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In Vivo CRISPR/Cas9-Based Targeted Disruption and Knockin of a Long Noncoding RNA.
The CRISPR/Cas9 system has been widely used as an efficient genome-editing tool for studying physiological functions of long noncoding RNAs (lncRNAs). In this chapter, we describe the experimental procedures for using the CRISPR/Cas9 system to genetically modify a long noncoding RNA in vivo through the targeted disruption and knockin approaches.
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First report on CRISPR/Cas9-targeted mutagenesis in the Colorado potato beetle, Leptinotarsa decemlineata.
Leptinotarsa decemlineata (Say), commonly known as the Colorado potato beetle (CPB), is an agricultural important pest for potatoes and other solanaceous plants. The CRISPR/Cas system is an efficient genome editing technology, which could be exploited to study the biology of CPB and possibly also lead to the development of better environmentally friendly pest management strategies. However, the use of CRISPR/Cas9 has been limited to only a few model insects. Here, for the first time, a CRISPR/Cas9 protocol for mutagenesis studies in CPB was developed. A gene with a clear phenotype such as the vestigial gene (vest), known to be involved in wing development in other insect species, was selected as a good indicator for the knockout study. First, vest was functionally characterized in CPB by using RNAi technology for knockdown studies. Once the expected deformed wing phenotypes were observed, a CRISPR/Cas9 work flow was established for mutagenesis in CPB. By co-injecting the Cas9 protein and a vest-guide RNA into 539 CPB eggs of <1 h old, sixty-two successfully developed to adults, among which mutation in the vest loci was confirmed in 5 of the 18 wingless CPBs (29% phenotypic mutation efficiency). The mutation in vest resulted in a clear phenotype in the CPBs, which developed to adulthood with no hindwing and elytron formed. Altogether, this study provides for the first time a useful methodology involving the use of the CRISPR/Cas9 system for mutagenesis studies in one of the most important pest insects.
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A novel gene editing lexicon strategy for the haemophilia community: Research plan for development and preliminary results.
INTRODUCTION: Despite the progress in gene editing platforms like CRISPR/Cas9 with the potential to transform the standard of care for haemophilia, the language used to explain and discuss gene editing is not aligned across the haemophilia community. Here, we present the objective and rationale for developing a clear, consistent, and globally aligned gene editing lexicon to address these communication gaps. METHODS: Effectively communicating complex gene editing concepts requires a clear and consistent vocabulary. Through collaboration with a diversity of haemophilia stakeholders, our main goal is to develop an accurate, informative lexicon which avoids overpromising or highly technical terminology. Using an innovative process, representatives from several patient and scientific haemophilia organizations and select biotechnology companies will develop and refine language concepts to be tested with approximately seventy participants across the United States of America, United Kingdom, and Germany. Participants will include lived experience experts (LEEs) and haematologists. The process will be overseen by the Lexicon Steering Committee of global experts from leading scientific and patient organizations in the haemophilia and gene editing fields. RESULTS: Initial feedback provided a robust foundation and rationale for building clear, consistent language around gene editing. This lexicon development framework will allow for increased understanding across the haemophilia community, including the development of valid informed consent and shared decision-making materials. CONCLUSION: Results provide important building blocks for stimuli development and highlight the need for a novel gene editing lexicon. In the next phase, language stimuli will be tested with LEEs and haematologists to better understand audience preferences and help shape the final lexicon.
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Efficient DNA-free co-targeting of nuclear genes in Chlamydomonas reinhardtii.
Chlamydomonas reinhardtii, a model organism for unicellular green microalgae, is widely used in basic and applied research. Nonetheless, proceeding towards synthetic biology requires a full set of manipulation techniques for inserting, removing, or editing genes. Despite recent advancements in CRISPR/Cas9, still significant limitations in producing gene knock-outs are standing, including (i) unsatisfactory genome editing (GE) efficiency and (ii) uncontrolled DNA random insertion of antibiotic resistance markers. Thus, obtaining efficient gene targeting without using marker genes is instrumental in developing a pipeline for efficient engineering of strains for biotechnological applications. We developed an efficient DNA-free gene disruption strategy, relying on phenotypical identification of mutants, to (i) precisely determine its efficiency compared to marker-relying approaches and (ii) establish a new DNA-free editing tool. This study found that classical CRISPR Cas9-based GE for gene disruption in Chlamydomonas reinhardtii is mainly limited by DNA integration. With respect to previous results achieved on synchronized cell populations, we succeeded in increasing the GE efficiency of single gene targeting by about 200 times and up to 270 times by applying phosphate starvation. Moreover, we determined the efficiency of multiplex simultaneous gene disruption by using an additional gene target whose knock-out did not lead to a visible phenotype, achieving a co-targeting efficiency of 22%. These results expand the toolset of GE techniques and, additionally, lead the way to future strategies to generate complex genotypes or to functionally investigate gene families. Furthermore, the approach provides new perspectives on how GE can be applied to (non-) model microalgae species, targeting groups of candidate genes of high interest for basic research and biotechnological applications.
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Efficient Separation of Single-Stranded DNA of Various Lengths for Applications in Bioimaging, Gene Editing, and Nanostructures.
Single-stranded DNA (ssDNA) has extremely high design flexibility and specific functions. Therefore, ssDNA is used in crucial practical applications in many fields, such as molecular detection, gene editing, and nanotechnology. However, the existing methods for ssDNA preparation often present limitations in terms of yield, purity, and length applicability. In order to overcome these challenges, the present study proposes a ssDNA separation method that utilizes the modification of linear polyacrylamide (LPA) combined with denaturing agarose gel electrophoresis (DAGE). The method is referred to as LPA-DAGE. The underlying principle is to produce LPA-modified double-stranded DNA (dsDNA) through PCR using the LPA primer and then achieve efficient ssDNA separation using denaturing agarose gel electrophoresis based on molecular weight differences. This method can stably recover ssDNA showing significant advantages over the existing methods in terms of purity and recovery rate. The results of this study demonstrate that the proposed ssDNA preparation method enables signal amplification using fluorescence in situ hybridization (FISH), CRISPR/Cas9-mediated homologous recombination repair, and nanostructure construction using DNA origami.
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De Novo Synthesis of Reticuline and Taxifolin Using Re-engineered Homologous Recombination in Yarrowia lipolytica.
Yarrowia lipolytica has been widely engineered as a eukaryotic cell factory to produce various important compounds. However, the difficulty of gene editing and the lack of efficient neutral sites make rewiring of Y. lipolytica metabolism challenging. Herein, a Cas9 system was established to redesign the Y. lipolytica homologous recombination system, which caused a more than 56-fold increase in the HR efficiency. The fusion expression of the hBrex27 sequence in the C-terminus of Cas9 recruited more Rad51 protein, and the engineered Cas9 decreased NHEJ, achieving 85% single-gene positive efficiency and 25% multigene editing efficiency. With this system, neutral sites on different chromosomes were characterized, and a deep learning model was developed for gRNA activity prediction, thus providing the corresponding integration efficiency and expression intensity. Subsequently, the tool and platform strains were validated by applying them for the de novo synthesis of (S)-reticuline and (2S)-taxifolin. The developed platform strains and tools helped transform Y. lipolytica into an easy-to-operate model cell factory, similar to Saccharomyces cerevisiae.
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Reversible perturbations of gene regulation after genome editing in Drosophila cells.
The prokaryotic phage defense CRISPR/cas-system has developed into a versatile toolbox for genome engineering and genetic studies in many organisms. While many efforts were spent on analyzing the consequences of off-target effects, only few studies addressed side-effects that occur due to the targeted manipulation of the genome. Here, we show that the CRISPR/cas9-mediated integration of an epitope tag in combination with a selection cassette can trigger an siRNA-mediated, epigenetic genome surveillance pathway in Drosophila melanogaster cells. After homology-directed insertion of the sequence coding for the epitope tag and the selection marker, a moderate level of siRNAs covering the inserted sequence and extending into the targeted locus was detected. This response affected protein levels less than two-fold and it persisted even after single cell cloning. However, removal of the selection cassette abolished the siRNA generation, demonstrating that this response is reversible. Consistently, marker-free genome engineering did not trigger the same surveillance mechanism. These two observations indicate that the selection cassette we employed induces an aberrant transcriptional arrangement and ultimately sets off the siRNA production. There have been prior concerns about undesirable effects induced by selection markers, but fortunately we were able to show that at least one of the epigenetic changes reverts as the marker gene is excised. Although the effects observed were rather weak (less than twofold de-repression upon ago2 or dcr-2 knock-down), we recommend that when selection markers are used during genome editing, a strategy for their subsequent removal should always be included.
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CRISPR/Cas System for Genome Editing: Progress and Prospects as a Therapeutic Tool.
CRISPR was first observed in 1987 in bacteria and archaea and was later confirmed as part of bacterial adaptive immunity against the attacking phage. The CRISPR/Cas restriction system involves a restriction endonuclease enzyme guided by a hybrid strand of RNA consisting of CRISPR RNA and trans-activating RNA, which results in gene knockout or knockin followed by nonhomologous end joining and homology-directed repair. Owing to its efficiency, specificity, and reproducibility, the CRISPR/Cas restriction system was said to be a breakthrough in the field of biotechnology. Apart from its application in biotechnology, CRISPR/Cas has been explored for its therapeutic potential in several diseases including cancer, Alzheimer's disease, sickle cell disease, Duchenne muscular dystrophy, neurologic disorders, etc., wherein CRISPR/Cas components such as Cas9/single guide RNA (sgRNA) ribonucleoprotein, sgRNA/mRNA, and plasmid were delivered. However, limitations including immunogenicity, low transfection, limited payload, instability, and off-target binding pose hurdles in its therapeutic use. Nonviral vectors (including cationic polymers, lipids, etc.), classically used as carriers for therapeutic genes, were used to deliver CRISPR/Cas components and showed interesting results. Herein, we discuss the CRISPR/Cas system and its brief history and classification, followed by its therapeutic applications using current nonviral delivery strategies.
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Human genetic diversity alters off-target outcomes of therapeutic gene editing.
CRISPR gene editing holds great promise to modify DNA sequences in somatic cells to treat disease. However, standard computational and biochemical methods to predict off-target potential focus on reference genomes. We developed an efficient tool called CRISPRme that considers single-nucleotide polymorphism (SNP) and indel genetic variants to nominate and prioritize off-target sites. We tested the software with a BCL11A enhancer targeting guide RNA (gRNA) showing promise in clinical trials for sickle cell disease and β-thalassemia and found that the top candidate off-target is produced by an allele common in African-ancestry populations (MAF 4.5%) that introduces a protospacer adjacent motif (PAM) sequence. We validated that SpCas9 generates strictly allele-specific indels and pericentric inversions in CD34+ hematopoietic stem and progenitor cells (HSPCs), although high-fidelity Cas9 mitigates this off-target. This report illustrates how genetic variants should be considered as modifiers of gene editing outcomes. We expect that variant-aware off-target assessment will become integral to therapeutic genome editing evaluation and provide a powerful approach for comprehensive off-target nomination.
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Mutation-specific reporter for optimization and enrichment of prime editing.
Prime editing is a versatile genome-editing technique that shows great promise for the generation and repair of patient mutations. However, some genomic sites are difficult to edit and optimal design of prime-editing tools remains elusive. Here we present a fluorescent prime editing and enrichment reporter (fluoPEER), which can be tailored to any genomic target site. This system rapidly and faithfully ranks the efficiency of prime edit guide RNAs (pegRNAs) combined with any prime editor variant. We apply fluoPEER to instruct correction of pathogenic variants in patient cells and find that plasmid editing enriches for genomic editing up to 3-fold compared to conventional enrichment strategies. DNA repair and cell cycle-related genes are enriched in the transcriptome of edited cells. Stalling cells in the G1/S boundary increases prime editing efficiency up to 30%. Together, our results show that fluoPEER can be employed for rapid and efficient correction of patient cells, selection of gene-edited cells, and elucidation of cellular mechanisms needed for successful prime editing.
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