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Construction of humanized CYP1A2 rats using CRISPR/Cas9 to promote drug metabolism and pharmacokinetic research.
Cytochrome P450 family 1 subfamily A member 2 (CYP1A2), performs an indispensable role in metabolism of both exogenous and endogenous substances. What is more, CYP1A2 functions in human diseases by regulating homeostasis of cholesterol. Despite the emergence of gene-editing animal models, genetically humanized animals that overcome species differences for further exploring the role of CYP1A2 in drug metabolism and human diseases have not yet been constructed. In this study, we inserted human CYP1A2 cDNA into the rat Cyp1a2 gene by using CRISPR/Cas9 technology. Results showed that human CYP1A2 was successfully expressed in humanized rat liver and there were no statistically significant differences of physiological symptoms compared with wild-type (WT) rats. In vitro incubation results indicated the different inhibition of furafylline on CYP1A2 activity in human liver microsomes, humanized CYP1A2 (hCYP1A2) rat liver microsomes, and WT rat liver microsomes, with IC50 values of 7.1 μM, 36.5 μM, and 285.8 μM, respectively. Meanwhile, pharmacokinetic characteristics of clozapine were conducted, and the results suggested that in hCYP1A2 rats, clozapine tended to be metabolized into norclozapine. Both the in vitro and in vivo results demonstrated the different metabolic functions of CYP1A2 in humanized and WT rats. We successfully constructed a novel humanized CYP1A2 rat model using the CRISPR/Cas9 system, providing a powerful tool for better predicting CYP1A2-mediated drug metabolism and pharmacokinetics. Significance Statement Human CYP1A2 takes active part both in the biotransformation of exogenous substances and endogenous substances. Meanwhile, it plays a regulatory role in human diseases, including hypercholesterolemia, hypertension as well as various malignant tumors. This study successfully constructed humanized CYP1A2 rat model by CRISPR/Cas9 technology, providing a powerful model for promoting drug development and safety evaluation, as well as further exploring the role of CYP1A2 in human diseases.
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Modeling Dominant and Recessive Forms of Retinitis Pigmentosa by Editing Three Rhodopsin-Encoding Genes in Xenopus Laevis Using Crispr/Cas9.
The utility of Xenopus laevis, a common research subject for developmental biology, retinal physiology, cell biology, and other investigations, has been limited by lack of a robust gene knockout or knock-down technology. Here we describe manipulation of the X. laevis genome using CRISPR/Cas9 to model the human disorder retinitis pigmentosa, and to introduce point mutations or exogenous DNA sequences. We introduced and characterized in-frame and out-of-frame insertions and deletions in three genes encoding rhodopsin by co-injection of Cas9 mRNA, eGFP mRNA, and single guide RNAs into fertilized eggs. Deletions were characterized by direct sequencing and cloning; phenotypes were assessed by assays of rod opsin in retinal extracts, and confocal microscopy of cryosectioned and immunolabeled contralateral eyes. We obtained germline transmission of editing to F1 offspring. In-frame deletions frequently caused dominant retinal degeneration associated with rhodopsin biosynthesis defects, while frameshift phenotypes were consistent with knockout. We inserted eGFP or point mutations into rhodopsin genes by co-injection of repair fragments with homology to the Cas9 target sites. Our techniques can produce high frequency gene editing in X. laevis, permitting analysis in the F0 generation, and advancing the utility of X. laevis as a subject for biological research and disease modeling.
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Genome editing of susceptibility and resistance genes for engineering disease-resistant crops: Recent advances, challenges, and future prospects.
Plant diseases caused by bacterial, fungal, oomycete, and viral pathogens remain a major constraint to crop productivity worldwide. Recent advances in genome-editing technologies have enabled precise manipulation of genes involved in plant-pathogen interactions, providing new opportunities for developing disease-resistant crops. In particular, CRISPR/Cas-based systems have facilitated functional dissection and engineering of susceptibility genes, resistance genes, and immune regulatory networks across diverse crop species. In this review, we initially discuss emerging genome-engineering platforms and highlighting their potential for precise disease-resistance engineering. We then summarize current progress in genome-editing strategies for improving resistance against bacterial, fungal, oomycete, and viral pathogens through targeted manipulation of host susceptibility factors, immune receptors, transcription factors, and defense-signaling components, while critically evaluating the limitations and challenges associated with the reported approaches. The review further examines pleiotropic effects and fitness costs associated with susceptibility-gene editing, alongside the functional validation and precision engineering of classical resistance genes and quantitative and multilocus resistance mechanisms. Recent field-validation studies are also discussed in relation to agronomic performance, while ethical, regulatory, and policy considerations governing the adoption of gene-edited crops are addressed. Emerging applications of CRISPR-based transcriptional regulation and de novo domestication are highlighted as complementary strategies for developing disease-resistant crops. Finally, we discuss current challenges and future directions, emphasizing AI-driven target discovery and gRNA design and the transition from single-gene modification toward integrated disease-resistance engineering. Collectively, these advances highlight the potential of genome editing to accelerate the development of durable, agronomically competitive, and precisely engineered disease-resistant crops.
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Optimized protocol for efficient generation, confirmation, transformation, and CRISPR editing of grapevine hairy roots.
Hairy root cultures (HRCs) are powerful tools in plant biotechnology but show variable establishment efficiencies, limiting broader applications. Here, we present a standardized and optimized reference methodology for the routine generation, multiplication, and maintenance of HRCs across diverse grapevine genotypes. Our workflow evaluated three Rhizobium strains, seven grapevine cultivars (three Vitis vinifera cultivars; four Vitis rootstock hybrids), multiple explant types, infection protocols, co-cultivation times, growth media types, and anti-browning agents. The resulting protocol was effective for all grapevine genotypes and, with minor adjustments, also yielded HRCs from two other important South African plant species, namely Sutherlandia frutescens and Aspalathus linearis. Useful molecular tools were developed for transformation and selection of HRCs, including universal multiplex primers for confirmation of transformation, tested antibiotic resistance markers (kanamycin and hygromycin), and fluorescent reporters (DsRed and eyGFPuv), with DsRed found to be particularly versatile. To test the system, we overexpressed the VviMYBA1 transcription factor gene, leading to increased anthocyanin accumulation and red pigmentation in HRCs. Additionally, we achieved CRISPR/Cas9 editing of the VviPUB19 gene, the first report of CRISPR-edited grapevine HRCs. Gene editing combined with HRCs can facilitate rapid gene function studies, offering an efficient alternative or pre-screening system to whole-plant transformations, that could support advanced functional genomics and biotechnological applications in grapevine.
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Generating waxy rice starch with target type of amylopectin fine structure and gelatinization temperature by waxy gene editing.
Waxy rice, which lacks amylose, is an important variant in rice, and its starches have been widely used. New waxy rice varieties generated via the CRISPR/Cas9 gene-editing system is described. Herein, four waxy rice starches with different physicochemical properties were successfully obtained by the CRISPR/Cas9 editing Waxy (Wx) gene. The results showed that the amylose content (AC) of wx mutant starches ranged from 0.26 to 1.78 %, and CZBwx1 starches had the best gel consistency and highest water solubility among all wx mutants. Mutations of Wxb allele produced more short-chains (degree of chain polymerization (DP) 6-11), and less medium- and long-chains (DP12-70) than that of Wxa, while the AC of Wxa allele mutants was higher than the mutations of Wxb allele. The gelatinization temperature (GT) of wxa mutant starches was higher than that of wxb mutant starches. Moreover, we found that the GT and amylopectin fine structure type of waxy rice starch did not change after Wx gene editing. Based on these findings, it is possible to produce waxy rice starch with different physicochemical properties, containing target GT and chain length distributions of amylopectin.
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Nuclear-targeted CRISPR/Cas delivery by using aptamer-modified polymer lipid nanoparticles.
Efficient nuclear delivery of CRISPR/Cas ribonucleoproteins (RNP) remains a significant hurdle for non-viral systems. To address this, we developed a polymer-lipid hybrid nanoparticle functionalized with the AS1411 aptamer, targeting nucleolin to facilitate nucleus-directed delivery. Confocal imaging confirmed the accumulation of these aptamer-modified nanoparticles within the cell nuclei. For precise quantification, we utilized an AI-assisted segmentation approach based on deep convolutional neural networks (CNN) to analyse nanoparticle and DAPI colocalization. We further evaluated in vitro gene knockout efficiency of Cas9/sgRNA by using this nucleus-targeted system. Aptamer-functionalised nanoparticles reduced GFP-positive cells to 30.0%, compared with 40.8% for untargeted nanoparticles. Further evaluation targeting the Lcn2 gene demonstrated higher knockout efficacy and a more potent inhibition of breast cancer cell proliferation. These findings indicate that aptamer-mediated nuclear targeting enhances CRISPR/Cas9 editing efficacy and may offer the potential to advance the performance of non-viral gene therapies.
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Challenges of CRISPR-Based Gene Editing in Primary T Cells.
Adaptive T-cell immunotherapy holds great promise for the successful treatment of leukemia, as well as other types of cancers. More recently, it was also shown to be an effective treatment option for chronic virus infections in immunosuppressed patients. Autologous or allogeneic T cells used for immunotherapy are usually genetically modified to express novel T-cell or chimeric antigen receptors. The production of such cells was significantly simplified with the CRISPR/Cas system, allowing for the deletion or insertion of novel genes at specific locations within the genome. In this review, we describe recent methodological breakthroughs that were important for the conduction of these genetic modifications, summarize crucial points to be considered when conducting such experiments, and highlight the potential pitfalls of these approaches.
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hpCasMINI: An engineered hypercompact CRISPR-Cas12f system with boosted gene editing activity.
Compact CRISPR-Cas systems have demonstrated potential for effective packaging into adeno-associated viruses (AAVs) for use in gene therapy. However, their applications are currently limited due to modest gene-editing activity. Here we introduce an engineered compact CRISPR-Cas12f (hpCasMINI, 554 aa), with hyper editing efficiency in mammalian cells via adding an α-helix structure to the N-terminus of an Un1Cas12f1 variant CasMINI (529 aa). The hpCasMINI system boosts gene activation and DNA cleavage activity with about 1.4-3.0-fold and 1.1-19.5-fold, respectively, and maintains the high specificity when compared to CasMINI. In addition, the system can activate luciferase reporter gene and endogenous Fgf21 gene in adult mouse liver, as well as construct liver tumorigenesis model via disrupting Trp53 and Pten genes and inserting oncogenic KrasG12D into the Trp53 locus. When compared to SpCas9 and LbCas12a, hpCasMINI displays higher gene activation and exhibits higher DNA cleavage specificity, although with lower activity, at the tested sites. Moreover, with a similar strategy, we engineer compact versions of hpOsCas12f1 (458 aa) from enOsCas12f1 and hpAsCas12f1 (447 aa) from AsCas12f1-HKRA, both of which display increased DNA cleavage activity, with hpAsCas12f1 also showing improved gene activation capability. Therefore, we develop activity-increased miniature hpCasMINI, hpOsCas12f1 and hpAsCas12f1 nucleases, which hold great potential for gene therapy in the future.
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Lessons from Domestication: Targeting Cis-Regulatory Elements for Crop Improvement.
Domestication of wild plant species has provided us with crops that serve our human nutritional needs. Advanced DNA sequencing has propelled the unveiling of underlying genetic changes associated with domestication. Interestingly, many changes reside in cis-regulatory elements (CREs) that control the expression of an unmodified coding sequence. Sequence variation in CREs can impact gene expression levels, but also developmental timing and tissue specificity of expression. When genes are involved in multiple pathways or active in several organs and developmental stages CRE modifications are favored in contrast to mutations in coding regions, due to the lack of detrimental pleiotropic effects. Therefore, learning from domestication, we propose that CREs are interesting targets for genome editing to create new alleles for plant breeding.
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Harnessing CRISPR-Cas9 for Genome Editing in Streptococcus pneumoniae D39V.
CRISPR-Cas systems provide bacteria and archaea with adaptive immunity against viruses and plasmids by the detection and cleavage of invading foreign DNA. Modified versions of this system can be exploited as a biotechnological tool for precise genome editing at a targeted locus. Here, we developed a replicative plasmid that carries the CRISPR-Cas9 system for RNA-programmable genome editing by counterselection in the opportunistic human pathogen Streptococcus pneumoniae Specifically, we demonstrate an approach for making targeted markerless gene knockouts and large genome deletions. After a precise double-stranded break (DSB) is introduced, the cells' DNA repair mechanism of homology-directed repair (HDR) is exploited to select successful transformants. This is achieved through the transformation of a template DNA fragment that will recombine in the genome and eliminate recognition of the target of the Cas9 endonuclease. Next, the newly engineered strain can be easily cured from the plasmid, which is temperature sensitive for replication, by growing it at the nonpermissive temperature. This allows for consecutive rounds of genome editing. Using this system, we engineered a strain with three major virulence factors deleted. The approaches developed here could potentially be adapted for use with other Gram-positive bacteria.IMPORTANCEStreptococcus pneumoniae (the pneumococcus) is an important opportunistic human pathogen killing more than 1 million people each year. Having the availability of a system capable of easy genome editing would significantly facilitate drug discovery and efforts to identify new vaccine candidates. Here, we introduced an easy-to-use system to perform multiple rounds of genome editing in the pneumococcus by putting the CRISPR-Cas9 system on a temperature-sensitive replicative plasmid. The approaches used here will advance genome editing projects in this important human pathogen.
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