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Transcriptomics and phenotypic analysis of OTOF gene knockdown in zebrafish mediated by CRISPR/Cas9.
Deafness is a common genetic disorder, where mutations,in the OTOF gene can disrupt the normal functionof the Otoferlin protein, leading to impaired neurotransmitter release in the inner ear and subsequent deafness. Despite the complexity of the pathogenic mechanism,it is not fully understood. Zebrafish are an excellent model for studying genetically-induced deafness,but there have been no previous reports on the pathogenesis of OTOF in zebrafish.This study successfully established a zebrafish model with mutated OTOF genes using CRISPR/Cas9 gene editing technology to investigate the molecular basis of OTOF-induced deafness. Compared to AB wild type zebrafish, those with low otof expression showed injury and apoptosis of hair cells in the posterior lateral neuromasts along with significant increase in the number of macrophages and apoptotic cells in this region. Additionally, these mutants exhibited a reduction in body length. To further elucidate differences at 5dpf (days post-fertilization) between mutant and wild type zebrafish embryos, RNA-seq analysis was conducted to examine differentially expressed genes (DEGs).A total of 334 up-regulated DEGs and 111 down-regulated DEGs were identified in mutants compared to wild types.KEGG and GO enrichment analyses were performed on these DEGs to identify key signaling pathways and hub DEGs. The findings revealedan increased expression of several genes involved in the HSP70 oxidative stress system, suggesting that OTOF may protect cochlear hair cell from apoptosis induced by oxidative stress through regulation of MAPK signal and HSP70 expression.In summary, the establishment of a zebrafish model with OTOF knockout provides a valuable tool for investigating the function of Otoferlin and understanding the role of the OTOF gene in deafness. These potential molecular insights offer significant contributions towards understanding the pathogenesis of deafness experimental models and serves as a foundation for comprehending the involvement of the OTOF gene.
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Induced Pluripotent Stem Cells and Genome-Editing Tools in Determining Gene Function and Therapy for Inherited Retinal Disorders.
Inherited retinal disorders (IRDs) affect millions of people worldwide and are a major cause of irreversible blindness. Therapies based on drugs, gene augmentation or transplantation approaches have been widely investigated and proposed. Among gene therapies for retinal degenerative diseases, the fast-evolving genome-editing CRISPR/Cas technology has emerged as a new potential treatment. The CRISPR/Cas system has been developed as a powerful genome-editing tool in ophthalmic studies and has been applied not only to gain proof of principle for gene therapies in vivo, but has also been extensively used in basic research to model diseases-in-a-dish. Indeed, the CRISPR/Cas technology has been exploited to genetically modify human induced pluripotent stem cells (iPSCs) to model retinal disorders in vitro, to test in vitro drugs and therapies and to provide a cell source for autologous transplantation. In this review, we will focus on the technological advances in iPSC-based cellular reprogramming and gene editing technologies to create human in vitro models that accurately recapitulate IRD mechanisms towards the development of treatments for retinal degenerative diseases.
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CRISPR/Cas9-based engineered Escherichia coli biosensor for sensitive and specific detection of Cd(II) in drinking water.
Cadmium (Cd) is a ubiquitous pollutant that poses a potential threat to human health. Monitoring Cd(II) in drinking water has significant implications for preventing potential threats of Cd(II) to human. However, the weak signal output and response to nontarget interference limit the detection of Cd(II) using bacterial biosensors. In this study, to enable sensitive and specific detection of Cd(II) in water, a stable whole-cell biosensor, K12-PMP-luxCDABE-△cysI, was constructed in a dual-promoter mode by fusing the mercury promoter Pmer, regulatory gene merR(m), and luciferase gene luxCDABE into the E.coli chromosome based on CRISPR/Cas9 gene editing technology. By knocking out the cadmium-resistance-gene cysI, the sensitivity of the biosensor to Cd(II) was further enhanced. The constructed E. coli biosensor K12-PMP-luxCDABE-△cysI exhibited good nonlinear responses to 0.005-2 mg/L Cd(II). Notably, among the three constructed E. coli biosensor, it exhibited the strongest fluorescence intensity, with the limit of detection meeting the allowable limit for Cd(II) in drinking water. Simultaneously, it could specifically detect Cd(II). Nontarget metal ions, such as Zn(II), Hg(II), and Pb(II), did not affect its performance. Furthermore, it exhibited superior performance in detecting Cd(II) in real drinking water samples by avoiding background interference, and showed excellent stability with the relative standard deviation under 5%. Thus, K12-PMP-luxCDABE-△cysI holds promise as a potential tool for the detection of Cd(II) in drinking water.
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High-Throughput Screening to Identify Novel Compounds Affecting the Genome Editing Efficiency of CRISPR System.
Genome editing is a promising therapeutic strategy for genetic disorders by modifying the genome precisely, especially the CRISPR/Cas9 system. However, a major limitation of CRISPR/Cas9 in gene therapy is the biosafety issues caused by off-target effects. Compounds that can modulate the genome editing efficiency of the CRISPR/Cas9 system, especially those reducing the off-target effects, are potentially useful pharmacological tools for improving the effectiveness and safety of genome editing. Here, we performed high-throughput screening in HEK 293FT cells to discover compounds that decrease or increase the genome editing efficiency of the CRISPR/Cas9 system from 9930 compounds. After two rounds of screening, we identified that CP-724714, a ErbB2 (HER2) tyrosine kinase inhibitor, decreased the CRISPR/Cas9 efficiency and reduced the off-target effects by suppressing the efficiency of CRISPR/Cas9, and was thus named a CRISPR decelerator (or inhibitor), while Clofarabine, a DNA synthesis inhibitor, increased the efficiency of CRISPR/Cas9, and was named a CRISPR accelerator. We further identified four compounds (Tranilast, Cerulenin, Rosolic acid and Resveratrol) that affected the efficiency of single-strand annealing (SSA) repair. Among them, Tranilast, Cerulenin and Rosolic acid are potential SSA decelerators, while Resveratrol is a potential SSA accelerator. These identified compounds may be useful in optimizing mammalian genetic manipulation techniques.
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Elimination of HIV-1 Genomes from Human T-lymphoid Cells by CRISPR/Cas9 Gene Editing.
We employed an RNA-guided CRISPR/Cas9 DNA editing system to precisely remove the entire HIV-1 genome spanning between 5' and 3' LTRs of integrated HIV-1 proviral DNA copies from latently infected human CD4+ T-cells. Comprehensive assessment of whole-genome sequencing of HIV-1 eradicated cells ruled out any off-target effects by our CRISPR/Cas9 technology that might compromise the integrity of the host genome and further showed no effect on several cell health indices including viability, cell cycle and apoptosis. Persistent co-expression of Cas9 and the specific targeting guide RNAs in HIV-1-eradicated T-cells protected them against new infection by HIV-1. Lentivirus-delivered CRISPR/Cas9 significantly diminished HIV-1 replication in infected primary CD4+ T-cell cultures and drastically reduced viral load in ex vivo culture of CD4+ T-cells obtained from HIV-1 infected patients. Thus, gene editing using CRISPR/Cas9 may provide a new therapeutic path for eliminating HIV-1 DNA from CD4+ T-cells and potentially serve as a novel and effective platform toward curing AIDS.
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Peptide display on small extracellular vesicles directs tissue-specific tropism and delivery of gene editing machinery.
Small extracellular vesicles (EVs) deliver nucleic acid and protein therapeutics that promote tissue repair, however, there are few approaches to direct the tropism of EVs to specific tissues. Here, we address this challenge by engineering EVs to display a peptide library on the surface of EVs that is linked to barcoded guide RNA payloads (gRNAs). We show how these EVs can be administered systemically into mouse models and the cellular uptake of these gRNA-bearing EVs assessed by recovery of the small RNAs followed by PCR amplification and barcode sequencing. Since the design of the peptide library was linked to specific barcodes encoded on the same plasmid, subsequent sequencing of barcode sequences recovered from tissues revealed profiles of EV uptake associated with the display of specific peptide sequences on the surface of EVs. Therefore, candidate peptide motifs were cloned for validation using in vivo and in vitro readouts. In addition to the barcode-based tissue profiling, gRNA-laded EVs mediated functional gene editing in the Cre-loxP R26 LSL-tdTomato reporter mouse model in vivo. These studies demonstrated how EV display linked with barcoded gRNA payloads can address barriers to EV-based delivery of gene editing therapies.
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Synthetic minichromosomes in plants: past, present, and promise.
The status of engineered mini-chromosomes/artificial chromosomes/synthetic chromosomes in plants is summarized. Their promise is that they provide a means to accumulate foreign genes on an independent entity other than the normal chromosomes, which would facilitate stacking of novel traits in a way that would not be linked to endogenous genes and that would facilitate transfer between lines. Centromeres in plants are epigenetic, and therefore the isolation of DNA underlying centromeres and reintroduction into plant cells will not establish a functional kinetochore, which obviates this approach for in vitro assembly of plant artificial chromosomes. This issue was bypassed by using telomere-mediated chromosomal truncation to produce mini-chromosomes with little more than an endogenous centromere that could in turn be used as a foundation to build synthetic chromosomes. Site-specific recombinases and various iterations of CRISPR-Cas9 editing provide many tools for the development and re-engineering of synthetic chromosomes.
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Efficient genome editing by CRISPR/Cas9 with a tRNA-sgRNA fusion in the methylotrophic yeast Ogataea polymorpha.
The methylotrophic yeast Ogataea polymorpha (syn. Hansenula polymorpha) is an attractive industrial non-conventional yeast showing high thermo-tolerance (up to 50°C) and xylose assimilation. However, genetic manipulation of O. polymorpha is often laborious and time-consuming because it has lower homologous recombination efficiency relative to Saccharomyces cerevisiae. To overcome this disadvantage, we applied the CRISPR/Cas9 system as a powerful genome editing tool in O. polymorpha. In this system, both single guide RNA (sgRNA) and endonuclease Cas9 were expressed by a single autonomously-replicable plasmid and the sgRNA portion could be easily changed by using PCR and In-Fusion cloning techniques. Because the mutation efficiency of the CRISPR/Cas9 system was relatively low when the sgRNA was expressed under the control of the OpSNR6 promoter, the tRNACUG gene was used for sgRNA expression. The editing efficiency of this system ranged from 17% to 71% of transformants in several target genes tested (ADE12, PHO1, PHO11, and PHO84). These findings indicate that genetic manipulation of O. polymorpha will be more convenient and accelerated by using this CRISPR/Cas9 system.
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Applications and explorations of CRISPR/Cas9 in CAR T-cell therapy.
Chimeric antigen receptor(CAR) T-cell therapy has shown remarkable effects and promising prospects in patients with refractory or relapsed malignancies, pending further progress in the next-generation CAR T cells with more optimized structure, enhanced efficacy and reduced toxicities. The clustered regulatory interspaced short palindromic repeat/CRISPR-associated protein 9 (CRISPR/Cas9) technology holds immense promise for advancing the field owing to its flexibility, simplicity, high efficiency and multiplexing in precise genome editing. Herein, we review the applications and explorations of CRISPR/Cas9 technology in constructing allogenic universal CAR T cells, disrupting inhibitory signaling to enhance potency and exploration of safer and more controllable novel CAR T cells.
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Electroporation-mediated genome editing in vitrified/warmed mouse zygotes created by IVF via ultra-superovulation.
Recently, genome editing in mouse zygotes has become convenient and scalable, in association with various technological developments and improvements such as novel nuclease tools, alternative delivery methods, and contemporary reproductive engineering techniques. We have so far demonstrated the applicability of ultra-superovulation, in vitro fertilization (IVF), and vitrification/warming of zygotes in microinjection-mediated mouse genome editing. Moreover, an electroporation-mediated method has rapidly become established for simple gene knockout and small precise modifications including single amino acid substitutions. Here, we present an updated example of an application coupling the following three latest technologies: 1) CRISPR-Cas9 ribonucleoprotein as the most convenient genome-editing reagent, 2) electroporation as the most effortless delivery method, and 3) cryopreserved oocytes created by IVF via ultra-superovulation as the most animal welfare- and user-friendly strategy. We successfully created gene knockout and knock-in mice carrying insertion/deletion mutations and single amino acid substitutions, respectively, using the streamlined production system of mouse genome editing described above, referred to as the CREATRE (CARD-based Reproductive Engineering-Assisted Technology for RNP Electroporation) system. Owing to its accessibility, robustness, and high efficiency, we believe that our CREATRE protocol will become widely used globally for the production of genome-edited mice.
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