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An MHV-68 Mutator Phenotype Mutant Virus, Confirmed by CRISPR/Cas9-Mediated Gene Editing of the Viral DNA Polymerase Gene, Shows Reduced Viral Fitness.
Drug resistance studies on human γ-herpesviruses are hampered by the absence of an in vitro system that allows efficient lytic viral replication. Therefore, we employed murine γ-herpesvirus-68 (MHV-68) that efficiently replicates in vitro as a model to study the antiviral resistance of γ-herpesviruses. In this study, we investigated the mechanism of resistance to nucleoside (ganciclovir (GCV)), nucleotide (cidofovir (CDV), HPMP-5azaC, HPMPO-DAPy) and pyrophosphate (foscarnet (PFA)) analogues and the impact of these drug resistance mutations on viral fitness. Viral fitness was determined by dual infection competition assays, where MHV-68 drug-resistant viral clones competed with the wild-type virus in the absence and presence of antivirals. Using next-generation sequencing, the composition of the viral populations was determined at the time of infection and after 5 days of growth. Antiviral drug resistance selection resulted in clones harboring mutations in the viral DNA polymerase (DP), denoted Y383SGCV, Q827RHPMP-5azaC, G302WPFA, K442TPFA, G302W+K442TPFA, C297WHPMPO-DAPy and C981YCDV. Without antiviral pressure, viral clones Q827RHPMP-5azaC, G302WPFA, K442TPFA and G302W+K442TPFA grew equal to the wild-type virus. However, in the presence of antivirals, these mutants had a growth advantage over the wild-type virus that was moderately to very strongly correlated with antiviral resistance. The Y383SGCV mutant was more fit than the wild-type virus with and without antivirals, except in the presence of brivudin. The C297W and C981Y changes were associated with a mutator phenotype and had a severely impaired viral fitness in the absence and presence of antivirals. The mutator phenotype caused by C297W in MHV-68 DP was validated by using a CRISPR/Cas9 genome editing approach.
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Digital droplet PCR and IDAA for the detection of CRISPR indel edits in the malaria species Anopheles stephensi.
CRISPR/Cas9 technology is a powerful tool for the design of gene-drive systems to control and/or modify mosquito vector populations; however, CRISPR/Cas9-mediated nonhomologous end joining mutations can have an important impact on generating alleles resistant to the drive and thus on drive efficiency. We demonstrate and compare the insertions or deletions (indels) detection capabilities of two techniques in the malaria vector mosquito Anopheles stephensi: Indel Detection by Amplicon Analysis (IDAA™) and Droplet Digital™ PCR (ddPCR™). Both techniques showed accuracy and reproducibility for indel frequencies across mosquito samples containing different ratios of indels of various sizes. Moreover, these techniques have advantages that make them potentially better suited for high-throughput nonhomologous end joining analysis in cage trials and contained field testing of gene-drive mosquitoes.
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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±35.0μg/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.
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Gene conversion following CRISPR/Cas9 DNA cleavage: an overlooked effect.
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The pSG5-based thermosensitive vector family for genome editing and gene expression in actinomycetes.
Actinomycetes are the most important producers of secondary metabolites for medical, agricultural and industrial applications. Efficient engineering of bacterial genomes to improve their biosynthetic capabilities largely depends on the available arsenal of tools and vectors. One of the most widely used vector systems for actinomycetes is derived from the Streptomyces ghanaensis DSM2932 plasmid pSG5. pSG5 is a broad host range multicopy plasmid replicating via a rolling circle mechanism. The unique feature of pSG5, which distinguishes it from other Streptomyces plasmids, is its naturally thermosensitive mode of replication. This allows the efficient elimination of the plasmid from its host by simply shifting the incubation temperature to non-permissive 37-39 °C. This property makes pSG5 derivatives ideal facultative suicide vectors required for selection of gene disruption/gene replacement, transposon delivery or CRISPR/Cas9-mediated genome editing. Whereas these techniques depend on the fast elimination of the vector, stably replicating expression vectors for the production of recombinant proteins have been constructed more recently. This mini-review describes the generation and application of the pSG5 vector family, highlighting the specific features of the distinct vector plasmids.
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Sequentially Self-Assembled Supramolecular Nanocomplexes Enable Systemic Cas9 RNP Delivery and In Vivo Tumor Genome Editing.
In vivo genome editing utilizing Cas9/sgRNA ribonucleoproteins (RNPs) holds substantial therapeutic promise, yet rapid bloodstream clearance and degradation of RNPs have hindered its accumulation within the target site and effective gene editing. Herein, we report a supramolecular ternary complex platform where RNPs are co-assembled with tannic acid (TA) and phenylboronic acid (PBA)-conjugated polymers through sequential self-assembly. This ∼30 nm core-shell structure protects RNPs from enzymatic degradation and dissociates selectively at endosomal pH. Upon intravenous administration in subcutaneous tumor-bearing mice, the ternary complex exhibits prolonged blood circulation and preferential tumor accumulation, achieving 37.2% gene editing at tumor sites compared with only 1.5% for free RNPs. The platform successfully disrupts PLK1 in subcutaneous tumors, accompanied by significant tumor growth suppression. This is further applied to a KRASG13D-mutant tumor model, in which treatment delayed tumor growth. By integrating sequential supramolecular self-assembly with stimuli-responsive cargo release, this strategy presents a promising approach for systemic RNP delivery and tumor genome editing.
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Advances in CRISPR-Cas9 in lineage tracing of model animals.
Cell lineage tracing is a key technology for describing the developmental history of individual progenitor cells and assembling them to form a lineage development tree. However, traditional methods have limitations of poor stability and insufficient resolution. As an efficient and flexible gene editing tool, CRISPR-Cas9 system has been widely used in biological research. Furthermore, CRISPR-Cas9 gene editing-based tracing methods can introduce fluorescent proteins, reporter genes, or DNA barcodes for high-throughput sequencing, enabling precise lineage analysis, significantly improving precision and resolution, and expanding its application range. In this review, we summarize applications of CRISPR-Cas9 system in cell lineage tracing, with special emphasis on its successful applications in traditional model animals (e.g., zebrafish and mice), large animal models (pigs), and human cells or organoids. We also discussed its potential prospects and challenges in xenotransplantation and regenerative medicine.
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Targeted genome editing of the non-model cyanobacterium Cyanothece PCC 7425 via CRISPR/Cas12a.
Cyanobacteria are diverse photosynthetic microorganisms of great interest for fundamental science and sustainable biotechnological applications. However, their polyploidy makes genetic manipulation challenging and time-consuming. The development of CRISPR/Cas tools has greatly accelerated genome editing and metabolic engineering of some cyanobacterial model species. In this work, we extend the CRISPR/Cas12a system for targeted gene deletion in the non-model cyanobacterium Cyanothece sp. PCC 7425, interesting for its ability to perform intracellular calcium carbonate (CaCO3) biomineralization, nitrogen fixation, etc. We demonstrate for the first time its tractability to gene knockout by generating deletion mutants of four genes (cax3-cax4, gor, and sodB) acting in metabolism and/or response to stresses, using Cas12a-mediated homologous recombination. Importantly, full chromosome segregation was rapidly achieved after a single round of selection in all cases. All mutants were genotypically and phenotypically characterised. Moreover, biochemical analysis in the case of the ΔsodB mutant further confirmed its targeted deletion. Overall, CRISPR/Cas12a provides a rapid and efficient system for genome editing in Cyanothece sp. PCC 7425, establishing this organism as a versatile model for studying oxidative stress pathways, metal toxicity, and moreover, the still poorly known mechanism(s) of intracellular CaCO3 biomineralization. KEY POINTS: • Rapid and efficient CRISPR/Cas12a editing established in Cyanothece sp. PCC 7425. • Fully segregated knockout mutants obtained after a single selection round. • Platform for exploring the biotechnological potential of Cyanothece sp. PCC 7425.
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Targeted genomic translocations and inversions generated using a paired prime editing strategy.
A variety of cancers have been found to have chromosomal rearrangements, and the genomic abnormalities often induced expression of fusion oncogenes. To date, a pair of engineered nucleases including ZFNs, TALENs, and CRISPR-Cas9 nucleases have been used to generate chromosomal rearrangement in living cells and organisms for disease modeling. However, these methods induce unwanted indel mutations at the DNA break junctions, resulting in incomplete disease modeling. Here, we developed prime editor nuclease-mediated translocation and inversion (PETI), a method for programmable chromosomal translocation and inversion using prime editor 2 nuclease (PE2 nuclease) and paired pegRNA. Using PETI method, we successfully introduced DNA recombination in episomal fluorescence reporters as well as precise chromosomal translocations in human cells. We applied PETI to create cancer-associated translocations and inversions such as NPM1-ALK and EML4-ALK in human cells. Our findings show that PETI generated chromosomal translocation and inversion in a programmable manner with efficiencies comparable of Cas9. PETI methods, we believe, could be used to create disease models or for gene therapy.
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Efficient gene knockout in primary human and murine myeloid cells by non-viral delivery of CRISPR-Cas9.
Myeloid cells play critical and diverse roles in mammalian physiology, including tissue development and repair, innate defense against pathogens, and generation of adaptive immunity. As cells that show prolonged recruitment to sites of injury or pathology, myeloid cells represent therapeutic targets for a broad range of diseases. However, few approaches have been developed for gene editing of these cell types, likely owing to their sensitivity to foreign genetic material or virus-based manipulation. Here we describe optimized strategies for gene disruption in primary myeloid cells of human and murine origin. Using nucleofection-based delivery of Cas9-ribonuclear proteins (RNPs), we achieved near population-level genetic knockout of single and multiple targets in a range of cell types without selection or enrichment. Importantly, we show that cellular fitness and response to immunological stimuli is not significantly impacted by the gene editing process. This provides a significant advance in the study of myeloid cell biology, thus enabling pathway discovery and drug target validation across species in the field of innate immunity.
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