CRISPR-Cas "Non-Target" Sites Inhibit On-Target Cutting Rates

CRISPR-Cas systems have become ubiquitous for genome editing in eukaryotic as well as bacterial systems. Cas9 forms a complex with a guide RNA (gRNA) and searches DNA for a matching sequence (target site) next to a protospacer adjacent motif (PAM). Once found, Cas9 cuts the DNA. Cas9 is revolutionary for the ability to change the RNA sequence and target a new site easily. However, while algorithms have been developed to predict gRNA-specific Cas9 activity, a fundamental biological understanding of gRNA-specific activity is lacking. The number of PAM sites in the genome is effectively a large pool of inhibitory substrates, competing with the target site for the Cas9/gRNA complex. We demonstrate that increasing the number of non-target sites for a given gRNA reduces on-target activity in a dose-dependent manner. Furthermore, we show that the use of Cas9 mutants with increased PAM specificity toward a smaller subset of PAMs (or smaller pool of competitive substrates) improves cutting rates, while increased PAM promiscuity decreases cutting rates. Decreasing the potential search space by increasing PAM specificity provides a path toward improving on-target activity for slower high-fidelity Cas9 variants. Engineering improved PAM specificity to reduce the competitive search space offers an alternative strategy to engineer Cas9 variants with increased specificity and maintained on-target activity.

Novel Type V-A CRISPR Effectors Are Active Nucleases with Expanded Targeting Capabilities

Cas12a enzymes are quickly being adopted for use in a variety of genome-editing applications. These programmable nucleases are part of adaptive microbial immune systems, the natural diversity of which has been largely unexplored. Here, we identified novel families of Type V-A CRISPR nucleases through a large-scale analysis of metagenomes collected from a variety of complex environments, and developed representatives of these systems into gene-editing platforms. The nucleases display extensive protein variation and can be programmed by a single-guide RNA with specific motifs. The majority of these enzymes are part of systems recovered from uncultivated organisms, some of which also encode a divergent Type V effector. Biochemical analysis uncovered unexpected protospacer adjacent motif diversity, indicating that these systems will facilitate a variety of genome-engineering applications. The simplicity of guide sequences and activity in human cell lines suggest utility in gene and cell therapies.

Generation of a human induced pluripotent stem cell line with Cas9 driven by Tet-on operator via AAVS1 safe harbor gene-editing

Human induced pluripotent stem (iPS) cells expressing Cas9 protein are valuable for the pathogenic mechanism study and drug discovery. These cells can be efficiently induced to differentiate into disease cell models with specific mutations through adding designed sgRNAs. Here, we generated a human gene-editable iPS cell line by gene editing method that Cas9 gene driven by Tet-on operator was perfectly integrated into the human AAVS1 safe harbor locus. The established Cas9 expression iPS cell line named as WMUi013-A can express endogenous pluripotent markers, has the ability to differentiate into the three germ layers, and possesses a normal karyotype.

Docking sites inside Cas9 for adenine base editing diversification and RNA off-target elimination

Base editing tools with diversified editing scopes and minimized RNA off-target activities are required for broad applications. Nevertheless, current Streptococcus pyogenes Cas9 (SpCas9)-based adenine base editors (ABEs) with minimized RNA off-target activities display constrained editing scopes with efficient editing activities at positions 4-8. Here, functional ABE variants with diversified editing scopes and reduced RNA off-target activities are identified using domain insertion profiling inside SpCas9 and with different combinations of TadA variants. Engineered ABE variants in this study display narrowed, expanded or shifted editing scopes with efficient editing activities across protospacer positions 2-16. And when combined with deaminase engineering, the RNA off-target activities of engineered ABE variants are further minimized. Thus, domain insertion profiling provides a framework to improve and expand ABE toolkits, and its combination with other strategies for ABE engineering deserves comprehensive explorations in the future.

A catalogue of biochemically diverse CRISPR-Cas9 orthologs

Bacterial Cas9 nucleases from type II CRISPR-Cas antiviral defence systems have been repurposed as genome editing tools. Although these proteins are found in many microbes, only a handful of variants are used for these applications. Here, we use bioinformatic and biochemical analyses to explore this largely uncharacterized diversity. We apply cell-free biochemical screens to assess the protospacer adjacent motif (PAM) and guide RNA (gRNA) requirements of 79 Cas9 proteins, thus identifying at least 7 distinct gRNA classes and 50 different PAM sequence requirements. PAM recognition spans the entire spectrum of T-, A-, C-, and G-rich nucleotides, from single nucleotide recognition to sequence strings longer than 4 nucleotides. Characterization of a subset of Cas9 orthologs using purified components reveals additional biochemical diversity, including both narrow and broad ranges of temperature dependence, staggered-end DNA target cleavage, and a requirement for long stretches of homology between gRNA and DNA target. Our results expand the available toolset of RNA-programmable CRISPR-associated nucleases.

An efficient KRAB domain for CRISPRi applications in human cells

Clustered regularly interspaced short palindromic repeat interference (CRISPRi), based on the fusion of inactive Cas9 (dCas9) to the Krüppel-associated box (KRAB) repressor, is a powerful platform for silencing gene expression. However, it suffers from incomplete silencing of target genes. We assayed 57 KRAB domains for their repressive potency and identified the ZIM3 KRAB domain as an exceptionally potent repressor. We establish that ZIM3 KRAB-dCas9 fusion silences gene expression more efficiently than existing platforms.

multicrispr: gRNA design for prime editing and parallel targeting of thousands of targets

Targeting the coding genome to introduce nucleotide deletions/insertions via the CRISPR/Cas9 technology has become a standard procedure. It has quickly spawned a multitude of methods such as prime editing, APEX proximity labeling, or homology directed repair, for which supporting bioinformatics tools are, however, lagging behind. New CRISPR/Cas9 applications often require specific gRNA design functionality, and a generic tool is critically missing. Here, we introduce multicrispr, an R/bioconductor tool, intended to design individual gRNAs and complex gRNA libraries. The package is easy to use; detects, scores, and filters gRNAs on both efficiency and specificity; visualizes and aggregates results per target or CRISPR/Cas9 sequence; and finally returns both genomic ranges and sequences of gRNAs. To be generic, multicrispr defines and implements a genomic arithmetic framework as a basis for facile adaptation to techniques recently introduced such as prime editing or yet to arise. Its performance and design concepts such as target set-specific filtering render multicrispr a tool of choice when dealing with screening-like approaches.

Prediction of the sequence-specific cleavage activity of Cas9 variants

Several Streptococcus pyogenes Cas9 (SpCas9) variants have been developed to improve an enzyme's specificity or to alter or broaden its protospacer-adjacent motif (PAM) compatibility, but selecting the optimal variant for a given target sequence and application remains difficult. To build computational models to predict the sequence-specific activity of 13 SpCas9 variants, we first assessed their cleavage efficiency at 26,891 target sequences. We found that, of the 256 possible four-nucleotide NNNN sequences, 156 can be used as a PAM by at least one of the SpCas9 variants. For the high-fidelity variants, overall activity could be ranked as SpCas9 ≥ Sniper-Cas9 > eSpCas9(1.1) > SpCas9-HF1 > HypaCas9 ≈ xCas9 >> evoCas9, whereas their overall specificities could be ranked as evoCas9 >> HypaCas9 ≥ SpCas9-HF1 ≈ eSpCas9(1.1) > xCas9 > Sniper-Cas9 > SpCas9. Using these data, we developed 16 deep-learning-based computational models that accurately predict the activity of these variants at any target sequence.

Cas9 gene therapy for Angelman syndrome traps Ube3a-ATS long non-coding RNA

Angelman syndrome (AS) is a severe neurodevelopmental disorder caused by a mutation or deletion of the maternally inherited UBE3A allele. In neurons, the paternally inherited UBE3A allele is silenced in cis by a long non-coding RNA called UBE3A-ATS. Here, as part of a systematic screen, we found that Cas9 can be used to activate ('unsilence') paternal Ube3a in cultured mouse and human neurons when targeted to Snord115 genes, which are small nucleolar RNAs that are clustered in the 3' region of Ube3a-ATS. A short Cas9 variant and guide RNA that target about 75 Snord115 genes were packaged into an adeno-associated virus and administered to a mouse model of AS during the embryonic and early postnatal stages, when the therapeutic benefit of restoring Ube3a is predicted to be greatest1,2. This early treatment unsilenced paternal Ube3a throughout the brain for at least 17 months and rescued anatomical and behavioural phenotypes in AS mice. Genomic integration of the adeno-associated virus vector into Cas9 target sites caused premature termination of Ube3a-ATS at the vector-derived polyA cassette, or when integrated in the reverse orientation, by transcriptional collision with the vector-derived Cas9 transcript. Our study shows that targeted genomic integration of a gene therapy vector can restore the function of paternally inherited UBE3A throughout life, providing a path towards a disease-modifying treatment for a syndromic neurodevelopmental disorder.

Shortening the sgRNA-DNA interface enables SpCas9 and eSpCas9(1.1) to nick the target DNA strand

The length of the sgRNA-DNA complementary sequence is a key factor influencing the cleavage activity of Streptococcus pyogenes Cas9 (SpCas9) and its variants. The detailed mechanism remains unknown. Here, based on in vitro cleavage assays and base editing analysis, we demonstrate that reducing the length of this complementary region can confer nickase activity on SpCas9 and eSpCas9(1.1). We also show that these nicks are made on the target DNA strand. These properties encouraged us to develop a dual-functional system that simultaneously carries out double-strand DNA cleavage and C-to-T base conversions at separate targets. This system provides a novel tool for achieving trait stacking in plants.

A cell cycle-dependent CRISPR-Cas9 activation system based on an anti-CRISPR protein shows improved genome editing accuracy

The development of genome editing systems based on the Cas9 endonuclease has greatly facilitated gene knockouts and targeted genetic alterations. Precise editing of target genes without off-target effects is crucial to prevent adverse effects in clinical applications. Although several methods have been reported to result in less off-target effects associated with the CRISPR technology, these often exhibit lower editing efficiency. Therefore, efficient, accurate, and innocuous CRISPR technology is still required. Anti-CRISPR proteins are natural inhibitors of CRISPR-Cas systems derived from bacteriophages. Here, the anti-CRISPR protein, AcrIIA4, was fused with the N terminal region of human Cdt1 that is degraded specifically in S and G2, the phases of the cell cycle when homology-directed repair (HDR) is dominant. Co-expression of SpyCas9 and AcrIIA4-Cdt1 not only increases the frequency of HDR but also suppress off-targets effects. Thus, the combination of SpyCas9 and AcrIIA4-Cdt1 is a cell cycle-dependent Cas9 activation system for accurate and efficient genome editing.

AcrIIA5 Inhibits a Broad Range of Cas9 Orthologs by Preventing DNA Target Cleavage

CRISPR-Cas9 is an adaptive immune system for prokaryotes to defend against invasive genetic elements such as phages and has been used as a powerful tool for genome editing and modulation. To overcome CRISPR immunity, phages encode anti-CRISPR proteins (Acrs) to inhibit Cas9, providing an efficient "off-switch" tool for Cas9-based applications. Here, we characterized AcrIIA5, which is a Cas9 inhibitor discovered in a virulent phage of Streptococcus thermophilus. We found that AcrIIA5 is a potent and broad-spectrum inhibitor of CRISPR-Cas9, which can inhibit diverse Cas9 orthologs of type II-A, type II-B, and type II-C. AcrIIA5 inhibits Cas9 by preventing DNA target cleavage, but DNA target binding of Cas9 is unaffected. Importantly, it can affect the activity of the RuvC nuclease domain of Cas9 independent of the HNH nuclease domain. Our work expands the diversity of the inhibitory mechanisms used by Acrs and provides the guidance for developing controlling tools in Cas9-based applications.

Campylobacter jejuni Cas9 Modulates the Transcriptome in Caco-2 Intestinal Epithelial Cells

The zoonotic human pathogen Campylobacter jejuni is known for its ability to induce DNA-damage and cell death pathology in humans. The molecular mechanism behind this phenomenon involves nuclear translocation by Cas9, a nuclease in C. jejuni (CjeCas9) that is the molecular marker of the Type II CRISPR-Cas system. However, it is unknown via which cellular pathways CjeCas9 drives human intestinal epithelial cells into cell death. Here, we show that CjeCas9 released by C. jejuni during the infection of Caco-2 human intestinal epithelial cells directly modulates Caco-2 transcriptomes during the first four hours of infection. Specifically, our results reveal that CjeCas9 activates DNA damage (p53, ATM (Ataxia Telangiectasia Mutated Protein)), pro-inflammatory (NF-κB (Nuclear factor-κB)) signaling and cell death pathways, driving Caco-2 cells infected by wild-type C. jejuni, but not when infected by a cas9 deletion mutant, towards programmed cell death. This work corroborates our previous finding that CjeCas9 is cytotoxic and highlights on a RNA level the basal cellular pathways that are modulated.

Production of genome-edited mice by visualization of nucleases introduced into the embryos using electroporation

Genome editing technology contributes to the quick and highly efficient production of genetically engineered animals. These animals are helpful in clarifying the mechanism of human disease. Recently, a new electroporation technique (TAKE: Technique for animal knockout system by electroporation) was developed to produce genome-edited animals by introducing nucleases into intact embryos using electroporation instead of the microinjection method. The aim of this study was to increase the efficiency of production of genome-edited animals using the TAKE method. In the conventional protocol, it was difficult to confirm the introduction of nucleases into embryos and energization during operation. Using only embryos that introduced nucleases for embryo transfer, it will lead to increased efficiency in the production of genome-edited animals. This study examined the visualization in the introduction of nucleases into the embryos by using nucleases fluorescent labeled with ATTO-550. The embryos were transfected with Cas9 protein and fluorescent labeled dual guide RNA (mixture with crRNA and tracrRNA with ATTO-550) targeted tyrosinase gene by the TAKE method. All embryos that survived after electroporation showed fluorescence. Of these embryos with fluorescence, 43.7% developed to morphologically normal offspring. In addition, 91.7% of offspring were edited by the tyrosinase gene. This study is the first to demonstrate that the introduction of nucleases into embryos by the TAKE method could be visualized using fluorescent-labeled nucleases. This improved TAKE method can be used to produce genome-edited animals and confirm energization during operation.

Enhancing site-specific DNA integration by a Cas9 nuclease fused with a DNA donor-binding domain

The CRISPR/Cas system is widely used for genome editing. However, robust and targeted insertion of a DNA segment remains a challenge. Here, we present a fusion nuclease (Cas9-N57) to enhance site-specific DNA integration via a fused DNA binding domain of Sleeping Beauty transposase to tether the DNA segment to the Cas9/sgRNA complex. The insertion was unidirectional and specific, and DNA fragments up to 12 kb in length were successfully integrated. As a test of the system, Cas9-N57 mediated the insertion of a CD19-specific chimeric antigen receptor (CD19-CAR) cassette into the AAVS1 locus in human T cells, and induced intrahepatic cholangiocarcinoma in mice by simultaneously mediating the insertion of oncogenic KrasG12D into the Rosa26 locus and disrupting Trp53 and Pten. Moreover, the nuclease-N57 fusion proteins based on AsCpf1 (AsCas12a) and CjCas9 exhibited similar activity. These findings demonstrate that CRISPR-associated nuclease-N57 protein fusion is a powerful tool for targeted DNA insertion and holds great potential for gene therapy applications.

In situ reprogramming of gut bacteria by oral delivery

Abundant links between the gut microbiota and human health indicate that modification of bacterial function could be a powerful therapeutic strategy. The inaccessibility of the gut and inter-connections between gut bacteria and the host make it difficult to precisely target bacterial functions without disrupting the microbiota and/or host physiology. Herein we describe a multidisciplinary approach to modulate the expression of a specific bacterial gene within the gut by oral administration. We demonstrate that an engineered temperate phage λ expressing a programmable dCas9 represses a targeted E. coli gene in the mammalian gut. To facilitate phage administration while minimizing disruption to host processes, we develop an aqueous-based encapsulation formulation with a microbiota-based release mechanism and show that it facilitates oral delivery of phage in vivo. Finally we combine these technologies and show that bacterial gene expression in the mammalian gut can be precisely modified in situ with a single oral dose.

Construction of wild-type Yarrowia lipolytica IMUFRJ 50682 auxotrophic mutants using dual CRISPR/Cas9 strategy for novel biotechnological approaches

Yarrowia lipolytica IMUFRJ 50682 is a Brazilian wild-type strain with potential application in bioconversion processes which can be improved through synthetic biology. In this study, we focused on a combinatorial dual cleavage CRISPR/Cas9-mediated for construction of irreversible auxotrophic mutants IMUFRJ 50682, which genomic information is not available, thought paired sgRNAs targeting upstream and downstream sites of URA3 gene. The disruption efficiency ranged from 5 to 28 % for sgRNAs combinations closer to URA3's start and stop codon and the auxotrophic mutants lost about 970 bp containing all coding sequence, validating this method for genomic edition of wild-type strains. In addition, we introduced a fluorescent phenotype and achieved cloning rates varying from 80 to 100 %. The ura3Δ strains IMUFRJ 50682 were also engineered for β-carotene synthesis as proof of concept. Carotenoid-producing strains exhibited a similar growth profile compared to the wild-type strain and were able to synthesized 30.54-50.06 mg/L (up to 4.8 mg/g DCW) of β-carotene in YPD and YNB flask cultures, indicating a promisor future of the auxotrophic mutants IMUFRJ 50682 as a chassis for production of novel value-added chemicals.

Engineered RNA-Interacting CRISPR Guide RNAs for Genetic Sensing and Diagnostics

CRISPR guide RNAs (gRNAs) can be programmed with relative ease to allow the genetic editing of nearly any DNA or RNA sequence. Here, we propose novel molecular architectures to achieve RNA-dependent modulation of CRISPR activity in response to specific RNA molecules. We designed and tested, in both living Escherichia coli cells and cell-free assays for rapid prototyping, cis-repressed RNA-interacting guide RNA (igRNA) that switch to their active state only upon interaction with small RNA fragments or long RNA transcripts, including pathogen-derived mRNAs of medical relevance such as the human immunodeficiency virus infectivity factor. The proposed CRISPR-igRNAs are fully customizable and easily adaptable to the majority if not all the available CRISPR-Cas variants to modulate a variety of genetic functions in response to specific cellular conditions, providing orthogonal activation and increased specificity. We thereby foresee a large scope of application for therapeutic, diagnostic, and biotech applications in both prokaryotic and eukaryotic systems.

Progress Toward Zygotic and Germline Gene Drives in Mice

CRISPR-based synthetic gene drives have the potential to deliver a more effective and humane method of invasive vertebrate pest control than current strategies. Relatively efficient CRISPR gene drive systems have been developed in insects and yeast but not in mammals. Here, we investigated the efficiency of CRISPR-Cas9-based gene drives in Mus musculus by constructing "split drive" systems where gRNA expression occurs on a separate chromosome to Cas9, which is under the control of either a zygotic (CAG) or germline (Vasa) promoter. While both systems generated double-strand breaks at their intended target site in vivo, no homology-directed repair between chromosomes ("homing") was detectable. Our data indicate that robust and specific Cas9 expression during meiosis is a critical requirement for the generation of efficient CRISPR-based synthetic gene drives in rodents.

In Vitro Validation of Transgene Expression in Gene-Edited Pigs Using CRISPR Transcriptional Activators

The use of CRISPR-Cas and RNA-guided endonucleases has drastically changed research strategies for understanding and exploiting gene function, particularly for the generation of gene-edited animal models. This has resulted in an explosion in the number of gene-edited species, including highly biomedically relevant pig models. However, even with error-free DNA insertion or deletion, edited genes are occasionally not expressed and/or translated as expected. Therefore, there is a need to validate the expression outcomes gene modifications in vitro before investing in the costly generation of a gene-edited animal. Unfortunately, many gene targets are tissue specific and/or not expressed in cultured primary cells, making validation difficult without generating an animal. In this study, using pigs as a proof of concept, we show that CRISPR-dCas9 transcriptional activators can be used to validate functional transgene insertion in nonexpressing easily cultured cells such as fibroblasts. This is a tool that can be used across disciplines and animal species to save time and resources by verifying expected outcomes of gene edits before generating live animals.

Harnessing endogenous repair mechanisms for targeted gene knock-in of bovine embryos

Introducing useful traits into livestock breeding programs through gene knock-ins has proven challenging. Typically, targeted insertions have been performed in cell lines, followed by somatic cell nuclear transfer cloning, which can be inefficient. An alternative is to introduce genome editing reagents and a homologous recombination (HR) donor template into embryos to trigger homology directed repair (HDR). However, the HR pathway is primarily restricted to actively dividing cells (S/G2-phase) and its efficiency for the introduction of large DNA sequences in zygotes is low. The homology-mediated end joining (HMEJ) approach has been shown to improve knock-in efficiency in non-dividing cells and to harness HDR after direct injection of embryos. The knock-in efficiency for a 1.8 kb gene was contrasted when combining microinjection of a gRNA/Cas9 ribonucleoprotein complex with a traditional HR donor template or an HMEJ template in bovine zygotes. The HMEJ template resulted in a significantly higher rate of gene knock-in as compared to the HR template (37.0% and 13.8%; P < 0.05). Additionally, more than a third of the knock-in embryos (36.9%) were non-mosaic. This approach will facilitate the one-step introduction of gene constructs at a specific location of the bovine genome and contribute to the next generation of elite cattle.

Development of an ObLiGaRe Doxycycline Inducible Cas9 system for pre-clinical cancer drug discovery

The CRISPR-Cas9 system has increased the speed and precision of genetic editing in cells and animals. However, model generation for drug development is still expensive and time-consuming, demanding more target flexibility and faster turnaround times with high reproducibility. The generation of a tightly controlled ObLiGaRe doxycycline inducible SpCas9 (ODInCas9) transgene and its use in targeted ObLiGaRe results in functional integration into both human and mouse cells culminating in the generation of the ODInCas9 mouse. Genomic editing can be performed in cells of various tissue origins without any detectable gene editing in the absence of doxycycline. Somatic in vivo editing can model non-small cell lung cancer (NSCLC) adenocarcinomas, enabling treatment studies to validate the efficacy of candidate drugs. The ODInCas9 mouse allows robust and tunable genome editing granting flexibility, speed and uniformity at less cost, leading to high throughput and practical preclinical in vivo therapeutic testing.

Conditional Recruitment to a DNA-Bound CRISPR-Cas Complex Using a Colocalization-Dependent Protein Switch

To spatially control biochemical functions at specific sites within a genome, we have engineered a synthetic switch that activates when bound to its DNA target site. The system uses two CRISPR-Cas complexes to colocalize components of a de novo-designed protein switch (Co-LOCKR) to adjacent sites in the genome. Colocalization triggers a conformational change in the switch from an inactive closed state to an active open state with an exposed functional peptide. We prototype the system in yeast and demonstrate that DNA binding triggers activation of the switch, recruitment of a transcription factor, and expression of a downstream reporter gene. This DNA-triggered Co-LOCKR switch provides a platform to engineer sophisticated functions that should only be executed at a specific target site within the genome, with potential applications in a wide range of synthetic systems including epigenetic regulation, imaging, and genetic logic circuits.

A knockdown of the herpes simplex virus type-1 gene in all-in-one CRISPR vectors

INTRODUCTION

Herpes simplex virus type 1 (HSV-1) is a virus that causes serious human disease and establishes a long-term latent infection. The latent form of this virus has shown to be resistant to antiviral drugs. Clustered Regularly Interspace Short Palindromic Repeats (CRISPR), is an important tool in genome engineering and composed of guide RNA (gRNA) and Cas9 nuclease that makes an RNA-protein complex to digest exclusive target sequences implementation of gRNA. Moreover, CRISPR-Cas9 system effectively suppresses HSV-1 infection by knockout of some viral genes.

MATERIALS AND METHODS

To survey the efficacy of Cas9 system on HSV-1 genome destruction, we designed several guide RNAs (gRNAs) that all packaged in one vector. Additionally, we performed a one-step restriction using BamHI and Esp3I enzymes.

RESULTS

CRISPR/Cas9 system targeted against the gD gene of HSV-1 was transfected into HEK-AD cells that showed a significant reduction of HSV-1 infection by plaque assay and real-time PCR.

CONCLUSION

The pCas-Guide-EF1a-GFP CRISPR vector can create a fast and efficient method for gRNA cloning by restriction enzymes (Esp3I (BsmBI) and BamHI). Therefore, the CRISPR/Cas9 system may be utilized for the screening of genes critical for the HSV-1 infection and developing new strategies for targeted therapy of viral infections caused by HSV-1.

Establishment of a PEG-mediated protoplast transformation system based on DNA and CRISPR/Cas9 ribonucleoprotein complexes for banana

BACKGROUND

To date, CRISPR/Cas9 RNP editing tools have not been applied to the genetic modification of banana. Here, the establishment of a PEG-mediated banana protoplast transformation system makes it possible to build an efficient DNA-free method for a site-directed mutagenesis system.

RESULTS

Protoplasts constitute a versatile platform for transient expression in plant science. In this study, we established a PEG-mediated banana protoplast transformation system. This system was further optimized for successfully delivering CRISPR/Cas9 and CRISPR/Cas12a plasmids and CRISPR/Cas9 ribonucleoproteins (RNPs) for targeted delivery of the PDS gene into banana protoplasts. Specific bands were observed in PCR-Restriction Enzyme Digestion (PCR-RE) assays, and Sanger sequencing of single clones further confirmed the occurrence of indels at target sites. Deep amplicon sequencing results showed that the editing efficiency of the CRISPR/Cas9 system was higher than that of the other two systems.

CONCLUSIONS

The PEG-mediated banana protoplast transformation system can serve as a rapid and effective tool for transient expression assays and sgRNA validation in banana. The application of the CRISPR/Cas9 RNP system enables the generation of banana plants engineered by DNA-free gene editing.