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A boosting upconversion luminescent resonance energy transfer and biomimetic periodic chip integrated CRISPR/Cas12a biosensor for functional DNA regulated transduction of non-nucleic acid targets.
Apart from gene editing capacity, the newly discovered CRISPR/Cas systems offer an exciting option for biosensing field because of their excellent target recognition accuracy. However, the currently constructed sensors are not only limited to nucleic acid analysis but also suffer from poor adaptability in complex samples and unsatisfying sensitivity. We herein introduce some advanced concepts to break through these bottlenecks. First, the sensing targets are extended by skillfully designing a functional DNA such as aptamer (for protein) and DNAzyme (for metal ion) to regulate the transduction of non-nucleic acid species and further activate the trans cleavage of CRISPR/Cas12a. Second, a boosting upconversion luminescent resonance energy is triggered by using a peculiar energy-confining notion, whereby the luminescence domain is intensively restricted in a very narrow space (~2.44 nm) and up to 92.9% of the green emission can be quenched by the approaching BHQ-1 modified reporters. Third, a bio-inspired periodic arrangement biomimetic chip (photonic crystal) is employed to selectively reflect the upconversion luminescence to achieve noteworthy signal enhancement (~35-fold). By utilizing very simple detection devices (a 980 nm portable laser and a smartphone), the CRISPR/Cas12a biosensor shows commendable sensitivity and specificity toward model targets (ATP and Na+, limits of detection are ~ 18 nM and ~0.37 μM, respectively). More importantly, the analysis of real complex samples demonstrate that the as-proposed platform can work as a powerful toolbox for monitoring the ATP fluctuation in single cell and point-of-care testing Na+ in human plasma, enabling a broad application prospect.
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Generating CRISPR/Cas9 Mediated Monoallelic Deletions to Study Enhancer Function in Mouse Embryonic Stem Cells.
Enhancers control cell identity by regulating tissue-specific gene expression in a position and orientation independent manner. These enhancers are often located distally from the regulated gene in intergenic regions or even within the body of another gene. The position independent nature of enhancer activity makes it difficult to match enhancers with the genes they regulate. Deletion of an enhancer region provides direct evidence for enhancer activity and is the gold standard to reveal an enhancer's role in endogenous gene transcription. Conventional homologous recombination based deletion methods have been surpassed by recent advances in genome editing technology which enable rapid and precisely located changes to the genomes of numerous model organisms. CRISPR/Cas9 mediated genome editing can be used to manipulate the genome in many cell types and organisms rapidly and cost effectively, due to the ease with which Cas9 can be targeted to the genome by a guide RNA from a bespoke expression plasmid. Homozygous deletion of essential gene regulatory elements might lead to lethality or alter cellular phenotype whereas monoallelic deletion of transcriptional enhancers allows for the study of cis-regulation of gene expression without this confounding issue. Presented here is a protocol for CRISPR/Cas9 mediated deletion in F1 mouse embryonic stem (ES) cells (Mus musculus(129) x Mus castaneus). Monoallelic deletion, screening and expression analysis is facilitated by single nucleotide polymorphisms (SNP) between the two alleles which occur on average every 125 bp in these cells.
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DNA-binding-domain fusions enhance the targeting range and precision of Cas9.
The CRISPR-Cas9 system is commonly used in biomedical research; however, the precision of Cas9 is suboptimal for applications that involve editing a large population of cells (for example, gene therapy). Variations on the standard Cas9 system have yielded improvements in the precision of targeted DNA cleavage, but they often restrict the range of targetable sequences. It remains unclear whether these variants can limit lesions to a single site in the human genome over a large cohort of treated cells. Here we show that by fusing a programmable DNA-binding domain (pDBD) to Cas9 and attenuating Cas9's inherent DNA-binding affinity, we were able to produce a Cas9-pDBD chimera with dramatically improved precision and an increased targeting range. Because the specificity and affinity of this framework can be easily tuned, Cas9-pDBDs provide a flexible system that can be tailored to achieve extremely precise genome editing at nearly any genomic locus.
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A modified pCas/pTargetF system for CRISPR-Cas9-assisted genome editing in Escherichia coli.
The clustered regularly interspaced short palindromic repeats (CRISPR)-associated nuclease 9 (Cas9)-based genome editing tool pCas/pTargetF system that we established previously has been widely used in Escherichia coli MG1655. However, this system failed to manipulate the genome of E. coli BL21(DE3), owing to the potential higher leaky transcription of the gRNA-pMB1 specific to pTargetF in this strain. In this study, we modified the pCas/pTargetF system by replacing the promoter of gRNA-pMB1 with a tightly regulated promoter PrhaB, changing the replicon of pCas to a nontemperature-sensitive replicon, adding the sacB gene into pCas, and replacing the original N20-specific sequence of pTargetF with ccdB gene. We call this updated system as pEcCas/pEcgRNA. We found that gRNA-pMB1 indeed showed a slightly higher leaky expression in the pCas/pTargetF system compared with pEcCas/pEcgRNA. We also confirmed that genome editing can successfully be performed in BL21(DE3) by pEcCas/pEcgRNA with high efficiency. The application of pEcCas/pEcgRNA was then expanded to the E. coli B strain BL21 StarTM (DE3), K-12 strains MG1655, DH5α, CGMCC3705, Nissle1917, W strain ATCC9637, and also another species of Enterobacteriaceae, Tatumella citrea DSM13699, without any specific modifications. Finally, the plasmid curing process was optimized to shorten the time from $\sim$60 h to $\sim$32 h. The entire protocol (including plasmid construction, editing, electroporation and mutant verification, and plasmid elimination) took only $\sim$5.5 days per round in the pEcCas/pEcgRNA system, whereas it took $\sim$7.5 days in the pCas/pTargetF system. This study established a faster-acting genome editing tool that can be used in a wider range of E. coli strains and will also be useful for other Enterobacteriaceae species.
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In vivo editing of the pan-endothelium by immunity evading simian adenoviral vector.
Biological applications deriving from the clustered regularly interspaced short palindromic repeats (CRISPR)-Cas9 site-specific nuclease system continue to impact and accelerate gene therapy strategies. Safe and effective in vivo co-delivery of the CRISPR/Cas9 system to target somatic cells is essential in the clinical therapeutic context. Both non-viral and viral vector systems have been applied for this delivery matter. Despite elegant proof-of-principle studies, available vector technologies still face challenges that restrict the application of CRISPR/Cas9-facilitated gene therapy. Of note, the mandated co-delivery of the gene-editing components must be accomplished in the potential presence of pre-formed anti-vector immunity. Additionally, methods must be sought to limit the potential of off-target editing. To this end, we have exploited the molecular promiscuities of adenovirus (Ad) to address the key requirements of CRISPR/Cas9-facilitated gene therapy. In this regard, we have endeavored capsid engineering of a simian (chimpanzee) adenovirus isolate 36 (SAd36) to achieve targeted modifications of vector tropism. The SAd36 vector with the myeloid cell-binding peptide (MBP) incorporated in the capsid has allowed selective in vivo modifications of the vascular endothelium. Importantly, vascular endothelium can serve as an effective non-hepatic cellular source of deficient serum factors relevant to several inherited genetic disorders. In addition to allowing for re-directed tropism, capsid engineering of nonhuman primate Ads provide the means to circumvent pre-formed vector immunity. Herein we have generated a SAd36. MBP vector that can serve as a single intravenously administered agent allowing effective and selective in vivo editing for endothelial target cells of the mouse spleen, brain and kidney. DATA AVAILABILITY: The data that support the findings of this study are available from the corresponding author upon reasonable request.
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Applying CRISPR-Cas9 screens to dissect hematological malignancies.
Bit by bit, over the last few decades, functional genomic tools have been piecing together the molecular puzzle driving tumorigenesis in human patients. Nevertheless, our understanding of the role of several genes and regulatory elements that drive critical cancer-associated physiological processes from disease development to progression to spread is very limited, which significantly affects our ability of applying these insights in the context of improved disease management. The recent advent of clustered regularly interspaced short palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9)-based technology and its application in cancer genomics has, however, allowed the generation of a wealth of knowledge that has helped decipher several critical questions associated with translational cancer research. Precisely, the high-throughput capability coupled with a high level of technological plasticity associated with the CRISPR-Cas9 screens have expanded our horizons from a mere struggle to appreciate cancer as a genetic disease to observing the integrated genomic/epigenomic network of numerous malignancies and correlating it with our present knowledge of drugging strategies to develop innovative approaches for next-generation precision cancer medicine. Specifically, within blood cancers, current CRISPR screens have specifically focused on improving our understanding of drug resistance mechanisms, disease biology, the development of novel therapeutic approaches, and identifying the molecular mechanisms of current therapies, with an underlying aim of improving disease outcomes. Here, we review the development of the CRISPR-Cas9 genome-editing strategy, explicitly focusing on the recent advances in the CRISPR-Cas9-based screening approaches, its current capabilities, limitations, and future applications in the context of hematological malignancies.
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Exploring the potential of CRISPR/Cas genome editing for vegetable crop improvement: An overview of challenges and approaches.
Vegetables provide many nutrients in the form of fiber, vitamins, and minerals, which make them an important part of our diet. Numerous biotic and abiotic stresses can affect crop growth, quality, and yield. Traditional and modern breeding strategies to improve plant traits are slow and resource intensive. Therefore, it is necessary to find new approaches for crop improvement. Clustered regularly interspaced short palindromic repeats/CRISPR associated 9 (CRISPR/Cas9) is a genome editing tool that can be used to modify targeted genes for desirable traits with greater efficiency and accuracy. By using CRISPR/Cas9 editing to precisely mutate key genes, it is possible to rapidly generate new germplasm resources for the promotion of important agronomic traits. This is made possible by the availability of whole genome sequencing data and information on the function of genes responsible for important traits. In addition, CRISPR/Cas9 systems have revolutionized agriculture, making genome editing more versatile. Currently, genome editing of vegetable crops is limited to a few vegetable varieties (tomato, sweet potato, potato, carrot, squash, eggplant, etc.) due to lack of regeneration protocols and sufficient genome sequencing data. In this article, we summarize recent studies on the application of CRISPR/Cas9 in improving vegetable trait development and the potential for future improvement.
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CRISPR prime editing for unconstrained correction of oncogenic KRAS variants.
KRAS is the most commonly mutated RAS family gene and is a primary cause of the occurrence of several types of cancer. However, KRAS mutations have several unique and diverse molecular identities, making it difficult to find specific treatments. Here, we developed universal pegRNAs which can correct all types of G12 and G13 oncogenic KRAS mutations with CRISPR-mediated prime editors (PEs). The universal pegRNA successfully corrected 12 types of KRAS mutations, accounting for 94% of all known KRAS mutations, by up to 54.8% correction frequency in HEK293T/17 cells. We also applied the universal pegRNA to correct endogenous KRAS mutations in human cancer cells and found that G13D KRAS mutation was successfully corrected to wild-type KRAS sequences with up to 40.6% correction frequency without indel mutations. We propose prime editing with the universal pegRNA as a 'one-to-many' potential therapeutic strategy for KRAS oncogene variants.
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Genome editing using FACS enrichment of nuclease-expressing cells and indel detection by amplicon analysis.
This protocol describes methods for increasing and evaluating the efficiency of genome editing based on the CRISPR-Cas9 (clustered regularly interspaced short palindromic repeats-CRISPR-associated 9) system, transcription activator-like effector nucleases (TALENs) or zinc-finger nucleases (ZFNs). First, Indel Detection by Amplicon Analysis (IDAA) determines the size and frequency of insertions and deletions elicited by nucleases in cells, tissues or embryos through analysis of fluorophore-labeled PCR amplicons covering the nuclease target site by capillary electrophoresis in a sequenator. Second, FACS enrichment of cells expressing nucleases linked to fluorescent proteins can be used to maximize knockout or knock-in editing efficiencies or to balance editing efficiency and toxic/off-target effects. The two methods can be combined to form a pipeline for cell-line editing that facilitates the testing of new nuclease reagents and the generation of edited cell pools or clonal cell lines, reducing the number of clones that need to be generated and increasing the ease with which they are screened. The pipeline shortens the time line, but it most prominently reduces the workload of cell-line editing, which may be completed within 4 weeks.
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Combined refinements to somatic cell nuclear transfer methods improve porcine embryo development.
The discovery of how to utilize CRISPR (clustered, regularly interspaced, short, palindromic repeats)-Cas (CRISPR-associated) systems for genome modification has accelerated development of the field of genome editing, especially in large animals such as pigs. The low efficiency of somatic cell nuclear transfer (SCNT) is now becoming a major obstacle in the production of genome-edited animals via cell-mediated approaches and improving efficacy of this technique is crucial. In this study, we propose a few simple modifications to a zona-free SCNT protocol that are effective to produce numerous high-quality blastocysts. To refine the SCNT protocol we modified the following steps/factors: 1) culture medium for SCNT embryos, 2) chemical treatment to prevent precocious activation of the manipulated/reconstructed oocytes and 3) donor cell serum starvation treatment. Although changes in each of these steps only resulted in small improvements, the combination of all modifications altogether significantly enhanced developmental competence of SCNT embryos. Our modified method yielded approximately three times greater blastocyst formation rates. Moreover, resulting blastocysts had roughly twice as many cells as compared to blastocysts produced by the conventional SCNT method. With these significant in vitro improvements, our refined SCNT method is potentially suited for use in the production of genome edited pigs.
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