Expanding the CRISPR Toolbox with ErCas12a in Zebrafish and Human Cells

Wierson, Wesley A.; Simone, Brandon W.; WareJoncas, Zachary; Mann, Carla M.; Welker, Jordan M.; Kar, Bibekananda; Gendron, William A.C.; Barry, Michael A.; Clark, Karl J.; Dobbs, Drena; McGrail, Maura; Ekker, Stephen C.; Essner, Jeffrey J.

doi:10.1101/650515

Cited by 4 publications

(9 citation statements)

References 44 publications

(44 reference statements)

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“…We next assessed the specificity and precision of the approach. It is possible that either concatemerization of inserts or off-target integrations could occur after foreign DNA delivery and a CRISPR/Cas9 mediated DSBs (Gutierrez-Triana et al, 2018, Doench et al, 2016, Fu et al, 2013, Paix et al, 2017a, Won and Dawid, 2017, Yan et al, 2013, Hackett et al, 2007, Wierson et al, 2020, Wierson et al, 2019. To identify off-target insertions genome-wide and verify single-copy integration we performed next generation Whole Genome Sequencing (WGS) with high coverage (for details of WGS, see material and methods) on three knock-in lines (Figure 1B-D); eGFP-cbx1b, mScarlet-pcna and mNeonGreen-myosinhc .…”

Section: Precise Single Copy Knock-ins Of Fluorescent Protein Reportersmentioning

confidence: 99%

Endogenous protein tagging in medaka using a simplified CRISPR/Cas9 knock-in approach

Seleit

Aulehla

Paix

2021

Preprint

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The CRISPR/Cas9 system has been used to generate fluorescently labelled fusion proteins by homology directed repair in a variety of species. Despite its success, there remains an urgent need for increased simplicity and efficiency of genome editing in research organisms. Here, we establish a simplified, highly efficient and precise strategy for CRISPR/Cas9 mediated endogenous protein tagging in medaka. We use a cloning-free approach that relies on PCR amplified donor fragments containing the fluorescent reporter sequences flanked by short homology arms (30-40bp), a synthetic sgRNA and streptavidin tagged Cas9. We generate six novel knock-in lines with high efficiency of F0 targeting and germline transmission. Whole Genome Sequencing results reveal single-copy integration events only at the targeted loci. We provide an initial characterization of these fusion-protein lines, significantly expanding the repertoire of genetic tools available in medaka. In particular, we show that the mScarlet-pcna knock-in has the potential to serve as an organismal-wide label for proliferative zones and an endogenous cell cycle reporter.

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Section: Precise Single Copy Knock-ins Of Fluorescent Protein Reportersmentioning

confidence: 99%

Endogenous protein tagging in medaka using a simplified CRISPR/Cas9 knock-in approach

Seleit

Aulehla

Paix

2021

Preprint

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“…• For many gene-edited cell therapy applications, scientists do not wish to fix single nucleotide polymorphisms, but need to add exogenous DNA to educate cells with new functions that nature did not evolve. In research applications, this could be the addition of DNA encoding fluorescent proteins to tag spatiotemporal gene expression patterns in model organisms and cell lines (Wierson et al, 2020). In the clinic, the best example is the addition of DNA encoding chimeric antigen receptors to impart tumor killing activity onto T cells.…”

Section: Homology-directed Repair (Hdr)mentioning

confidence: 99%

“…Just like in templated SNP alterations, the homologous recombination pathway of DNA repair is a commonly employed mechanism for these alterations. There are publications demonstrating the efficacy of many types of templates to accomplish this feat; single stranded or double stranded linear DNAs (Roth et al, 2018) (Shin et al, 2014), plasmid DNA (Wierson et al, 2020), or rAAV donors (MacLeod et al, 2017) to mediate site-specific integration of this DNA. More recently, scientists have discovered that delivering a donor template with CRISPR sites flanking homology arms and cargo DNA is an efficient way of inducing a sub-pathway of HDR, dubbed homology-mediated end joining, to create engineered model organisms and cell lines (Wierson et al, 2020) (Yao et al, 2017) (Hisano et al, 2015).…”

Section: Homology-directed Repair (Hdr)mentioning

confidence: 99%

“…Indeed, the University of Pennsylvania is currently recruiting dogs to participate in a trial with human CAR-T cells engineered to fight canine B cell lymphoma, though it is not publicly known how the CAR module is integrated into the genome in this approach (are they using lentivirus or gene editing approaches?). In industry, LEAH Labs is developing a similar approach using CRISPR to knock-out the TCR while simultaneously integrating CAR DNA using their nonviral GeneWeld platform (Wierson et al, 2020. Of note, a combination approach using lentivirus to introduce the CAR transgene and gene editing to knock out the TCR to generate xenogeneic CAR-T cells for canine use may be an effective strategy toward the application of this therapy.…”

Section: Proof Of Concept Car-t Cell Therapy In Dogsmentioning

confidence: 99%

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Gene Editing and Gene Therapy: Entering Uncharted Territory in Veterinary Oncology

Wierson¹,

Abel²,

Siegler³

et al. 2021

Preprint

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With rapid advances in gene editing and gene therapy technologies, the development of genetic, cell, or protein-based cures to disease are no longer the realm of science fiction but that of today’s practice. The impact of these technologies are rapidly bringing them to the veterinary market as both enhanced therapeutics and towards modeling their outcomes for translational application. Simply put, gene editing enables scientists to modify an organism’s DNA a priori through the use of site-specific DNA targeting tools like clustered regularly interspaced short palindromic repeats and CRISPR-associated protein 9 (CRISPR/Cas9). Gene therapy is a broader definition that encompasses the addition of exogenous genetic materials into specific cells to correct a genetic defect. More precisely, the U.S Food and Drug Administration (FDA) defines gene therapy as “a technique that modifies a person’s genes to treat or cure disease” by either (i) replacing a disease-causing gene with a healthy copy of the gene; (ii) inactivating a disease-causing gene that was not functioning properly; or (iii) introducing a new or modified gene into the body to help treat a disease. In some instances, this can be accomplished through direct transfer of DNA or RNA into target cells of interest or more broadly through gene editing. While gene therapy is possible through the simple addition of genetic information into cells of interest, gene editing allows the genome to be reprogrammed intentionally through the deletion of diseased alleles, reconstitution of wild type sequence, or targeted integration of exogenous DNA to impart new function. Cells can be removed from the body, altered, and reinfused, or edited in vivo. Indeed, manufacturing and production efficiencies in gene editing and gene therapy in the 21st century has brought the therapeutic potential of in vitro and in vivo reprogrammed cells, to the front lines of therapeutic intervention (Brooks et al., 2016). For example, CAR-T cell therapy is revolutionizing hematologic cancer care in humans and is being translated to canines by us and others, and gene therapy trials are ongoing for mitral valve disease in dogs.

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“…pSC-CMV-Cre was previously developed by the Barry lab and consists of a CMV driven Cre flanked by AAV ITRs [12]. ErCas12a plasmid was cloned as previously described in Wierson et al [2].…”

Section: Plasmids and Cloningmentioning

confidence: 99%

Unlocking LoxP to Track Genome Editing In Vivo

Gendron¹,

Rubin²,

Simone³

et al. 2021

Preprint

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The development of CRISPR associated proteins, such as Cas9, has led to increased accessibility and ease of use in genome editing. However, additional tools are needed to quantify and identify successful genome editing events in living animals. We developed a method to rapidly and quantitatively monitor gene editing activity non-invasively in living animals that also facilitates confocal microscopy and nucleotide level analyses at the end of study. Here we report a new CRISPR “footprinting” approach to activate luciferase and fluorescent proteins in mice as a function of gene editing. This system is based on experience with our prior Cre-detector system and is designed for Cas editors able to target LoxP including gRNAs including SaCas9 and ErCas12a [1, 2]. These CRISPRs cut specifically within LoxP, an approach that is a departure from previous gene editing in vivo activity detection techniques that targeted adjacent stop sequences. In this sensor paradigm, CRISPR activity was monitored non-invasively in living Cre reporter mice (FVB.129S6(B6)-Gt(ROSA)26Sortm1(Luc)Kael/J and Gt(ROSA)26Sortm4(ACTB-tdTomato,-EGFP)Luo/J, which will be referred to as LSL and mT/mG throughout the paper) after intramuscular or intravenous hydrodynamic plasmid injections, demonstrating utility in two diverse organ systems. The same genome-editing event was examined at the cellular level in specific tissues by confocal microscopy to determine the identity and frequency of successfully genome-edited cells. Further, SaCas9 induced targeted editing at efficiencies that were comparable to Cre recombinase demonstrating high effective delivery and activity in a whole animal. This work establishes genome editing tools and models to track CRISPR editing in vivo non-invasively and to fingerprint the identity of targeted cells. This approach also enables similar utility for any of the thousands of previously generated LoxP animal models.

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Expanding the CRISPR Toolbox with ErCas12a in Zebrafish and Human Cells

Cited by 4 publications

References 44 publications

Endogenous protein tagging in medaka using a simplified CRISPR/Cas9 knock-in approach

Endogenous protein tagging in medaka using a simplified CRISPR/Cas9 knock-in approach

Gene Editing and Gene Therapy: Entering Uncharted Territory in Veterinary Oncology

Unlocking LoxP to Track Genome Editing In Vivo

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