CRISPR-Cas9 mediated gene deletions in lager yeast <Emphasis Type="Italic">Saccharomyces pastorianus</Emphasis>

Background

The ease of use of CRISPR-Cas9 reprogramming, its high efficacy, and its multiplexing capabilities have brought this technology at the forefront of genome editing techniques. Saccharomyces pastorianus is an aneuploid interspecific hybrid of Saccharomyces cerevisiae and Saccharomyces eubayanus that has been domesticated for centuries and is used for the industrial fermentation of lager beer. For yet uncharacterised reasons, this hybrid yeast is far more resilient to genetic alteration than its ancestor S. cerevisiae.

Results

This study reports a new CRISPR-Cas9 method for accurate gene deletion in S. pastorianus. This method combined the Streptococcus pyogenes cas9 gene expressed from either a chromosomal locus or from a mobile genetic element in combination with a plasmid-borne gRNA expression cassette. While the well-established gRNA expression system using the RNA polymerase III dependent SNR52 promoter failed, expression of a gRNA flanked with Hammerhead and Hepatitis Delta Virus ribozymes using the RNA polymerase II dependent TDH3 promoter successfully led to accurate deletion of all four alleles of the SeILV6 gene in strain CBS1483. Furthermore the expression of two ribozyme-flanked gRNAs separated by a 10-bp linker in a polycistronic array successfully led to the simultaneous deletion of SeATF1 and SeATF2, genes located on two separate chromosomes. The expression of this array resulted in the precise deletion of all five and four alleles mediated by homologous recombination in the strains CBS1483 and Weihenstephan 34/70 respectively, demonstrating the multiplexing abilities of this gRNA expression design.

Conclusions

These results firmly established that CRISPR-Cas9 significantly facilitates and accelerates genome editing in S. pastorianus.

     

Development of a genome editing technique using the CRISPR/Cas9 system in the industrial filamentous fungus <Emphasis Type="Italic">Aspergillus oryzae</Emphasis>

Objectives

To develop a genome editing method using the CRISPR/Cas9 system in Aspergillus oryzae, the industrial filamentous fungus used in Japanese traditional fermentation and for the production of enzymes and heterologous proteins.

Results

To develop the CRISPR/Cas9 system as a genome editing technique for A. oryzae, we constructed plasmids expressing the gene encoding Cas9 nuclease and single guide RNAs for the mutagenesis of target genes. We introduced these into an A. oryzae strain and obtained transformants containing mutations within each target gene that exhibited expected phenotypes. The mutational rates ranged from 10 to 20 %, and 1 bp deletions or insertions were the most commonly induced mutations.

Conclusions

We developed a functional and versatile genome editing method using the CRISPR/Cas9 system in A. oryzae. This technique will contribute to the use of efficient targeted mutagenesis in many A. oryzae industrial strains.

     

Design of a CRISPR-Cas system to increase resistance of <Emphasis Type="Italic">Bacillus subtilis</Emphasis> to bacteriophage SPP1

Clustered regularly interspaced short palindromic repeats (CRISPR) together with CRISPR-associated (cas) genes form an adaptive prokaryotic immune system which provides acquired resistance against viruses and plasmids. Bacillus subtilis presently is the best-characterized laboratory model for Gram-positive bacteria and also widely used for industrial production of enzymes, vitamins and antibiotics. In this study, we show that type II-A CRISPR-Cas system from Streptococcus thermophilus can be transferred into B. subtilis and provides heterologous protection against phage infection. We engineered a heterologous host by cloning S. thermophilus Cas9 and a spacer targeting bacteriophage SPP1 into the chromosome of B. subtilis, which does not harbor its own CRISPR-Cas systems. We found that the heterologous CRISPR-Cas system is functionally active in B. subtilis and provides resistance against bacteriophage SPP1 infection. The high efficiency of the acquired immunity against phage could be useful in generation of biotechnologically important B. subtilis strains with engineered chromosomes.     

Cpf1-assisted efficient genomic integration of in vivo assembled DNA parts in <Emphasis Type="Italic">Saccharomyces cerevisiae</Emphasis>

Objectives

To test the applicability of Cpf1 from Francisella novivida in genomic integration of in vivo assembled DNA parts in Saccharomyces cerevisiae.

Results

An easy-to-use vector toolkit, containing a CEN6/ARS4 plasmid expressing Cpf1 from Francisella novivida (FnCpf1) and a 2 μ plasmid for crRNA or crRNA array expressing, was constructed for Cpf1-assisted genomic integration in S. cerevisiae. Our results showed that FnCpf1 allowed for targeted singleplex, doubleplex, and tripleplex genomic integration of in vivo assembled DNA parts with efficiencies of 95, 52, and 43%, respectively.

Conclusions

CRISPR-Cpf1 system allows for efficient genomic integration of in vivo assembled DNA parts in S. cerevisiae, and thus provides an alternative CRISPR-Cas method for metabolic pathway engineering in addition to CRISPR-Cas9 system previously reported for yeast.

     

Knockout of <Emphasis Type="Italic">CTNNB1</Emphasis> by CRISPR-Cas9 technology inhibits cell proliferation through the Wnt/β-catenin signaling pathway

Objective

To study the effects of CTNNB1 gene knockout by CRISPR-Cas9 technology on cell adhesion, proliferation, apoptosis, and Wnt/β-catenin signaling pathway.

Results

CTNNB1 gene of HEK 293T cells was knocked out by CRISPR-Cas9. This was confirmed by sequencing and western blotting. Methylthiazolyl-tetrazolium bromide assays indicated that deletion of β-catenin significantly weakened adhesion ability and inhibited proliferation rate (P < 0.01) of HEK 293T cells. Nevertheless, deletion of β-catenin did not affect apoptosis of HEK 293T cells, which was analyzed by flow cytometry with Annexin V-fluorescein isothiocyanate/propidium iodide double staining. In addition, expression level of GSK-3β, CCND1, and CCNE1 detected by qPCR and expression level of N-Cadherin and cyclin D1 detected by western blotting were significantly decreased (P < 0.01) while expression of γ-catenin detected by western blotting was significantly increased (P < 0.001).

Conclusions

Knockout of CTNNB1 disturbed Wnt/β-catenin signaling pathway and significantly inhibited adhesion and proliferation of HEK 293T cells.

     

Multiplex gene editing of the <Emphasis Type="Italic">Yarrowia lipolytica</Emphasis> genome using the CRISPR-Cas9 system

Yarrowia lipolytica is categorized as a generally recognized as safe (GRAS) organism and is a heavily documented, unconventional yeast that has been widely incorporated into multiple industrial fields to produce valuable biochemicals. This study describes the construction of a CRISPR-Cas9 system for genome editing in Y. lipolytica using a single plasmid (pCAS1yl or pCAS2yl) to transport Cas9 and relevant guide RNA expression cassettes, with or without donor DNA, to target genes. Two Cas9 target genes, TRP1 and PEX10, were repaired by non-homologous end-joining (NHEJ) or homologous recombination, with maximal efficiencies in Y. lipolytica of 85.6 % for the wild-type strain and 94.1 % for the ku70/ku80 double-deficient strain, within 4 days. Simultaneous double and triple multigene editing was achieved with pCAS1yl by NHEJ, with efficiencies of 36.7 or 19.3 %, respectively, and the pCASyl system was successfully expanded to different Y. lipolytica breeding strains. This timesaving method will enable and improve synthetic biology, metabolic engineering and functional genomic studies of Y. lipolytica.     

Genetic tools for reliable gene expression and recombineering in <Emphasis Type="Italic">Pseudomonas putida</Emphasis>

Pseudomonas putida is a promising bacterial host for producing natural products, such as polyketides and nonribosomal peptides. In these types of projects, researchers need a genetic toolbox consisting of plasmids, characterized promoters, and techniques for rapidly editing the genome. Past reports described constitutive promoter libraries, a suite of broad host range plasmids that replicate in P. putida, and genome-editing methods. To augment those tools, we have characterized a set of inducible promoters and discovered that IPTG-inducible promoter systems have poor dynamic range due to overexpression of the LacI repressor. By replacing the promoter driving lacI expression with weaker promoters, we increased the fold induction of an IPTG-inducible promoter in P. putida KT2440 to 80-fold. Upon discovering that gene expression from a plasmid was unpredictable when using a high-copy mutant of the BBR1 origin, we determined the copy numbers of several broad host range origins and found that plasmid copy numbers are significantly higher in P. putida KT2440 than in the synthetic biology workhorse, Escherichia coli. Lastly, we developed a λRed/Cas9 recombineering method in P. putida KT2440 using the genetic tools that we characterized. This method enabled the creation of scarless mutations without the need for performing classic two-step integration and marker removal protocols that depend on selection and counterselection genes. With the method, we generated four scarless deletions, three of which we were unable to create using a previously established genome-editing technique.     

Gene editing and mutagenesis reveal inter-cultivar differences and additivity in the contribution of <Emphasis Type="Italic">TaGW2</Emphasis> homoeologues to grain size and weight in wheat

Key message

CRISPR-Cas9-based genome editing and EMS mutagenesis revealed inter-cultivar differences and additivity in the contribution of TaGW2 homoeologues to grain size and weight in wheat.

Abstract

The TaGW2 gene homoeologues have been reported to be negative regulators of grain size (GS) and thousand grain weight (TGW) in wheat. However, the contribution of each homoeologue to trait variation among different wheat cultivars is not well documented. We used the CRISPR-Cas9 system and TILLING to mutagenize each homoeologous gene copy in cultivars Bobwhite and Paragon, respectively. Plants carrying single-copy nonsense mutations in different genomes showed different levels of GS/TGW increase, with TGW increasing by an average of 5.5% (edited lines) and 5.3% (TILLING mutants). In any combination, the double homoeologue mutants showed higher phenotypic effects than the respective single-genome mutants. The double mutants had on average 12.1% (edited) and 10.5% (TILLING) higher TGW with respect to wild-type lines. The highest increase in GS and TGW was shown for triple mutants of both cultivars, with increases in 16.3% (edited) and 20.7% (TILLING) in TGW. The additive effects of the TaGW2 homoeologues were also demonstrated by the negative correlation between the functional gene copy number and GS/TGW in Bobwhite mutants and an F₂ population. The highest single-genome increases in GS and TGW in Paragon and Bobwhite were obtained by mutations in the B and D genomes, respectively. These inter-cultivar differences in the phenotypic effects between the TaGW2 gene homoeologues coincide with inter-cultivar differences in the homoeologue expression levels. These results indicate that GS/TGW variation in wheat can be modulated by the dosage of homoeologous genes with inter-cultivar differences in the magnitude of the individual homoeologue effects.

     

Concurrent modifications in the three homeologs of <Emphasis Type="Italic">Ms45</Emphasis> gene with CRISPR-Cas9 lead to rapid generation of male sterile bread wheat (<Emphasis Type="Italic">Triticum aestivum</Emphasis> L.)

Key message

Hexaploid bread wheat is not readily amenable to traditional mutagenesis approaches. In this study, we show efficient utilization of CRISPR-Cas system and Next Generation Sequencing for mutant analysis in wheat.

Abstract

Identification and manipulation of male fertility genes in hexaploid bread wheat is important for understanding the molecular basis of pollen development and to obtain novel sources of nuclear genetic male sterility (NGMS). The maize Male sterile 45 (Ms45) gene encodes a strictosidine synthase-like enzyme and has been shown to be required for male fertility. To investigate the role of Ms45 gene in wheat, mutations in the A, B and D homeologs were produced using CRISPR-Cas9. A variety of mutations in the three homeologs were recovered, including a plant from two different genotypes each with mutations in all three homeologs. Genetic analysis of the mutations demonstrated that all three wheat Ms45 homeologs contribute to male fertility and that triple homozygous mutants are required to abort pollen development and achieve male sterility. Further, it was demonstrated that a wild-type copy of Ms45 gene from rice was able to restore fertility to these wheat mutant plants. Taken together, these observations provide insights into the conservation of MS45 function in a polyploid species. Ms45 based NGMS can be potentially utilized for a Seed Production Technology (SPT)-like hybrid seed production system in wheat.

     

On the global CRISPR array behavior in class I systems

Background

Much effort is underway to build and upgrade databases and tools related to occurrence, diversity, and characterization of CRISPR-Cas systems. As microbial communities and their genome complements are unearthed, much emphasis has been placed on details of individual strains and model systems within the CRISPR-Cas classification, and that collection of information as a whole affords the opportunity to analyze CRISPR-Cas systems from a quantitative perspective to gain insight into distribution of CRISPR array sizes across the different classes, types and subtypes. CRISPR diversity, nomenclature, occurrence, and biological functions have generated a plethora of data that created a need to understand the size and distribution of these various systems to appreciate their features and complexity.

Results

By utilizing a statistical framework and visual analytic techniques, we have been able to test several hypotheses about CRISPR loci in bacterial class I systems. Quantitatively, though CRISPR loci can expand to hundreds of spacers, the mean and median sizes are 40 and 25, respectively, reflecting rather modest acquisition and/or retention overall. Histograms uncovered that CRISPR array size displayed a parametric distribution, which was confirmed by a goodness-of fit test. Mapping the frequency of CRISPR loci on a standardized chromosome plot revealed that CRISPRs have a higher probability of occurring at clustered locations along the positive or negative strand. Lastly, when multiple arrays occur in a particular system, the size of a particular CRISPR array varies with its distance from the cas operon, reflecting acquisition and expansion biases.

Conclusions

This study establishes that bacterial Class I CRISPR array size tends to follow a geometric distribution; these CRISPRs are not randomly distributed along the chromosome; and the CRISPR array closest to the cas genes is typically larger than loci in trans. Overall, we provide an analytical framework to understand the features and behavior of CRISPR-Cas systems through a quantitative lens.

Reviewers

This article was reviewed by Eugene Koonin (NIH-NCBI) and Uri Gophna (Tel Aviv University).

     

Recent advances in CRISPR/Cas9 mediated genome editing in <Emphasis Type="Italic">Bacillus subtilis</Emphasis>

Genome editing using engineered nucleases has rapidly transformed from a niche technology to a mainstream method used in various host cells. Its widespread adoption has been largely developed by the emergence of the clustered regularly interspaced short palindromic repeats (CRISPR) system, which uses an easily customizable specificity RNA-guided DNA endonuclease, such as Cas9. Recently, CRISPR/Cas9 mediated genome engineering has been widely applied to model organisms, including Bacillus subtilis, enabling facile, rapid high-fidelity modification of endogenous native genes. Here, we reviewed the recent progress in B. subtilis gene editing using CRISPR/Cas9 based tools, and highlighted state-of-the-art strategies for design of CRISPR/Cas9 system. Finally, future perspectives on the use of CRISPR/Cas9 genome engineering for sequence-specific genome editing in B. subtilis are provided.     

I-SceI endonuclease: a new tool for DNA repair studies and genetic manipulations in streptomycetes

Actinomycetes are Gram-positive bacteria with a complex life cycle. They produce many pharmaceutically relevant secondary metabolites, including antibiotics and anticancer drugs. However, there is a limited number of biotechnological applications available as opposed to genetic model organisms like Bacillus subtilis or Escherichia coli. We report here a system for the functional expression of a synthetic gene encoding the I-SceI homing endonuclease in several streptomycetes. Using the synthetic sce(a) gene, we were able to create controlled genomic DNA double-strand breaks. A mutagenesis system, based on the homing endonuclease I-SceI, has been developed to construct targeted, non-polar, unmarked gene mutations in Streptomyces sp. Tü6071. In addition, we have shown that homologous recombination is a major pathway in streptomycetes to repair an I-SceI-generated DNA double-strand break. This novel I-SceI-based tool will be useful in fundamental studies on the repair mechanism of DNA double-strand breaks and for a variety of biotechnological applications.     

Two acetyl-CoA synthetase isoenzymes are encoded by distinct genes in marine yeast <Emphasis Type="Italic">Rhodosporidium diobovatum</Emphasis>

Objectives

Two genes encoding two acetyl-CoA synthetase (ACS) isoenzymes have been identified in the marine yeast Rhodosporidium diobovatum MCCC 2A00023.

Results

ACS1 encoded a polypeptide with a sequence of 578 amino acid residues, a predicted molecular weight of 63.73 kDa, and pI of 8.14, while the ACS2 encoded a polypeptide containing 676 amino acid residues with a deduced molecular mass of 75.61 kDa and a pI of 5.95. Biological activity of Acs1p and Acs2p was confirmed by heterologous expression in Escherichia coli. A 1.5-kb DNA fragment of the ACS1 gene and a 2.7-kb DNA fragment of the ACS2 gene were deleted using the RNA guide CRISPR-Cas9 system. The strain lacking ACS1 was unable to grow on acetate and ethanol media, while the ACS2 deletant was unable to grow on glucose medium. ACS1-ACS2 double mutants of R. diobovatum were non-viable.

Conclusions

ACS isoenzymes are essential to the yeast metabolism, and other sources of ACSs cannot compensate for the lack of ACSs encoded by the two genes.

     

Targeted mutagenesis in the progeny of maize transgenic plants

We have demonstrated that targeted mutagenesis can be accomplished in maize plants by excision, activation, and subsequent elimination of an endonuclease in the progeny of genetic crosses. The yeast FLP/FRT site-specific recombination system was used to excise and transiently activate the previously integrated yeast I-SceI homing endonuclease in maize zygotes and/or developing embryos. An artificial I-SceI recognition sequence integrated into genomic DNA was analyzed for mutations to indicate the I-SceI endonuclease activity. Targeted mutagenesis of the I-SceI site occurred in about 1% of analyzed F1 plants. Short deletions centered on the I-SceI-produced double-strand break were the predominant genetic lesions observed in the F1 plants. The I-SceI expression cassette was not detected in the mutant F1 plants and their progeny. However, the original mutations were faithfully transmitted to the next generation indicating that the mutations occurred early during the F1 plant development. The procedure offers simultaneous production of double-strand breaks and delivery of DNA template combined with a large number of progeny plants for future gene targeting experiments.     

<Emphasis Type="Italic">Sfi</Emphasis>NX: a method for assembly of protein coding sequences with high success rates

Objective

Concatenation of two NdeI–XhoI gene fragments via an oligonucleotide linker on a plasmid vector with an SfiI site was performed to evaluate success rates in construction of polycistronic genes expressible in Escherichia coli.

Results

A series of plasmids with an SfiI site between the selection marker and the replication origin were constructed. The three wheat eEF1B subunit genes inserted between the NdeI and XhoI sites of pET-22b were transferred to the SfiI-containing plasmid with a spectinomycin-resistance gene. Then, the marker gene in the resultant plasmids was substituted with the ampicillin-resistance gene. These plasmids were used for concatenation of two different genes via a linker oligonucleotide containing a ribosome-binding site. During these operations, 42 clones were picked up out of which 41 had the intended product plasmid.

Conclusion

This method, named as the SfiNX method, is useful for trial-and-error based testing of different combinations of fusion and co-expression partners for optimization of recombinant protein production.

     

Site-directed mutagenesis in <Emphasis Type="Italic">Petunia</Emphasis> × <Emphasis Type="Italic">hybrida</Emphasis> protoplast system using direct delivery of purified recombinant Cas9 ribonucleoproteins

Key message

Site-directed mutagenesis of nitrate reductase genes using direct delivery of purified Cas9 protein preassembled with guide RNA produces mutations efficiently in Petunia × hybrida protoplast system.

Abstract

The clustered, regularly interspaced, short palindromic repeat (CRISPR)-CRISPR associated endonuclease 9 (CRISPR/Cas9) system has been recently announced as a powerful molecular breeding tool for site-directed mutagenesis in higher plants. Here, we report a site-directed mutagenesis method targeting Petunia nitrate reductase (NR) gene locus. This method could create mutations efficiently using direct delivery of purified Cas9 protein and single guide RNA (sgRNA) into protoplast cells. After transient introduction of RNA-guided endonuclease (RGEN) ribonucleoproteins (RNPs) with different sgRNAs targeting NR genes, mutagenesis at the targeted loci was detected by T7E1 assay and confirmed by targeted deep sequencing. T7E1 assay showed that RGEN RNPs induced site-specific mutations at frequencies ranging from 2.4 to 21 % at four different sites (NR1, 2, 4 and 6) in the PhNR gene locus with average mutation efficiency of 14.9 ± 2.2 %. Targeted deep DNA sequencing revealed mutation rates of 5.3–17.8 % with average mutation rate of 11.5 ± 2 % at the same NR gene target sites in DNA fragments of analyzed protoplast transfectants. Further analysis from targeted deep sequencing showed that the average ratio of deletion to insertion produced collectively by the four NR-RGEN target sites (NR1, 2, 4, and 6) was about 63:37. Our results demonstrated that direct delivery of RGEN RNPs into protoplast cells of Petunia can be exploited as an efficient tool for site-directed mutagenesis of genes or genome editing in plant systems.