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[Gene editing in drug discovery and therapeutic innovation].
The idea according to which the most recent therapeutic methods will overcome the more traditional pharmacopoeia is widespread in recent publications. Biomedicine and gene therapies are booming, but we realize, as for other therapeutic approaches, that they suffer intrinsic constraints and limitations and that their most relevant therapeutic fields are complementary to those of traditional drugs. They are now viewed as potentially synergistic with these traditional drugs, rather than competitors. This review puts into perspective the potential of genome editing in the field of drug discovery and therapeutic innovation.
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Precision editing in the human embryo.
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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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CRISPR/Cas9 as a Genome Editing Tool for Targeted Gene Integration in CHO Cells.
The emergence of CRISPR/Cas9 system as a precise and affordable method for genome editing has prompted its rapid adoption for the targeted integration of transgenes in Chinese hamster ovary (CHO) cells. Targeted gene integration allows the generation of stable cell lines with a controlled and predictable behavior, which is an important feature for the rational design of cell factories aimed at the large-scale production of recombinant proteins. Here we present the protocol for CRISPR/Cas9-mediated integration of a gene expression cassette into a specific genomic locus in CHO cells using homology-directed DNA repair.
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Mild phenotype of knockouts of the major apurinic/apyrimidinic endonuclease APEX1 in a non-cancer human cell line.
The major human apurinic/apyrimidinic (AP) site endonuclease, APEX1, is a central player in the base excision DNA repair (BER) pathway and has a role in the regulation of DNA binding by transcription factors. In vertebrates, APEX1 knockouts are embryonic lethal, and only a handful of knockout cell lines are known. To facilitate studies of multiple functions of this protein in human cells, we have used the CRISPR/Cas9 system to knock out the APEX1 gene in a widely used non-cancer hypotriploid HEK 293FT cell line. Two stable knockout lines were obtained, one carrying two single-base deletion alleles and one single-base insertion allele in exon 3, another homozygous in the single-base insertion allele. Both mutations cause a frameshift that leads to premature translation termination before the start of the protein's catalytic domain. Both cell lines totally lacked the APEX1 protein and AP site-cleaving activity, and showed significantly lower levels of the APEX1 transcript. The APEX1-null cells were unable to support BER on uracil- or AP site-containing substrates. Phenotypically, they showed a moderately increased sensitivity to methyl methanesulfonate (MMS; ~2-fold lower EC50 compared with wild-type cells), and their background level of natural AP sites detected by the aldehyde-reactive probe was elevated ~1.5-2-fold. However, the knockout lines retained a nearly wild-type sensitivity to oxidizing agents hydrogen peroxide and potassium bromate. Interestingly, despite the increased MMS cytotoxicity, we observed no additional increase in AP sites in knockout cells upon MMS treatment, which could indicate their conversion into more toxic products in the absence of repair. Overall, the relatively mild cell phenotype in the absence of APEX1-dependent BER suggests that mammalian cells possess mechanisms of tolerance or alternative repair of AP sites. The knockout derivatives of the extensively characterized HEK 293FT cell line may provide a valuable tool for studies of APEX1 in DNA repair and beyond.
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CRISPR/Cas9 and Chlorophyll Coordination Micelles for Cancer Treatment by Genome Editing and Photodynamic Therapy.
CRISPR/Cas9-based gene therapy and photodynamic therapy both show promise for cancer treatment but still have their drawbacks limited by tumor microenvironment and long treatment duration. Herein, CRISPR/Cas9 genome editing and photodynamic strategy for a synergistic anti-tumor therapeutic modality is merged. Chlorophyll (Chl) extracted from natural green vegetables is encapsulated in Pluronic F127 (F127) micelles and Histidine-tagged Cas9 can be effectively chelated onto micelles via metal coordination by simple incubation, affording Cas9-Chl@F127 micelles. Mg2+ acts as an enzyme cofactor to correlatively enhance Cas9 gene-editing activity. Upon laser irradiation, Chl as an effective photosensitizer generates reactive oxygen species (ROS) to kill tumor cells. Meanwhile, CRISPR/Cas9, mediated by dual deliberately designed gRNAs of APE1 and NRF2, can reprogram the tumor microenvironment by increasing the intracellular oxygen accumulation and impairing the oxidative defense system of tumor cells. Cas9-Chl@F127 micelles can responsively release Cas9 in the presence of abundant ATP or low pH in tumor cells. In a murine tumor model, Cas9-Chl@F127 complexed with dual gRNAs including APE1 and NRF2 significantly inhibits the tumor growth. Taken together, Cas9-Chl@F127 micelles, representing the first Chl-based green biomaterial for the delivery of Cas9, show great promise for the synergistic anti-tumor treatment by PDT and gene editing.
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Efficient genetic perturbation of murine sensory neurons in vivo using CRISPR/Cas9.
Gene editing using CRISPR/Cas9 in vivo offers a powerful tool to investigate pain mechanisms. We generated a conditional knock-in mouse model where Streptococcus pyogenes CRISPR-associated protein 9 (Cas9) expression is restricted to cells that express SCN9A. Transgenic markers were detected in key tissues including the dorsal root ganglia (DRG) and sciatic nerve. To assess in vivo editing efficacy, RNA guides targeting TRPV1 were intrathecally administered. Two injections of guide RNAs resulted in a significant reduction of TRPV1 in both the DRG and sciatic nerve without triggering caspase-3-mediated apoptosis or motor deficits. Edited animals exhibited increased withdrawal latencies to heat and reduced nocifensive behaviors following capsaicin injection. Capsaicin-evoked thermal hyperalgesia and mechanical allodynia were diminished. This approach enables rapid and efficient sensory neuron-specific CRISPR/Cas9 gene perturbations for pain research in mice. We envisage that this method can be employed both for the exploration of molecular mechanisms underlying nociception and for the validation of therapeutic targets associated with pain. PERSPECTIVE: There are tremendous opportunities afforded by facile multi-locus genome perturbation of sensory neurons in vivo. This model and approach enables rapid and low-cost genetic depletion experiments in mice.
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Learning from the Past: Discussing Lessons from Reproductive Justice in the Gene-Editing Sphere.
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Implementing CRISPR-Cas9 Yeast Practicals into Biology Curricula.
CRISPR-Cas9 is a genome-editing technique that has been widely adopted thanks to its simplicity, efficiency, and broad application potential. Due to its advantages and pervasive use, there have been attempts to include this method in the existing curricula for students majoring in various disciplines of biology. In this perspective, we summarize the existing CRISPR-Cas courses that harness a well-established model organism: baker's yeast, Saccharomyces cerevisiae. As an example, we present a detailed description of a fully hands-on, flexible, robust, and cost-efficient practical CRISPR-Cas9 course, where students participate in yeast genome editing at every stage-from the bioinformatic design of single-guide RNA, through molecular cloning and yeast transformation, to the final confirmation of the introduced mutation. Finally, we emphasize that in addition to providing experimental skills and theoretical knowledge, the practical courses on CRISPR-Cas represent ideal platforms for discussing the ethical implications of the democratization of biology.
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