CRISPR Journal最新文献

Flax is an important crop used for oil and fiber production. Although genetic engineering has been possible in flax, it is not commonly used to produce cultivars. However, the use of genome editing technology, which can produce site-specific mutations without introducing foreign genes, may be a valuable tool for creating elite cultivars that can be easily cultivated. The purpose of this study was to investigate the potential of genome editing in flax by establishing the clustered regularly interspaced short palindromic repeats (CR ISPR)-CRISPR-associated protein 9 (CRISPR-Cas9) genome editing system using the phytoene desaturase (PDS) gene, which produces albino mutants that are easily identifiable. Four sgRNAs were designed from two PDS genes of Flax (LuPDS1 and LuPDS2), and CRISPR-Cas9 genome editing vectors were constructed. After gene transformation, albino phenotypes were observed in transformed callus and regenerated plantlets on selection media. Polymerase chain reaction (PCR) amplification and sequencing of the PDS genes revealed deletions and insertions in the albino tissues, indicating successful editing of the PDS genes. Potential off-target sites were analyzed, but no off-target mutations were found, indicating the specificity of the CRISPR-Cas9 system. The establishment of a flax genome editing system using the CRISPR-Cas9 technology opens up new possibilities for the genetic engineering of flax. This study demonstrates the potential of genome editing in creating elite cultivars that can be easily cultivated, which can have significant implications for the flax industry.

Tuberous sclerosis complex (TSC) is an autosomal dominant disorder caused by mutations in either the TSC1 or TSC2 genes. Though TSC causes the formation of nonmalignant tumors throughout multiple organs, the most frequent causes of mortality and morbidity are due to neurological complications. In two-thirds of cases, TSC occurs sporadically and TSC2 pathogenic variants are approximately three times more prevalent than TSC1 pathogenic variants. Here, we utilized CRISPR-Cas9-mediated homology directed repair in patient induced pluripotent stem cells (iPSCs) to correct two types of TSC2 pathogenic variants generating two isogenic lines. In one line, we corrected a splice acceptor variant (c.2743-1G>A), which causes the skipping of coding exon 23 and subsequent frameshift and introduction of a stop codon in coding exon 25. In the second line, we corrected a missense variant in coding exon 40 within the GTPase-activating protein domain (c.5228G>A, p.R1743Q). The generation of TSC2 patient iPSCs in parallel with their corresponding CRISPR-corrected isogenic lines will be an important tool for disease modeling applications and for developing therapeutics.

In mice, naturally occurring and induced mutations in the suppressor of cytokine signaling-2 (Socs2) gene are associated with a high growth phenotype characterized by rapid post-weaning weight gain and 30-50% heavier mature body weight. In this work, we demonstrate an electroporation-based method of producing SOCS2 knock-out (KO) sheep. Electroporation of dual-guide CRISPR-Cas9 ribonucleoprotein complexes targeting SOCS2 was performed 6 h post-fertilization in sheep zygotes. Fifty-two blastocysts were transferred to 13 estrus-synchronized recipients, yielding five live lambs and one stillborn. These lambs all carried mutations predicted to result in SOCS2 KO. Three carried large deletion alleles which evaded detection in initial PCR screening. Off-target analysis using whole genome sequencing comparing the frequency of mutations in regions within 100 bp of possible sgRNA binding sites (up to 4 bp mismatches) and elsewhere in the genome showed no significant difference when comparing unedited control sheep to edited animals (p = 0.71). In conclusion, electroporation of zygotes with dual-guide CRISPR-Cas9 RNPs was effective at generating knock-out sheep with no substantial off-target activity.

The clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 system has revolutionized targeted mutagenesis, but screening for mutations in large sample pools can be time-consuming and costly. We present an efficient and cost-effective polymerase chain reaction (PCR)-based strategy for identifying edited mutants in the T₁ generation. Unlike previous methods, our approach addresses the challenges of large progeny populations by using T₀ generation sequencing results for genotype prediction. The T₁ generation plants were then divided into two scenarios: ≥4 bp indels and 1-2 bp indels. Specific primers are designed for these categories, employing dual-primers critical annealing temperature PCR for ≥4 bp indels and the derived cleaved amplified polymorphic sequences (dCAPS) method for 1-2 bp indels. This method is straightforward, cost-effective, and allows rapid and precise identification of T₁ editing outcomes, distinguishing between wild-type, heterozygous, and homozygous plants. This strategy accelerates gene functional analysis in plants and beyond.

Clustered regularly interspaced short palindromic repeats (CRISPR)-CRISPR-associated protein (Cas) technology has revolutionized genome editing across various biological systems, including the Apicomplexa phylum. This review describes the status, challenges, and applications of CRISPR-Cas9 editing technology in apicomplexan parasites, such as Plasmodium, Toxoplasma, Theileria, Babesia, and Cryptosporidium. The discussion encompasses successfully implemented CRISPR-Cas9-based techniques in these parasites, highlighting the achieved milestones, from precise gene modifications to genome-wide screening. In addition, the review addresses the challenges hampering efficient genome editing, including the parasites' complex life cycles, multiple intracellular stages, and the lack of robust genetic tools. It further explores the ethical and policy considerations surrounding genome editing and the future perspectives of CRISPR-Cas applications in apicomplexan parasites.

Genome-wide genetic screens using CRISPR-guide RNA libraries are widely performed in mammalian cells to functionally characterize individual genes and for the discovery of new anticancer therapeutic targets. As the effectiveness of such powerful and precise tools for cancer pharmacogenomics is emerging, tools and methods for their quality assessment are becoming increasingly necessary. Here, we provide an R package and a high-quality reference data set for the assessment of novel experimental pipelines through which a single calibration experiment has been executed: a screen of the HT-29 human colorectal cancer cell line with a commercially available genome-wide library of single-guide RNAs. This package and data allow experimental researchers to benchmark their screens and produce a quality-control report, encompassing several quality and validation metrics. The R code used for processing the reference data set, for its quality assessment, as well as to evaluate the quality of a user-provided screen, and to reproduce the figures presented in this article is available at https://github.com/DepMap-Analytics/HT29benchmark. The reference data is publicly available on FigShare.

Wildlife diseases are a considerable threat to human health, conservation, and the economy. Surveillance is a critical component to mitigate the impact of animal diseases in these sectors. To monitor human diseases, CRISPR-Cas (clustered regularly interspaced short palindromic repeats-CRISPR-associated protein) biosensors have proven instrumental as diagnostic tools capable of detecting unique DNA and RNA sequences related to their associated pathogens. However, despite the significant advances in the general development of CRISPR-Cas biosensors, their use to support wildlife disease management is lagging. In some cases, wildlife diseases of concern could be rapidly surveyed using these tools with minimal technical, operational, or cost requirements to end users. This review explores the potential to further leverage this technology to advance wildlife disease monitoring and highlights how concerted standardization of protocols can help to ensure data reliability.