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CRISPR genome editing in plants without tissue culture.
Conventional plant genome editing relies on tissue culture-mediated somatic cell regeneration, a technically demanding process that limits its application across diverse species. Emerging strategies now circumvent this bottleneck by enabling direct genome editing of meristematic or germline cells. Key advances include (i) genome editing via de novo meristem induction or dormant meristem activation; (ii) germline editing facilitated by graft-mobile tRNA-like sequence systems and haploid induction technologies; and (iii) optimized viral delivery platforms that exploit mobile RNA elements and compact editors such as TnpB to achieve efficient, transgene-free, heritable modifications across a broad range of genotypes and species. The development of robust, tissue culture-free editing platforms promises to revolutionize crop improvement pipelines and accelerate trait development for sustainable agriculture.
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Single nucleotide replacement in the Atlantic salmon genome using CRISPR/Cas9 and asymmetrical oligonucleotide donors.
BACKGROUND: New breeding technologies (NBT) using CRISPR/Cas9-induced homology directed repair (HDR) has the potential to expedite genetic improvement in aquaculture. The long generation time in Atlantic salmon makes breeding an unattractive solution to obtain homozygous mutants and improving the rates of perfect HDR in founder (F0) fish is thus required. Genome editing can represent small DNA changes down to single nucleotide replacements (SNR). This enables edits such as premature stop codons or single amino acid changes and may be used to obtain fish with traits favorable to aquaculture, e.g. disease resistance. A method for SNR has not yet been demonstrated in salmon. RESULTS: Using CRISPR/Cas9 and asymmetrical ODNs, we were able to perform precise SNR and introduce a premature stop codon in dnd in F0 salmon. Deep sequencing demonstrated up to 59.2% efficiency in single embryos. In addition, using the same asymmetrical ODN design, we inserted a FLAG element into slc45a2 and dnd, showing high individual perfect HDR efficiencies (up to 36.7 and 32.7%, respectively). CONCLUSIONS: In this work, we demonstrate that precise SNR and knock-in (KI) can be performed in F0 salmon embryos using asymmetrical oligonucleotide (ODN) donors. We suggest that HDR-induced SNR can be applied as a powerful NBT, allowing efficient introgression of favorable alleles and bypassing challenges associated with traditional selective breeding.
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Francisella novicida Cas9 interrogates genomic DNA with very high specificity and can be used for mammalian genome editing.
Genome editing using the CRISPR/Cas9 system has been used to make precise heritable changes in the DNA of organisms. Although the widely used Streptococcus pyogenes Cas9 (SpCas9) and its engineered variants have been efficiently harnessed for numerous gene-editing applications across different platforms, concerns remain regarding their putative off-targeting at multiple loci across the genome. Here we report that Francisella novicida Cas9 (FnCas9) shows a very high specificity of binding to its intended targets and negligible binding to off-target loci. The specificity is determined by its minimal binding affinity with DNA when mismatches to the target single-guide RNA (sgRNA) are present in the sgRNA:DNA heteroduplex. FnCas9 produces staggered cleavage, higher homology-directed repair rates, and very low nonspecific genome editing compared to SpCas9. We demonstrate FnCas9-mediated correction of the sickle cell mutation in patient-derived induced pluripotent stem cells and propose that it can be used for precise therapeutic genome editing for a wide variety of genetic disorders.
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A single-plasmid-based, easily curable CRISPR/Cas9 system for rapid, iterative genome editing in Pseudomonas putida KT2440.
BACKGROUND: Pseudomonas putida KT2440, a non-pathogenic soil bacterium, is a key platform strain in synthetic biology and industrial applications due to its robustness and metabolic versatility. Various systems have been developed for genome editing in P. putida, including transposon modules, integrative plasmids, recombineering systems, and CRISPR/Cas systems. However, rapid iterative genome editing is limited by complex and lengthy processes. RESULTS: We discovered that the pBBR1MCS2 plasmid carrying the CRISPR/Cas9 module could be easily cured in P. putida KT2440 at 30 oC. We then developed an all-in-one CRISPR/Cas9 system for yqhD and ech-vdh-fcs deletions, respectively, and further optimized the editing efficiency by varying homology arm lengths and target sites. Sequential gene deletions of vdh and vanAB were carried out rapidly using single-round processing and easy plasmid curing. This system's user-friendliness was validated by 3 researchers from two labs for 9 deletions, 3 substitutions, and 2 insertions. Finally, iterative genome editing was used to engineer P. putida for valencene biosynthesis, achieving a 10-fold increase in yield. CONCLUSIONS: We developed and applied a rapid all-in-one plasmid CRISPR/Cas9 system for genome editing in P. putida. This system requires less than 1.5 days for one edit due to simplified plasmid construction, electroporation and curing processes, thus accelerating the cycle of genome editing. To our knowledge, this is the fastest iterative genome editing system for P. putida. Using this system, we rapidly engineered P. putida for valencene biosynthesis for the first time, showcasing the system's potential for expanding biotechnological applications.
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Reduced Susceptibility to Phytophthora in Non-Transgenic Cacao Progeny Through CRISPR-Cas9 Mediated TcNPR3 Mutagenesis.
Black pod disease, caused by a complex of Phytophthora species, poses a severe threat to global cacao production. This study explores the use of CRISPR-Cas9 genome editing to reduce disease susceptibility in Theobroma cacao L. by targeting the TcNPR3 gene, a known negative regulator of plant defence. Transgenic T0 lines carrying mutations predicted to disrupt TcNPR3 function exhibited reduced susceptibility to Phytophthora infection in in vitro foliar assays. These T0 plants were advanced to maturity and outcrossed with non-transgenic cacao to eliminate T-DNA sequences associated with the CRISPR-Cas9 transgene. Whole-genome sequencing of the T0 parents and 22 progeny revealed single T-DNA insertion sites in each T0 line; seven progeny retained the edited npr3 alleles but lacked T-DNA insertions. Transcriptome analysis of the mutant lines showed upregulation of genes associated with reactive oxygen species (ROS) generation, defence-related transcription factors and pathogenesis-related proteins. Several genes were also downregulated, suggesting that TcNPR3 may function as both a repressor and an activator in regulating basal transcriptional states. Genome-edited plants were phenotypically comparable to wild-type controls and displayed a 42% reduction in lesion size upon Phytophthora challenge. These findings demonstrate the feasibility of generating non-transgenic cacao with reduced susceptibility to Phytophthora through CRISPR-Cas9-mediated genome editing, offering a promising strategy for sustainable cacao cultivation and improved farmer livelihoods. Field trials are underway to evaluate long-term agronomic performance under natural conditions.
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Identifying and Isolating Meiotic Mutants in a Polyploid Brassica Crop.
This chapter provides a detailed description of TILLING and CRISPR-Cas9 approaches for the purpose of studying genes/factors involved in meiotic recombination in the polyploid species B. napus. The TILLING approach involves the screening and identification of EMS-mutagenized M2 B. napus plants. The strategy for high-throughput plant pooling, the set up for microfluidic PCR and sequencing is provided and the parameters for the analysis of sequence results and the detection of mutants are explained. The CRISPR-Cas system relies on the optimal design of guide RNAs and their efficient expression. The procedure for the generation and detection of knockout mutants is described with the aims to simultaneously target homologous genes.
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Oligo-Modified Poly(β-Amino Ester)-Mediated mRNA Delivery: A Protocol for Gene Editing Applications.
Oligopeptide-modified poly(β-amino ester)s (OM-pBAEs) are polymer-based delivery systems that enable the encapsulation and intracellular transport of nucleic acids, including messenger RNA (mRNA) and mixtures of mRNA with DNA. OM-pBAEs form cationic nanoparticles (NPs) through electrostatic interactions with nucleic acids, protecting them from degradation and enhancing cellular uptake. Terminal oligopeptides composed of lysine, arginine, or histidine enhance polyplex stability, nucleic acid binding, and endosomal escape. This chapter describes a protocol for preparing OM-pBAE nanoparticles, including polymer synthesis, peptide modification, controlled mixing with nucleic acids to form polyplexes, precipitation in RNase-free water, stabilization with excipients, and optional freeze-drying for long-term storage. Nanoparticle formation occurs at a constant temperature of 25°C, with incubation to ensure reproducible self-assembly. Nanoparticles are characterized by using dynamic light scattering (DLS) to determine particle size, polydispersity, and surface charge, while nucleic acid encapsulation efficiency is assessed to evaluate loading capacity. This protocol provides a versatile, nonviral platform for efficient nucleic acid delivery, supporting applications such as gene-editing experiments.
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Viral vectors as carriers of genome-editing reagents.
The presence of a transgene in the genome of plants is a regulatory challenge. Recently, Liu et al. reported an engineered tomato spotted wilt virus (TSWV) that can carry large clustered regularly interspaced palindromic repeats (CRISPR)/Cas reagents for targeted genome editing in various crops without the integration of the transgene into the genome.
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Developing a robust genome editing tool based on an endogenous type I-B CRISPR-Cas system in Saccharopolyspora spinosa.
Saccharopolyspora spinosa is an industrial rare actinomycete capable of producing important environmental-friendly biopesticides, spinosyns. However, exploitation of S. spinosa has been limited due to its genetic inaccessibility and lack of effective genome engineering tools. In this work, we characterized the activity of an endogenous type I-B CRISPR-Cas system as well as its recognized protospacer adjacent motifs (PAMs) based on bioinformatics analysis combined with a plasmid interference assay in S. spinosa. By delivering editing plasmids containing a designed miniCRISPR array (repeat+self-targeting spacer+repeat) and repair templates, we achieved 100% editing efficiency for gene deletion. Using this tool, the genetic barrier composed of the restriction-modification (RM) systems was systematically disarmed. We showed that by disarming one type I RM system (encoded by A8926_1903/1904/1905) and two type II RM systems (encoded by A8926_1725/1726 and A8926_2652/2653) simultaneously, the transformation efficiency of the replicative and integrative plasmids (pSP01 and pSI01) was increased by approximately 3.9-fold and 4.2-fold, respectively. Using the engineered strain with simultaneous knock-out of these three RM genes as the starting strain, we achieved the deletion of 75-kb spinosyns biosynthetic gene cluster (BGC) as well as gene insertion at high efficiency. Collectively, we developed a reliable and highly efficient genome editing tool based on the endogenous type I CRISPR-Cas system combined with the disarmament of the RM systems in S. spinosa. This is the first time to establish an endogenous CRISPR-Cas-based genome editing tool in the non-model industrial actinomycetes.
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Insights Gained from RNA Editing Targeted by the CRISPR-Cas13 Family.
Clustered regularly interspaced short palindromic repeat (CRISPR)/CRISPR-associated protein (Cas) systems, especially type II (Cas9) systems, have been widely developed for DNA targeting and formed a set of mature precision gene-editing systems. However, the basic research and application of the CRISPR-Cas system in RNA is still in its early stages. Recently, the discovery of the CRISPR-Cas13 type VI system has provided the possibility for the expansion of RNA targeting technology, which has broad application prospects. Most type VI Cas13 effectors have dinuclease activity that catalyzes pre-crRNA into mature crRNA and produces strong RNA cleavage activity. Cas13 can specifically recognize targeted RNA fragments to activate the Cas13/crRNA complex for collateral cleavage activity. To date, the Cas13X protein is the smallest effector of the Cas13 family, with 775 amino acids, which is a promising platform for RNA targeting due to its lack of protospacer flanking sequence (PFS) restrictions, ease of packaging, and absence of permanent damage. This study highlighted the latest progress in RNA editing targeted by the CRISPR-Cas13 family, and discussed the application of Cas13 in basic research, nucleic acid diagnosis, nucleic acid tracking, and genetic disease treatment. Furthermore, we clarified the structure of the Cas13 protein family and their molecular mechanism, and proposed a future vision of RNA editing targeted by the CRISPR-Cas13 family.
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