Rapid, Efficient, and Cost-Effective Gene Editing of Enterococcus faecium with CRISPR-Cas12a.

Considered a serious threat by the Centers for Disease Control and Prevention, multidrug-resistant Enterococcus faecium is an increasing cause of hospital-acquired infection. Here, we provide details on a single-plasmid CRISPR-Cas12a system for generating clean deletions and insertions. Single manipulations were carried out in under 2 weeks, with successful deletions/insertions present in >80% of the clones tested. Using this method, we generated three individual clean deletion mutations in the acpH, treA, and lacL genes and inserted codon-optimized unaG, enabling green fluorescent protein (GFP)-like fluorescence under the control of the trehalase operon. The use of in vivo recombination for plasmid construction kept costs to a minimum. IMPORTANCE Enterococcus faecium is increasingly associated with hard-to-treat antibiotic-resistant infections. The ability to generate clean genomic alterations is the first step in generating a complete mechanistic understanding of how E. faecium acquires pathogenic traits and causes disease. Here, we show that CRISPR-Cas12a can be used to quickly (under 2 weeks) and cheaply delete or insert genes into the E. faecium genome. This substantial improvement over current methods should speed up research on this important opportunistic pathogen.

Rapid generation of conditional knockout mice using the CRISPR-Cas9 system and electroporation for neuroscience research.

The Cre/LoxP-based conditional knockout technology is a powerful tool for gene function analysis that allows region- and time-specific gene manipulation. However, inserting a pair of LoxP cassettes to generate conditional knockout can be technically challenging and thus time- and resource-consuming. This study proposes an efficient, low-cost method to generate floxed mice using in vitro fertilization and the CRISPR-Cas9 system over two consecutive generations. This method allowed us to produce floxed mice targeting exons 5 and 6 of CaMK1 in a short period of 125 days, using only 16 mice. In addition, we directly edited the genome of fertilized eggs of mice with our target genetic background, C57BL/6 N, to eliminate additional backcrossing steps. We confirmed that the genome of the generated floxed mice was responsive to the Cre protein. This low-cost, time-saving method for generating conditional knockout will facilitate comprehensive, tissue-specific genome analyses.

Efficient and Economical Targeted Insertion in Plant Genomes via Protoplast Regeneration.

Versatile genome editing can be facilitated by the insertion of DNA sequences into specific locations. Current protocols involving CRISPR and Cas proteins rely on low efficiency homology-directed repair or non-homologous end joining with modified double-stranded DNA oligonucleotides as donors. Our simple protocol eliminates the need for expensive equipment, chemical and enzymatic donor DNA modification, or plasmid construction by using polyethylene glycol-calcium to deliver non-modified single-stranded DNA oligonucleotides and CRISPR-Cas9 ribonucleoprotein into protoplasts. Plants regenerated via edited protoplasts achieved targeted insertion frequencies of up to 50% in Nicotiana benthamiana and 13.6% in rapid cycling Brassica oleracea without antibiotic selection. Using a 60 nt donor containing 27 nt in each homologous arm, 6/22 regenerated N. benthamiana plants showed targeted insertions, and one contained a precise insertion of a 6 bp HindIII site. The inserted sequences were transmitted to the next generation and invite the possibility of future exploration of versatile genome editing by targeted DNA insertion in plants.

Rapid and cost-effective screening of CRISPR/Cas9-induced mutants by DNA-guided Argonaute nuclease.

OBJECTIVE: With the widespread application of CRISPR/Cas9 gene editing technology, new methods are needed to screen mutants quickly and effectively. Here, we aimed to develop a simple and cost-effective method to screen CRISPR/Cas9-induced mutants. RESULT: We report a novel method to identify CRISPR/Cas9-induced mutants through a DNA-guided Argonaute nuclease derived from the archaeon Pyrococcus furiosus. We demonstrated that the Pyrococcus furiosus Argonaute (PfAgo)-based method could distinguish among biallelic mutants, monoallelic mutants and wild type (WT). Furthermore, this method was able to identify 1 bp indel mutations. CONCLUSION: The PfAgo-based method is simple to implement and can be applied to screen biallelic mutants and mosaic mutants generated by CRISPR-Cas9 or other kinds of gene editing tools.

Moving somatic gene editing to the clinic: routes to market access and reimbursement in Europe.

Somatic gene editing (SGE) holds great promise for making genetic therapy possible for many monogenic conditions very soon. Is our current system of European market authorization and reimbursement ready for the expected tsunami of gene therapies? At a recent workshop of the Netherlands ZonMw consortium on ethical, legal, and social implications of personalized medicine, we discussed the current possibilities for bringing new gene therapies to the clinic. In Europe, it is not yet clear whether the route via the European medicines agency as an advanced therapy medicinal product is the most appropriate for evaluation of highly personalized SGE applications, although this may optimally guarantee safety and effectiveness. Compassionate use may ensure faster access than the centralized procedure but does not stimulate the commercial development of products. Prescription to named patients may only provide adequate access for single patients. Temporary authorization of use may allow access to medication half a year before formal market authorization has been granted, but may also have large budget impacts. Magistral compounding under a hospital exemption may be an attractive solution for rare, tailor-made applications at an acceptable price. To approve local experimental use of a therapy on a case-by-case basis may be fast, but does not guarantee optimal safety, effectiveness, and broad implementation. We argue that alternative routes should be considered for products developed for a market of large groups of patients versus unique personalized treatments. A balance between scientific evidence for safety and effectiveness, affordability, and fast access may demand a range of alternative solutions.

CRISPR/Cas9 facilitates rapid generation of constitutive forms of transcription factors in Aspergillus niger through specific on-site genomic mutations resulting in increased saccharification of plant biomass.

The CRISPR/Cas9 system has been successfully applied for gene editing in filamentous fungi. Previous studies reported that single stranded oligonucleotides can be used as repair templates to induce point mutations in some filamentous fungi belonging to genus Aspergillus. In Aspergillus niger, extensive research has been performed on regulation of plant biomass degradation, addressing transcription factors such as XlnR or GaaR, involved in (hemi-)cellulose and pectin utilization, respectively. Single nucleotide mutations leading to constitutively active forms of XlnR and GaaR have been previously reported. However, the mutations were performed by the introduction of versions obtained through site-directed or UV-mutagenesis into the genome. Here we report a more time- and cost-efficient approach to obtaining constitutively active versions by application of the CRISPR/Cas9 system to generate the desired mutation on-site in the A. niger genome. This was also achieved using only 60-mer single stranded oligonucleotides, shorter than the previously reported 90-mer strands. In this study, we show that CRISPR/Cas9 can also be used to efficiently change functional properties of the proteins encoded by the target gene by on-site genomic mutations in A. niger. The obtained strains with constitutively active XlnR and GaaR versions resulted in increased production of plant biomass degrading enzymes and improved release of d-xylose and l-arabinose from wheat bran, and d-galacturonic acid from sugar beet pulp.

Mapping the coevolution, leadership and financing of research on viral vectors, RNAi, CRISPR/Cas9 and other genomic editing technologies.

Genomic editing technologies are developing rapidly, promising significant developments for biomedicine, agriculture and other fields. In the present investigation, we analyzed and compared the process of innovation for six genomic technologies: viral vectors, RNAi, TALENs, meganucleases, ZFNs and CRISPR/Cas including the profile of the main research institutions and their funders, to understand how innovation evolved and what institutions influenced research trajectories. A Web of Science search of papers on viral vectors RNAi, CRISPR/Cas, TALENs, ZFNs and meganucleases was used to build a citation network of 16,746 papers. An analysis of network clustering combined with text mining was performed. For viral vectors, a long-term process of incremental innovation was identified, which was largely publicly funded in the United States and the European Union. The trajectory of RNAi research included clusters related to the study of RNAi as a biological phenomenon and its use in functional genomics, biomedicine and pest control. A British philanthropic organization and a US pharmaceutical company played a key role in the development of basic RNAi research and clinical application respectively, in addition to government and academic institutions. In the case of CRISPR/Cas research, basic science discoveries led to the technical improvements, and these two in turn provided the information required for the development of biomedical, agricultural, livestock and industrial applications. The trajectory of CRISPR/Cas research exhibits a geopolitical division of the investigation efforts between the US, as the main producer and funder of basic research and technical improvements, and Chinese research institutions increasingly leading applied research. Our results reflect a change in the model for financing science, with reduced public financing for basic science and applied research on publicly funded technological developments in the US, and the emergence of China as a scientific superpower, with implications for the development of applications of genomic technologies.

Evaluation and minimization of Cas9-independent off-target DNA editing by cytosine base editors.

Cytosine base editors (CBEs) enable targeted C•G-to-T•A conversions in genomic DNA. Recent studies report that BE3, the original CBE, induces a low frequency of genome-wide Cas9-independent off-target C•G-to-T•A mutation in mouse embryos and in rice. Here we develop multiple rapid, cost-effective methods to screen the propensity of different CBEs to induce Cas9-independent deamination in Escherichia coli and in human cells. We use these assays to identify CBEs with reduced Cas9-independent deamination and validate via whole-genome sequencing that YE1, a narrowed-window CBE variant, displays background levels of Cas9-independent off-target editing. We engineered YE1 variants that retain the substrate-targeting scope of high-activity CBEs while maintaining minimal Cas9-independent off-target editing. The suite of CBEs characterized and engineered in this study collectively offer ~10-100-fold lower average Cas9-independent off-target DNA editing while maintaining robust on-target editing at most positions targetable by canonical CBEs, and thus are especially promising for applications in which off-target editing must be minimized.

The Daunting Economics of Therapeutic Genome Editing.

There is no shortage of enthusiasm for the clinical potential of CRISPR-based genome editing: many life-changing cures appear to be just around the corner. However, as mature genetic therapies reach the market, it seems that million-dollar price tags are the new normal. Several factors contribute to the extreme pricing of next-generation medicines, including the need to recoup development costs, the undeniable value of these powerful therapies, and the inherent technical challenges of manufacture and delivery. CRISPR technology has been hailed as a great leveler and a democratizing force in biomedicine. But for this principle to hold true in clinical contexts, therapeutic genome editing must avoid several pitfalls that could substantially limit access to its transformative potential, especially in the developing world.

An artificial triazole backbone linkage provides a split-and-click strategy to bioactive chemically modified CRISPR sgRNA.

As the applications of CRISPR-Cas9 technology diversify and spread beyond the laboratory to diagnostic and therapeutic use, the demands of gRNA synthesis have increased and access to tailored gRNAs is now restrictive. Enzymatic routes are time-consuming, difficult to scale-up and suffer from polymerase-bias while existing chemical routes are inefficient. Here, we describe a split-and-click convergent chemical route to individual or pools of sgRNAs. The synthetic burden is reduced by splitting the sgRNA into a variable DNA/genome-targeting 20-mer, produced on-demand and in high purity, and a fixed Cas9-binding chemically-modified 79-mer, produced cost-effectively on large-scale, a strategy that provides access to site-specific modifications that enhance sgRNA activity and in vivo stability. Click ligation of the two components generates an artificial triazole linkage that is tolerated in functionally critical regions of the sgRNA and allows efficient DNA cleavage in vitro as well as gene-editing in cells with no unexpected off-target effects.

Removal of extra sequences with I-SceI in combination with CRISPR/Cas9 technique for precise gene editing in Drosophila.

The CRISPR/Cas9 system has recently emerged as a powerful tool for functional genomic studies and has been adopted for many organisms, including Drosophila. Previously, an efficient two-step strategy was developed to engineer the fly genome by combining CRISPR/Cas9 with recombinase-mediated cassette exchange (RMCE). This strategy allows the introduction of designed mutations into a gene of interest in vivo. However, the loxP or frt site remains in the edited locus. Here, we propose a modification of this approach for rapid and efficient seamless genome editing with CRISPR/Cas9 and site-specific recombinase-mediated integration (SSRMI) combined with recombination between homologous sequences induced by the rare-cutting endonuclease I-SceI. The induced homological recombination leads to the removal of the remaining extraneous sequences from the target locus.

Genome editing in primary cells and in vivo using viral-derived Nanoblades loaded with Cas9-sgRNA ribonucleoproteins.

Programmable nucleases have enabled rapid and accessible genome engineering in eukaryotic cells and living organisms. However, their delivery into target cells can be technically challenging when working with primary cells or in vivo. Here, we use engineered murine leukemia virus-like particles loaded with Cas9-sgRNA ribonucleoproteins (Nanoblades) to induce efficient genome-editing in cell lines and primary cells including human induced pluripotent stem cells, human hematopoietic stem cells and mouse bone-marrow cells. Transgene-free Nanoblades are also capable of in vivo genome-editing in mouse embryos and in the liver of injected mice. Nanoblades can be complexed with donor DNA for "all-in-one" homology-directed repair or programmed with modified Cas9 variants to mediate transcriptional up-regulation of target genes. Nanoblades preparation process is simple, relatively inexpensive and can be easily implemented in any laboratory equipped for cellular biology.

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.

Unlocking the potential of CRISPR technology for improving livelihoods in Africa.

Africa is burdened with food shortages and plant, animal and human diseases. Some of these can be ameliorated by adopting genome editing technologies such as CRISPR. This technology is considered better than its predecessors, Zinc-finger nucleases (ZFNs) and transcription activator-like effector nucleases (TALENs), because it is cheaper, easy to use, has high gene modification efficiency and is less time consuming. CRISPR technology has wide applications in the African context ranging from crop and animal improvement to disease diagnosis and treatment as well as improving food shelf life, organoleptic properties and food safety. It has the potential to bring back species of organisms that are extinct. However, some African countries have not taken advantage of the potential of CRISPR to solve many of their problems. This paper explores possible applications of CRISPR towards improvement of African livelihoods.

Efficient Genome Engineering of a Virulent Klebsiella Bacteriophage Using CRISPR-Cas9.

Klebsiella pneumoniae is one of the most common nosocomial opportunistic pathogens and usually exhibits multiple-drug resistance. Phage therapy, a potential therapeutic to replace or supplement antibiotics, has attracted much attention. However, very few Klebsiella phages have been well characterized because of the lack of efficient genome-editing tools. Here, Cas9 from Streptococcus pyogenes and a single guide RNA (sgRNA) were used to modify a virulent Klebsiella bacteriophage, phiKpS2. We first evaluated the distribution of sgRNA activity in phages and proved that it is largely inconsistent with the predicted activity from current models trained on eukaryotic cell data sets. A simple CRISPR-based phage genome-editing procedure was developed based on the discovery that homologous arms as short as 30 to 60 bp were sufficient to introduce point mutation, gene deletion, and swap. We also demonstrated that weak sgRNAs could be used for precise phage genome editing but failed to select random recombinants, possibly because inefficient cleavage can be tolerated through continuous repair by homologous recombination with the uncut genomes. Small frameshift deletion was proved to be an efficient way to evaluate the essentiality of phage genes. By using the abovementioned strategies, a putative promoter and nine genes of phiKpS2 were successfully deleted. Interestingly, the holin gene can be deleted with little effect on phiKpS2 infection, but the reason is not yet clear. This study established an efficient, time-saving, and cost-effective procedure for phage genome editing, which is expected to significantly promote the development of bacteriophage therapy.IMPORTANCE In the present study, we have addressed efficient, time-saving, and cost-effective CRISPR-based phage genome editing of Klebsiella phage, which has the potential to significantly expand our knowledge of phage-host interactions and to promote applications of phage therapy. The distribution of sgRNA activity was first evaluated in phages. Short homologous arms were proven to be enough to introduce point mutation, small frameshift deletion, gene deletion, and swap into phages, and weak sgRNAs were proven useful for precise phage genome editing but failed to select random recombinants, all of which makes the CRISPR-based phage genome-editing method easier to use.