A CRISPR-Cas9 Assisted Non-Homologous End-Joining Strategy for One-step Engineering of Bacterial Genome

Efficient, continuous mutagenesis in human cells using a pseudo-random DNA editor

Here we describe TRACE (T7 polymerase-driven continuous editing), a method that enables continuous, targeted mutagenesis in human cells using a cytidine deaminase fused to T7 RNA polymerase. TRACE induces high rates of mutagenesis over multiple cell generations in genes under the control of a T7 promoter integrated in the genome. We used TRACE in a MEK1 inhibitor-resistance screen, and identified functionally correlated mutations.

Strategies for Applying Nonhomologous End Joining-Mediated Genome Editing in Prokaryotes

The emergence of genome editing technology based on the CRISPR/Cas system enabled revolutionary progress in genetic engineering. Double-strand breaks (DSBs), which can be induced by the CRISPR/Cas9 system, cause serious DNA damage that can be repaired by a homologous recombination (HR) system or the nonhomologous end joining (NHEJ) pathway. However, many bacterial species have a very weak HR system. Thus, the NHEJ pathway can be used in prokaryotes. Starting with a brief introduction of the mechanism of the NHEJ pathway, this review focuses on current research and details of applications of NHEJ in eukaryotes, which forms the theoretical basis for the application of the NHEJ system in prokaryotes.

Establishment and application of a CRISPR-Cas12a assisted genome-editing system in Zymomonas mobilis

BACKGROUND

Efficient and convenient genome-editing toolkits can expedite genomic research and strain improvement for desirable phenotypes. Zymomonas mobilis is a highly efficient ethanol-producing bacterium with a small genome size and desirable industrial characteristics, which makes it a promising chassis for biorefinery and synthetic biology studies. While classical techniques for genetic manipulation are available for Z. mobilis, efficient genetic engineering toolkits enabling rapidly systematic and high-throughput genome editing in Z. mobilis are still lacking.

RESULTS

Using Cas12a (Cpf1) from Francisella novicida, a recombinant strain with inducible cas12a expression for genome editing was constructed in Z. mobilis ZM4, which can be used to mediate RNA-guided DNA cleavage at targeted genomic loci. gRNAs were then designed targeting the replicons of native plasmids of ZM4 with about 100% curing efficiency for three native plasmids. In addition, CRISPR-Cas12a recombineering was used to promote gene deletion and insertion in one step efficiently and precisely with efficiency up to 90%. Combined with single-stranded DNA (ssDNA), CRISPR-Cas12a system was also applied to introduce minor nucleotide modification precisely into the genome with high fidelity. Furthermore, the CRISPR-Cas12a system was employed to introduce a heterologous lactate dehydrogenase into Z. mobilis with a recombinant lactate-producing strain constructed.

CONCLUSIONS

This study applied CRISPR-Cas12a in Z. mobilis and established a genome editing tool for efficient and convenient genome engineering in Z. mobilis including plasmid curing, gene deletion and insertion, as well as nucleotide substitution, which can also be employed for metabolic engineering to help divert the carbon flux from ethanol production to other products such as lactate demonstrated in this work. The CRISPR-Cas12a system established in this study thus provides a versatile and powerful genome-editing tool in Z. mobilis for functional genomic research, strain improvement, as well as synthetic microbial chassis development for economic biochemical production.

CRISPR-Cas9-assisted native end-joining editing offers a simple strategy for efficient genetic engineering in Escherichia coli

Unlike eukaryotes, prokaryotes are less proficient in homologous recombination (HR) and non-homologous end-joining (NHEJ). All existing genomic editing methods for Escherichia coli (E. coli) rely on exogenous HR or NHEJ systems to repair DNA double-strand breaks (DSBs). Although an E. coli native end-joining (ENEJ) system has been reported, its potential in genetic engineering has not yet been explored. Here, we present a CRISPR-Cas9-assisted native end-joining editing and show that ENEJ-dependent DNA repair can be used to conduct rapid and efficient deletion of chromosome fragments up to 83 kb or gene inactivation. Moreover, the positive rate and editing efficiency are independent of high-efficiency competent cells. The method requires neither exogenous DNA repair systems nor introduced editing template. The Cas9-sgRNA complex is the only foreign element in this method. This study is the first successful engineering effort to utilize ENEJ mechanism in genomic editing and provides an effective strategy for genetic engineering in bacteria that are inefficient in HR and NHEJ.

Stress-inducible NHEJ in bacteria: function in DNA repair and acquisition of heterologous DNA

DNA double-strand breaks (DSB) in bacteria can be repaired by non-homologous end-joining (NHEJ), a two-component system relying on Ku and LigD. While performing a genetic characterization of NHEJ in Sinorhizobium meliloti, a representative of bacterial species encoding several Ku and LigD orthologues, we found that at least two distinct functional NHEJ repair pathways co-exist: one is dependent on Ku2 and LigD2, while the other depends on Ku3, Ku4 and LigD4. Whereas Ku2 likely acts as canonical bacterial Ku homodimers, genetic evidences suggest that Ku3-Ku4 form eukaryotic-like heterodimers. Strikingly, we found that the efficiency of both NHEJ systems increases under stress conditions, including heat and nutrient starvation. We found that this stimulation results from the transcriptional up-regulation of the ku and/or ligD genes, and that some of these genes are controlled by the general stress response regulator RpoE2. Finally, we provided evidence that NHEJ not only repairs DSBs, but can also capture heterologous DNA fragments into genomic breaks. Our data therefore suggest that NHEJ could participate to horizontal gene transfer from distantly related species, bypassing the need of homology to integrate exogenous DNA. This supports the hypothesis that NHEJ contributes to evolution and adaptation of bacteria under adverse environmental conditions.

CRISPR Genome Editing Systems in the Genus Clostridium: a Timely Advancement

The genus Clostridium is composed of bioproducers, which are important for the industrial production of chemicals, as well as pathogens, which are a significant burden to the patients and on the health care industry. Historically, even though these bacteria are well known and are commonly studied, the genetic technologies to advance our understanding of these microbes have lagged behind other systems. New tools would continue the advancement of our understanding of clostridial physiology. The genetic modification systems available in several clostridia are not as refined as in other organisms and each exhibit their own drawbacks. With the advent of the repurposing of the CRISPR-Cas systems for genetic modification, the tools available for clostridia have improved significantly over the past four years. Several CRISPR-Cas systems such as using wild-type Cas9, Cas9n, dCas9/CRISPR interference (CRISPRi) and a newly studied Cpf1/Cas12a, are reported. These have the potential to greatly advance the study of clostridial species leading to future therapies or the enhanced production of industrially relevant compounds. Here we discuss the details of the CRISPR-Cas systems as well as the advances and current issues in the developed clostridial systems.

The phage T4 DNA ligase in vivo improves the survival-coupled bacterial mutagenesis

BACKGROUND

Microbial mutagenesis is an important avenue to acquire microbial strains with desirable traits for industry application. However, mutagens either chemical or physical used often leads narrow library pool due to high lethal rate. The T4 DNA ligase is one of the most widely utilized enzymes in modern molecular biology. Its contribution to repair chromosomal DNA damages, therefore cell survival during mutagenesis will be discussed.

RESULTS

Expression of T4 DNA ligase in vivo could substantially increase cell survival to ionizing radiation in multiple species. A T4 mediated survival-coupled mutagenesis approach was proposed. When polyhydroxybutyrate (PHB)-producing E. coli with T4 DNA ligase expressed in vivo was subjected to ionizing radiation, mutants with improved PHB production were acquired quickly owing to a large viable mutant library generated. Draft genome sequence analysis showed that the mutants obtained possess not only single nucleotide variation (SNV) but also DNA fragment deletion, indicating that T4 DNA ligase in vivo may contribute to the repair of DNA double strand breaks.

CONCLUSIONS

Expression of T4 DNA ligase in vivo could notably enhance microbial survival to excess chromosomal damages caused by various mutagens. Potential application of T4 DNA ligase in microbial mutagenesis was explored by mutating and screening PHB producing E. coli XLPHB strain. When applied to atmospheric and room temperature plasma (ARTP) microbial mutagenesis, large survival pool was obtained. Mutants available for subsequent screening for desirable features. The use of T4 DNA ligase we were able to quickly improve the PHB production by generating a larger viable mutants pool. This method is a universal strategy can be employed in wide range of bacteria. It indicated that traditional random mutagenesis became more powerful in combine with modern genetic molecular biology and has exciting prospect.

The phage T4 DNA ligase mediates bacterial chromosome DSBs repair as single component non-homologous end joining

DNA double-strand breaks (DSBs) are one of the most lethal forms of DNA damage that is not efficiently repaired in prokaryotes. Certain microorganisms can handle chromosomal DSBs using the error-prone non-homologous end joining (NHEJ) system and ultimately cause genome mutagenesis. Here, we demonstrated that Enterobacteria phage T4 DNA ligase alone is capable of mediating in vivo chromosome DSBs repair in Escherichia coli. The ligation efficiency of DSBs with T4 DNA ligase is one order of magnitude higher than the NHEJ system from Mycobacterium tuberculosis. This process introduces chromosome DNA excision with different sizes, which can be manipulated by regulating the activity of host-exonuclease RecBCD. The DNA deletion length reduced either by inactivating recB or expressing the RecBCD inhibitor Gam protein from λ phage. Furthermore, we also found single nucleotide substitutions at the DNA junction, suggesting that T4 DNA ligase, as a single component non-homologous end joining system, has great potential in genome mutagenesis, genome reduction and genome editing.

Editing the microbiome the CRISPR way

Our bodies are colonized by a complex ecosystem of bacteria, unicellular eukaryotes and their viruses that together play a major role in our health. Over the past few years tools derived from the prokaryotic immune system known as CRISPR-Cas have empowered researchers to modify and study organisms with unprecedented ease and efficiency. Here we discuss how various types of CRISPR-Cas systems can be used to modify the genome of gut microorganisms and bacteriophages. CRISPR-Cas systems can also be delivered to bacterial population and programmed to specifically eliminate members of the microbiome. Finally, engineered CRISPR-Cas systems can be used to control gene expression and modulate the production of metabolites and proteins. Together these tools provide exciting opportunities to investigate the complex interplay between members of the microbiome and our bodies, and present new avenues for the development of drugs that target the microbiome. This article is part of a discussion meeting issue 'The ecology and evolution of prokaryotic CRISPR-Cas adaptive immune systems'.

Synthetic Genome Defenses against Selfish DNA Elements Stabilize Engineered Bacteria against Evolutionary Failure

Mobile genetic elements drive evolution by disrupting genes and rearranging genomes. Eukaryotes have evolved epigenetic mechanisms, including DNA methylation and RNA interference, that silence mobile elements and thereby preserve the integrity of their genomes. We created an artificial reprogrammable epigenetic system based on CRISPR interference to give engineered bacteria a similar line of defense against transposons and other selfish elements in their genomes. We demonstrate that this CRISPR interference against mobile elements (CRISPRi-ME) approach can be used to simultaneously repress two different transposon families in Escherichia coli, thereby increasing the evolutionary stability of costly protein expression. We further show that silencing a transposon in Acinetobacter baylyi ADP1 reduces mutation rates by a factor of 5, nearly as much as deleting all copies of this element from its genome. By deploying CRISPRi-ME on a broad-host-range vector, we have created a generalizable platform for stabilizing the genomes of engineered bacterial cells for applications in metabolic engineering and synthetic biology.

Development of a Bicistronic Vector for the Expression of a CRISPR/Cas9-mCherry System in Fish Cell Lines

The clustered regularly interspaced short palindromic repeats/CRISPR-associated protein 9 (CRISPR/Cas9) system has been widely used in animals as an efficient genome editing tool. In fish cells, the technique has been difficult to implement due to the lack of proper vectors that use active promoters to drive the expression of both small guide RNA (sgRNA) and the S. pyogenes Cas9 (spCas9) protein within a single expression platform. Until now, fish cells have been modified using co-transfection of the mRNA of both the sgRNA and the spCas9. In the present study, we describe the optimization of a new vector for the expression of a CRISPR/Cas9 system, designed to edit the genome of fish cell lines, that combines a gene reporter (mCherry), sgRNA, and spCas9 in a single vector, facilitating the study of the efficiency of piscine and non-piscine promoters. A cassette containing the zebrafish U6 RNA III polymerase (U6ZF) promoter was used for the expression of the sgRNA. The new plasmid displayed the expression of spCas9, mCherry, and sgRNA in CHSE/F fish cells. The results demonstrate the functionality of the mammalian promoter and the U6ZF promoter in fish cell lines. This is the first approach aimed at developing a unified genome editing system in fish cells using bicistronic vectors, thus creating a powerful biotechnological platform to study gene function.

Characterization and Repurposing of Type I and Type II CRISPR-Cas Systems in Bacteria

CRISPR-Cas systems constitute the adaptive immune system of bacteria and archaea, as a sequence-specific nucleic acid targeting defense mechanism. The sequence-specific recognition and cleavage of Cas effector complexes has been harnessed to developed CRISPR-based technologies and drive the genome editing revolution underway, due to their efficacy, efficiency, and ease of implementation in a broad range of organisms. CRISPR-based technologies offer a wide variety of opportunities in genome remodeling and transcriptional regulation, opening new avenues for therapeutic and biotechnological applications. To repurpose CRISPR-Cas systems for these applications, the various elements of the system need to be first identified and functionally characterized in their native host. Bioinformatic tools are first used to identify putative CRISPR arrays and their associated genes, followed by a comprehensive characterization of the CRISPR-Cas system, encompassing predictions for guide and target sequences. Subsequently, interference assays and transcriptomic analyses should be performed to probe the functionality of the CRISPR-Cas system. Once an endogenous CRISPR-Cas system is characterized as functional, they can be readily repurposed by delivering an engineered synthetic CRISPR array or a small RNA guide for targeted gene manipulation. Alternatively, developing a plasmid-based system for heterologous expression of the necessary CRISPR components can enable exploitation in other organisms. Altogether, there is a wide diversity of native CRISPR-Cas systems in many bacteria and most archaea that await functional characterization and repurposing for genome editing applications in prokaryotes.

Chemical transformation mediated CRISPR/Cas9 genome editing in Escherichia coli

OBJECTIVES

To develop a convenient chemical transformation mediated CRISPR/Cas9 (CT-CRISPR/Cas9) system for genome editing in Escherichia coli.

RESULTS

Here, we have constructed a CT-CRISPR/Cas9 system, which can precisely edit bacterial genome (replacing, deleting, inserting or point mutating a target gene) through chemical transformation. Compared with the traditional electroporation mediated CRISPR/Cas9 (ET-CRISPR/Cas9) system, genome editing with the CT-CRISPR/Cas9 system is much cheaper and simpler. In the CT-CRISPR/Cas9 system, we observed efficient genome editing on LB-agar plates. The CT-CRISPR/Cas9 system has successfully modified the target gene with the editing template flanked by short homologous DNA fragments (~ 50 bp) which were designed in primers. We used the lab-made CaCl₂ solution to perform the CT-CRISPR/Cas9 experiment and successfully edited the genome of E. coli. Potential application of the CT-CRISPR/Cas9 system in high-throughput genome editing was evaluated in two E. coli strains by using a multiwell plate.

CONCLUSIONS

Our work provides a simple and cheap genome-editing method, that is expected to be widely applied as a routine genetic engineering method.

Serine integrase recombinational engineering (SIRE): A versatile toolbox for genome editing

Chromosomal integration of biosynthetic pathways for the biotechnological production of high-value chemicals is a necessity to develop industrial strains with a high long-term stability and a low production variability. However, the introduction of multiple transcription units into the microbial genome remains a difficult task. Despite recent advances, current methodologies are either laborious or efficiencies highly fluctuate depending on the length and the type of the construct. Here we present serine integrase recombinational engineering (SIRE), a novel methodology which combines the ease of recombinase-mediated cassette exchange (RMCE) with the selectivity of orthogonal att sites of the PhiC31 integrase. As a proof of concept, this toolbox is developed for Escherichia coli. Using SIRE we were able to introduce a 10.3 kb biosynthetic gene cluster on different locations throughout the genome with an efficiency of 100% for the integrating step and without the need for selection markers on the knock-in cassette. Next to integrating large fragments, the option for multitargeting, for deleting operons, as well as for performing in vivo assemblies further expand and proof the versatility of the SIRE toolbox for E. coli. Finally, the serine integrase PhiC31 was also applied in the yeast Saccharomyces cerevisiae as a marker recovery tool, indicating the potential and portability of this toolbox.

CRISPR/Cpf1 facilitated large fragment deletion in Saccharomyces cerevisiae

In this study, we focused on the applicability of CRISPR/Cpf1 in genome simplification of Saccharomyces cerevisiae and established a CRISPR/Cpf1 assisted method for rapid markerless large fragment deletion to facilitate laboratory evolution of geome of S. cerevisiae by rational genetic engineering. This method uses a Cpf1 expression plasmid and a crRNA array expression plasmid. The DNA fragment between two DSBs generated by CRISPR/Cpf1 can be cut off from the chromosome, along with re-ligation of the genomic endpoints of the DSBs. Using this method, the large DNA fragment of ∼38 kb between the two genes of TRM10 and REX4 was successfully and rapidly deleted, which was verified by PCR and Sanger DNA Sequencing. This method is simple and rapid, and can be easily implemented for large fragment deletion in S. cerevisiae.

Targeted Nucleotide Editing Technologies for Microbial Metabolic Engineering

Since the emergence of programmable RNA-guided nucleases based on clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein (Cas) systems, genome editing technologies have become a simplified and versatile tool for genome editing in various organisms and cell types. Although genome editing enables efficient genome manipulations, such as gene disruptions, gene knockins, and chromosomal translocations via DNA double-strand break (DSB) repair in eukaryotes, DSBs induced by the CRISPR/Cas system are lethal or severely toxic to many microorganisms. Therefore, in many prokaryotes, including industrially useful microbes, the CRISPR/Cas system is often used as a negative selection component in combination with recombineering or other related strategies. Novel and revolutionary technologies have been recently developed to re-write targeted nucleotides (C:G to T:A and A:T to G:C) without DSBs and donor DNA templates. These technologies rely on the recruitment of deaminases at specific target loci using the nuclease-deficient CRISPR/Cas system. Here, the authors review and compare CRISPR-based genome editing, current base editing platforms and their spectra. The authors discuss how these technologies can be applied in various aspects of microbial metabolic engineering to overcome barriers to cellular regulation in prokaryotes.

High GC Content Cas9-Mediated Genome-Editing and Biosynthetic Gene Cluster Activation in Saccharopolyspora erythraea

The overexpression of bacterial secondary metabolite biosynthetic enzymes is the basis for industrial overproducing strains. Genome editing tools can be used to further improve gene expression and yield. Saccharopolyspora erythraea produces erythromycin, which has extensive clinical applications. In this study, the CRISPR-Cas9 system was used to edit genes in the S. erythraea genome. A temperature-sensitive plasmid containing the PermE promoter, to drive Cas9 expression, and the Pj23119 and PkasO promoters, to drive sgRNAs, was designed. Erythromycin esterase, encoded by S. erythraea SACE_1765, inactivates erythromycin by hydrolyzing the macrolactone ring. Sequencing and qRT-PCR confirmed that reporter genes were successfully inserted into the SACE_1765 gene. Deletion of SACE_1765 in a high-producing strain resulted in a 12.7% increase in erythromycin levels. Subsequent PermE- egfp knock-in at the SACE_0712 locus resulted in an 80.3% increase in erythromycin production compared with that of wild type. Further investigation showed that PermE promoter knock-in activated the erythromycin biosynthetic gene clusters at the SACE_0712 locus. Additionally, deletion of indA (SACE_1229) using dual sgRNA targeting without markers increased the editing efficiency to 65%. In summary, we have successfully applied Cas9-based genome editing to a bacterial strain, S. erythraea, with a high GC content. This system has potential application for both genome-editing and biosynthetic gene cluster activation in Actinobacteria.

A CRISPR-Cas9-Based Toolkit for Fast and Precise In Vivo Genetic Engineering of Bacillus subtilis Phages

Phages are currently under discussion as a solution for the antibiotic crisis, as they may cure diseases caused by multi-drug-resistant pathogens. However, knowledge of phage biology and genetics is limited, which impedes risk assessment of therapeutic applications. In order to enable advances in phage genetic research, the aim of this work was to create a toolkit for simple and fast genetic engineering of phages recruiting Bacillus subtilis as host system. The model organism B. subtilis represents a non-pathogenic surrogate of its harmful relatives, such as Bacillus anthracis or Bacillus cereus. This toolkit comprises the application CutSPR, a bioinformatic tool for rapid primer design, and facilitates the cloning of specific CRISPR-Cas9-based mutagenesis plasmids. The employment of the prophage-free and super-competent B. subtilis TS01 strain enables an easy and fast introduction of specific constructs for in vivo phage mutagenesis. Clean gene deletions and a functional clean gene insertion into the genome of the model phage vB_BsuP-Goe1 served as proof of concept and demonstrate reliability and high efficiency. The here presented toolkit allows comprehensive investigation of the diverse phage genetic pool, a better understanding of phage biology, and safe phage applications.

Development of an Efficient Genome Editing Tool in Bacillus licheniformis Using CRISPR-Cas9 Nickase

Bacillus strains are important industrial bacteria that can produce various biochemical products. However, low transformation efficiencies and a lack of effective genome editing tools have hindered its widespread application. Recently, clustered regularly interspaced short palindromic repeat (CRISPR)-Cas9 techniques have been utilized in many organisms as genome editing tools because of their high efficiency and easy manipulation. In this study, an efficient genome editing method was developed for Bacillus licheniformis using a CRISPR-Cas9 nickase integrated into the genome of B. licheniformis DW2 with overexpression driven by the P43 promoter. The yvmC gene was deleted using the CRISPR-Cas9n technique with homology arms of 1.0 kb as a representative example, and an efficiency of 100% was achieved. In addition, two genes were simultaneously disrupted with an efficiency of 11.6%, and the large DNA fragment bacABC (42.7 kb) was deleted with an efficiency of 79.0%. Furthermore, the heterologous reporter gene aprN, which codes for nattokinase in Bacillus subtilis, was inserted into the chromosome of B. licheniformis with an efficiency of 76.5%. The activity of nattokinase in the DWc9nΔ7/pP43SNT-SsacC strain reached 59.7 fibrinolytic units (FU)/ml, which was 25.7% higher than that of DWc9n/pP43SNT-SsacC Finally, the engineered strain DWc9nΔ7 (Δepr ΔwprA Δmpr ΔaprE Δvpr ΔbprA ΔbacABC), with multiple disrupted genes, was constructed using the CRISPR-Cas9n technique. Taken together, we have developed an efficient genome editing tool based on CRISPR-Cas9n in B. licheniformis This tool could be applied to strain improvement for future research.IMPORTANCE As important industrial bacteria, Bacillus strains have attracted significant attention due to their production of biological products. However, genetic manipulation of these bacteria is difficult. The CRISPR-Cas9 system has been applied to genome editing in some bacteria, and CRISPR-Cas9n was proven to be an efficient and precise tool in previous reports. The significance of our research is the development of an efficient, more precise, and systematic genome editing method for single-gene deletion, multiple-gene disruption, large DNA fragment deletion, and single-gene integration in Bacillus licheniformis via Cas9 nickase. We also applied this method to the genetic engineering of the host strain for protein expression.

Recent Developments of the Synthetic Biology Toolkit for Clostridium

The Clostridium genus is a large, diverse group consisting of Gram-positive, spore-forming, obligate anaerobic firmicutes. Among this group are historically notorious pathogens as well as several industrially relevant species with the ability to produce chemical commodities, particularly biofuels, from renewable biomass. Additionally, other species are studied for their potential use as therapeutics. Although metabolic engineering and synthetic biology have been instrumental in improving product tolerance, titer, yields, and feed stock consumption capabilities in several organisms, low transformation efficiencies and lack of synthetic biology tools and genetic parts make metabolic engineering within the Clostridium genus difficult. Progress has recently been made to overcome challenges associated with engineering various Clostridium spp. For example, developments in CRISPR tools in multiple species and strains allow greater capability to produce edits with greater precision, faster, and with higher efficiencies. In this mini-review, we will highlight these recent advances and compare them to established methods for genetic engineering in Clostridium. In addition, we discuss the current state and development of Clostridium-based promoters (constitutive and inducible) and reporters. Future progress in this area will enable more rapid development of strain engineering, which would allow for the industrial exploitation of Clostridium for several applications including bioproduction of several commodity products.

CRISPR-Cas9 Genetic Analysis of Virus-Host Interactions

Clustered regularly interspaced short palindromic repeats (CRISPR) has greatly expanded the ability to genetically probe virus–host interactions. CRISPR systems enable focused or systematic, genomewide studies of nearly all aspects of a virus lifecycle. Combined with its relative ease of use and high reproducibility, CRISPR is becoming an essential tool in studies of the host factors important for viral pathogenesis. Here, we review the use of CRISPR–Cas9 for the loss-of-function analysis of host dependency factors. We focus on the use of CRISPR-pooled screens for the systematic identification of host dependency factors, particularly in Epstein–Barr virus-transformed B cells. We also discuss the use of CRISPR interference (CRISPRi) and gain-of-function CRISPR activation (CRISPRa) approaches to probe virus–host interactions. Finally, we comment on the future directions enabled by combinatorial CRISPR screens.

Recent Developments of the Synthetic Biology Toolkit for Clostridium

The Clostridium genus is a large, diverse group consisting of Gram-positive, spore-forming, obligate anaerobic firmicutes. Among this group are historically notorious pathogens as well as several industrially relevant species with the ability to produce chemical commodities, particularly biofuels, from renewable biomass. Additionally, other species are studied for their potential use as therapeutics. Although metabolic engineering and synthetic biology have been instrumental in improving product tolerance, titer, yields, and feed stock consumption capabilities in several organisms, low transformation efficiencies and lack of synthetic biology tools and genetic parts make metabolic engineering within the Clostridium genus difficult. Progress has recently been made to overcome challenges associated with engineering various Clostridium spp. For example, developments in CRISPR tools in multiple species and strains allow greater capability to produce edits with greater precision, faster, and with higher efficiencies. In this mini-review, we will highlight these recent advances and compare them to established methods for genetic engineering in Clostridium. In addition, we discuss the current state and development of Clostridium-based promoters (constitutive and inducible) and reporters. Future progress in this area will enable more rapid development of strain engineering, which would allow for the industrial exploitation of Clostridium for several applications including bioproduction of several commodity products.

Genome editing of Ralstonia eutropha using an electroporation-based CRISPR-Cas9 technique

BACKGROUND

Ralstonia eutropha is an important bacterium for the study of polyhydroxyalkanoates (PHAs) synthesis and CO₂ fixation, which makes it a potential strain for industrial PHA production and attractive host for CO₂ conversion. Although the bacterium is not recalcitrant to genetic manipulation, current methods for genome editing based on group II introns or single crossover integration of a suicide plasmid are inefficient and time-consuming, which limits the genetic engineering of this organism. Thus, developing an efficient and convenient method for R. eutropha genome editing is imperative.

RESULTS

An efficient genome editing method for R. eutropha was developed using an electroporation-based CRISPR-Cas9 technique. In our study, the electroporation efficiency of R. eutropha was found to be limited by its restriction-modification (RM) systems. By searching the putative RM systems in R. eutropha H16 using REBASE database and comparing with that in E. coli MG1655, five putative restriction endonuclease genes which are related to the RM systems in R. eutropha were predicated and disrupted. It was found that deletion of H16_A0006 and H16_A0008-9 increased the electroporation efficiency 1658 and 4 times, respectively. Fructose was found to reduce the leaky expression of the arabinose-inducible pBAD promoter, which was used to optimize the expression of cas9, enabling genome editing via homologous recombination based on CRISPR-Cas9 in R. eutropha. A total of five genes were edited with efficiencies ranging from 78.3 to 100%. The CRISPR-Cpf1 system and the non-homologous end joining mechanism were also investigated, but failed to yield edited strains.

CONCLUSIONS

We present the first genome editing method for R. eutropha using an electroporation-based CRISPR-Cas9 approach, which significantly increased the efficiency and decreased time to manipulate this facultative chemolithoautotrophic microbe. The novel technique will facilitate more advanced researches and applications of R. eutropha for PHA production and CO₂ conversion.

Inhibition of NHEJ repair by type II-A CRISPR-Cas systems in bacteria

Type II CRISPR-Cas systems introduce double-strand breaks into DNA of invading genetic material and use DNA fragments to acquire novel spacers during adaptation. These breaks can be the substrate of several DNA repair pathways, paving the way for interactions. We report that non-homologous end-joining (NHEJ) and type II-A CRISPR-Cas systems only co-occur once among 5563 fully sequenced prokaryotic genomes. We investigated experimentally the possible molecular interactions using the NHEJ pathway from Bacillus subtilis and the type II-A CRISPR-Cas systems from Streptococcus thermophilus and Streptococcus pyogenes. Our results suggest that the NHEJ system has no effect on CRISPR immunity. On the other hand, we provide evidence for the inhibition of NHEJ repair by the Csn2 protein. Our findings give insights on the complex interactions between CRISPR-Cas systems and repair mechanisms in bacteria, contributing to explain the scattered distribution of CRISPR-Cas systems in bacterial genome.

CRISPR/Cas9-Based Counterselection Boosts Recombineering Efficiency in Pseudomonas putida

While adoption of single-stranded DNA recombineering techniques has greatly eased genetic design of the platform strain Pseudomonas putida KT2440, available methods still produce the desired modifications/deletions at low frequencies. This makes isolation of mutants that do not display selectable or conspicuous phenotypes considerably difficult. To overcome this limitation, the authors have merged ssDNA recombineering with CRISPR/Cas9 technology in this bacterium for efficient killing of unmodified cells and thus non-phenotypic selection of bacteria bearing the mutations of interest. After incorporating the system into standardized pSEVA plasmids the authors tested its functional efficiency by targeting different types of changes that ranged from single nucleotide substitutions to one-gene deletions-to even the removal of the large flagellar cluster (≈69 kb). Simultaneous introduction of two independent gene deletions was tested as well. In all cases, directing the crRNA/Cas9 complexes toward non-modified, wild-type genomic sequences boosted dramatically the appearance of the mutants at stake in the absence of any phenotypic selection. The results presented here upgrade the engineering possibilities of the genome of this environmental bacterium (and possibly other Gram-negatives) to obtain modifications that are otherwise cumbersome to generate.

An efficient system for deletion of large DNA fragments in Escherichia coli via introduction of both Cas9 and the non-homologous end joining system from Mycobacterium smegmatis

Accompanied with the internal non-homologous end joining (NHEJ) system, Cas9 can be used to easily inactivate a gene or delete a fragment through introduction of DNA double-stranded breaks (DSBs) in eukaryotic cells. While in most prokaryotes (e.g. Escherichia coli), due to the lack of NHEJ, homologous recombination (HR) is required for repair of DSBs, which is less convenient. Here, a markerless system was developed for rapid gene inactivation or fragment deletion in E. coli via introduction of both Cas9 and a bacterial NHEJ system. Three bacterial NHEJ systems, i.e. Mycobacterium smegmatis (Msm), Mycobacterium tuberculosis (Mtb) and Bacillus subtilis (Bs), were tested in E. coli, and the MsmNHEJ system showed the best efficiency. With the employment of Cas9 and MsmNHEJ, we efficiently mutated lacZ gene, deleted glnALG operon and two large DNA fragments (67 kb and 123 kb) in E. coli, respectively. Moreover, the system was further designed to allow for continuous inactivation of genes or deletion of DNA fragments in E. coli. We envision this system can be extended to other bacteria, especially those with low HR efficiency.