gene from transgenic yellow fever mosquitoes, Aedes aegypti. A total of 33.3% of all fertile families resulting from excision protocols showed evidence of cre–loxP-mediated site-specific excision. Excision frequencies were as high as 99.4% within individual families. The cre recombinase was shown to precisely recognize loxP sites in the mosquito genome and catalyze excision. Similar experiments with the FLP/FRT site-specific recombination system failed to demonstrate excision of the marker gene from the mosquito chromosomes.     

Pre-selection of integration sites imparts repeatable transgene expression

Variable gene expression amongst transgenic lines occurs due to copy number and to random associations of incoming DNA with chromosomal elements at the site of integration. Here we describe a method of identifying sites permissive for transgene expression and their use for efficient introduction of single copy transgenes by homologous recombination. ES clones were selected in HAT medium for expression of a randomly integrated HPRT marker lying 5′ to an Oct4/lacZ transgene. 794 clones were assessed in vitro for appropriate down-regulation of lacZ following differentiation. Two clones were chosen for further analysis which displayed appropriate and inappropriate gene regulation (clones 710 and 91, respectively). Three developmental promoters (thyroglobulin, Hox2.6 and Myf5) were then sequentially introduced into the original insertion sites in each clone (710 and 91) by homologous recombination, to drive expression of lacZ. Transgenic embryos were assessed for their ability to direct lacZ expression to tissues in which the respective promoter sequences are normally active. The site which appropriately down-regulated lacZ in vitro (710) also showed appropriate in vivo regulation of lacZ from the three developmental promoters. Site 91, however, directed an additional pattern of ectopic expression, which was common to all four promoters. Pre-selection of genomic sites for the introduction of transgenes by gene targeting improves the repeatability of transgene expression and provides an efficient means of single copy transgene introduction by homologous recombination.