1
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Venema WJ, Hiddingh S, van Loosdregt J, Bowes J, Balliu B, de Boer JH, Ossewaarde-van Norel J, Thompson SD, Langefeld CD, de Ligt A, van der Veken LT, Krijger PHL, de Laat W, Kuiper JJW. A cis-regulatory element regulates ERAP2 expression through autoimmune disease risk SNPs. CELL GENOMICS 2024; 4:100460. [PMID: 38190099 PMCID: PMC10794781 DOI: 10.1016/j.xgen.2023.100460] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Received: 03/16/2023] [Revised: 10/04/2023] [Accepted: 11/09/2023] [Indexed: 01/09/2024]
Abstract
Single-nucleotide polymorphisms (SNPs) near the ERAP2 gene are associated with various autoimmune conditions, as well as protection against lethal infections. Due to high linkage disequilibrium, numerous trait-associated SNPs are correlated with ERAP2 expression; however, their functional mechanisms remain unidentified. We show by reciprocal allelic replacement that ERAP2 expression is directly controlled by the splice region variant rs2248374. However, disease-associated variants in the downstream LNPEP gene promoter are independently associated with ERAP2 expression. Allele-specific conformation capture assays revealed long-range chromatin contacts between the gene promoters of LNPEP and ERAP2 and showed that interactions were stronger in patients carrying the alleles that increase susceptibility to autoimmune diseases. Replacing the SNPs in the LNPEP promoter by reference sequences lowered ERAP2 expression. These findings show that multiple SNPs act in concert to regulate ERAP2 expression and that disease-associated variants can convert a gene promoter region into a potent enhancer of a distal gene.
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Affiliation(s)
- Wouter J Venema
- Department of Ophthalmology, University Medical Center Utrecht, Utrecht University, Utrecht, the Netherlands; Center for Translational Immunology, University Medical Center Utrecht, Utrecht University, Utrecht, the Netherlands
| | - Sanne Hiddingh
- Department of Ophthalmology, University Medical Center Utrecht, Utrecht University, Utrecht, the Netherlands; Center for Translational Immunology, University Medical Center Utrecht, Utrecht University, Utrecht, the Netherlands
| | - Jorg van Loosdregt
- Center for Translational Immunology, University Medical Center Utrecht, Utrecht University, Utrecht, the Netherlands
| | - John Bowes
- Centre for Genetics and Genomics Versus Arthritis, Centre for Musculoskeletal Research, Manchester Academic Health Science Centre, The University of Manchester, Manchester, UK
| | - Brunilda Balliu
- Department of Computational Medicine, David Geffen School of Medicine, University of California, Los Angeles, Los Angeles, CA, USA
| | - Joke H de Boer
- Department of Ophthalmology, University Medical Center Utrecht, Utrecht University, Utrecht, the Netherlands
| | | | - Susan D Thompson
- Department of Pediatrics, University of Cincinnati College of Medicine, Division of Human Genetics, Cincinnati Children's Hospital Medical Center, Cincinnati, OH, USA
| | - Carl D Langefeld
- Department of Biostatistics and Data Science, and Center for Precision Medicine, Wake Forest University School of Medicine, Winston-Salem, NC, USA
| | - Aafke de Ligt
- Department of Ophthalmology, University Medical Center Utrecht, Utrecht University, Utrecht, the Netherlands; Center for Translational Immunology, University Medical Center Utrecht, Utrecht University, Utrecht, the Netherlands
| | - Lars T van der Veken
- Department of Genetics, Division Laboratories, Pharmacy and Biomedical Genetics, University Medical Center Utrecht, Utrecht University, Utrecht, the Netherlands
| | - Peter H L Krijger
- Oncode Institute, Hubrecht Institute-KNAW and University Medical Center Utrecht, 3584 CT Utrecht, the Netherlands
| | - Wouter de Laat
- Oncode Institute, Hubrecht Institute-KNAW and University Medical Center Utrecht, 3584 CT Utrecht, the Netherlands
| | - Jonas J W Kuiper
- Department of Ophthalmology, University Medical Center Utrecht, Utrecht University, Utrecht, the Netherlands; Center for Translational Immunology, University Medical Center Utrecht, Utrecht University, Utrecht, the Netherlands.
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2
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Prykhozhij SV, Berman JN. Mutation Knock-in Methods Using Single-Stranded DNA and Gene Editing Tools in Zebrafish. Methods Mol Biol 2024; 2707:279-303. [PMID: 37668920 DOI: 10.1007/978-1-0716-3401-1_19] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 09/06/2023]
Abstract
Introduction or knock-in of precise genomic modifications remains one of the most important applications of CRISPR/Cas9 in all model systems including zebrafish. The most widely used type of donor template containing the desired modification is single-stranded DNA (ssDNA), either in the form of single-stranded oligodeoxynucleotides (ssODN) (<150 nucleotides (nt)) or as long ssDNA (lssDNA) molecules (up to about 2000 nt). Despite the challenges posed by DNA repair after DNA double-strand breaks, knock-in of precise mutations is relatively straightforward in zebrafish. Knock-in efficiency can be enhanced by careful donor template design, using lssDNA as template or tethering the donor template DNA to the Cas9-guide RNA complex. Other point mutation methods such as base editing and prime editing are starting to be applied in zebrafish and many other model systems. However, these methods may not always be sufficiently accessible or may have limited capacity to perform all desired mutation knock-ins which are possible with ssDNA-based knock-in methods. Thus, it is likely that there will be complementarity in the technologies used for generating precise mutants. Here, we review and describe a suite of CRISPR/Cas9 knock-in procedures utilizing ssDNA as the donor template in zebrafish, point out the potential challenges and suggest possible approaches for their solution ultimately leading to successful generation of precise mutant lines.
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Affiliation(s)
- Sergey V Prykhozhij
- Children's Hospital of Eastern Ontario Research Institute, Ottawa, ON, Canada
| | - Jason N Berman
- Children's Hospital of Eastern Ontario Research Institute, Ottawa, ON, Canada.
- Departments of Pediatrics and Cellular and Molecular Medicine, University of Ottawa, Ottawa, ON, Canada.
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3
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Chen C, Wang Z, Kang M, Lee KB, Ge X. High-fidelity large-diversity monoclonal mammalian cell libraries by cell cycle arrested recombinase-mediated cassette exchange. Nucleic Acids Res 2023; 51:e113. [PMID: 37941133 PMCID: PMC10711435 DOI: 10.1093/nar/gkad1001] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/08/2023] [Revised: 09/26/2023] [Accepted: 10/18/2023] [Indexed: 11/10/2023] Open
Abstract
Mammalian cells carrying defined genetic variations have shown great potentials in both fundamental research and therapeutic development. However, their full use was limited by lack of a robust method to construct large monoclonal high-quality combinatorial libraries. This study developed cell cycle arrested recombinase-mediated cassette exchange (aRMCE), able to provide monoclonality, precise genomic integration and uniform transgene expression. Via optimized nocodazole-mediated mitotic arrest, 20% target gene replacement efficiency was achieved without antibiotic selection, and the improved aRMCE efficiency was applicable to a variety of tested cell clones, transgene targets and transfection methods. As a demonstration of this versatile method, we performed directed evolution of fragment crystallizable (Fc), for which error-prone libraries of over 107 variants were constructed and displayed as IgG on surface of CHO cells. Diversities of constructed libraries were validated by deep sequencing, and panels of novel Fc mutants were identified showing improved binding towards specific Fc gamma receptors and enhanced effector functions. Due to its large cargo capacity and compatibility with different mutagenesis approaches, we expect this mammalian cell platform technology has broad applications for directed evolution, multiplex genetic assays, cell line development and stem cell engineering.
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Affiliation(s)
- Chuan Chen
- Department of Chemical and Environmental Engineering, University of California Riverside, Riverside, CA 92521, USA
| | - Zening Wang
- Department of Chemical and Environmental Engineering, University of California Riverside, Riverside, CA 92521, USA
- Institute of Molecular Medicine, University of Texas Health Science Center at Houston, Houston, TX 77030, USA
| | - Minhyo Kang
- Department of Chemical and Environmental Engineering, University of California Riverside, Riverside, CA 92521, USA
| | - Ki Baek Lee
- Institute of Molecular Medicine, University of Texas Health Science Center at Houston, Houston, TX 77030, USA
| | - Xin Ge
- Department of Chemical and Environmental Engineering, University of California Riverside, Riverside, CA 92521, USA
- Institute of Molecular Medicine, University of Texas Health Science Center at Houston, Houston, TX 77030, USA
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4
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Bekaert B, Boel A, De Witte L, Vandenberghe W, Popovic M, Stamatiadis P, Cosemans G, Tordeurs L, De Loore AM, Chuva de Sousa Lopes SM, De Sutter P, Stoop D, Coucke P, Menten B, Heindryckx B. Retained chromosomal integrity following CRISPR-Cas9-based mutational correction in human embryos. Mol Ther 2023; 31:2326-2341. [PMID: 37376733 PMCID: PMC10422011 DOI: 10.1016/j.ymthe.2023.06.013] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/09/2022] [Revised: 04/11/2023] [Accepted: 06/22/2023] [Indexed: 06/29/2023] Open
Abstract
Human germline gene correction by targeted nucleases holds great promise for reducing mutation transmission. However, recent studies have reported concerning observations in CRISPR-Cas9-targeted human embryos, including mosaicism and loss of heterozygosity (LOH). The latter has been associated with either gene conversion or (partial) chromosome loss events. In this study, we aimed to correct a heterozygous basepair substitution in PLCZ1, related to infertility. In 36% of the targeted embryos that originated from mutant sperm, only wild-type alleles were observed. By performing genome-wide double-digest restriction site-associated DNA sequencing, integrity of the targeted chromosome (i.e., no deletions larger than 3 Mb or chromosome loss) was confirmed in all seven targeted GENType-analyzed embryos (mutant editing and absence of mutation), while short-range LOH events (shorter than 10 Mb) were clearly observed by single-nucleotide polymorphism assessment in two of these embryos. These results fuel the currently ongoing discussion on double-strand break repair in early human embryos, making a case for the occurrence of gene conversion events or partial template-based homology-directed repair.
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Affiliation(s)
- Bieke Bekaert
- Ghent-Fertility and Stem Cell Team (G-FaST), Department for Reproductive Medicine, Ghent University Hospital, Corneel Heymanslaan 10, 9000 Ghent, Belgium
| | - Annekatrien Boel
- Ghent-Fertility and Stem Cell Team (G-FaST), Department for Reproductive Medicine, Ghent University Hospital, Corneel Heymanslaan 10, 9000 Ghent, Belgium
| | - Lisa De Witte
- Center for Medical Genetics Ghent, Department of Biomolecular Medicine, Ghent University, Corneel Heymanslaan 10, 9000 Ghent, Belgium
| | - Winter Vandenberghe
- Ghent-Fertility and Stem Cell Team (G-FaST), Department for Reproductive Medicine, Ghent University Hospital, Corneel Heymanslaan 10, 9000 Ghent, Belgium
| | - Mina Popovic
- Ghent-Fertility and Stem Cell Team (G-FaST), Department for Reproductive Medicine, Ghent University Hospital, Corneel Heymanslaan 10, 9000 Ghent, Belgium
| | - Panagiotis Stamatiadis
- Ghent-Fertility and Stem Cell Team (G-FaST), Department for Reproductive Medicine, Ghent University Hospital, Corneel Heymanslaan 10, 9000 Ghent, Belgium
| | - Gwenny Cosemans
- Ghent-Fertility and Stem Cell Team (G-FaST), Department for Reproductive Medicine, Ghent University Hospital, Corneel Heymanslaan 10, 9000 Ghent, Belgium
| | - Lise Tordeurs
- Ghent-Fertility and Stem Cell Team (G-FaST), Department for Reproductive Medicine, Ghent University Hospital, Corneel Heymanslaan 10, 9000 Ghent, Belgium
| | - Athina-Maria De Loore
- Ghent-Fertility and Stem Cell Team (G-FaST), Department for Reproductive Medicine, Ghent University Hospital, Corneel Heymanslaan 10, 9000 Ghent, Belgium
| | - Susana Marina Chuva de Sousa Lopes
- Ghent-Fertility and Stem Cell Team (G-FaST), Department for Reproductive Medicine, Ghent University Hospital, Corneel Heymanslaan 10, 9000 Ghent, Belgium; Department of Anatomy and Embryology, Leiden University Medical Center, 2333 ZA Leiden, the Netherlands
| | - Petra De Sutter
- Ghent-Fertility and Stem Cell Team (G-FaST), Department for Reproductive Medicine, Ghent University Hospital, Corneel Heymanslaan 10, 9000 Ghent, Belgium
| | - Dominic Stoop
- Ghent-Fertility and Stem Cell Team (G-FaST), Department for Reproductive Medicine, Ghent University Hospital, Corneel Heymanslaan 10, 9000 Ghent, Belgium
| | - Paul Coucke
- Center for Medical Genetics Ghent, Department of Biomolecular Medicine, Ghent University, Corneel Heymanslaan 10, 9000 Ghent, Belgium
| | - Björn Menten
- Center for Medical Genetics Ghent, Department of Biomolecular Medicine, Ghent University, Corneel Heymanslaan 10, 9000 Ghent, Belgium
| | - Björn Heindryckx
- Ghent-Fertility and Stem Cell Team (G-FaST), Department for Reproductive Medicine, Ghent University Hospital, Corneel Heymanslaan 10, 9000 Ghent, Belgium.
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5
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Asuncion JD, Eamani A, Rohrbach EW, Knapp EM, Deshpande SA, Bonanno SL, Murphy JE, Lawal HO, Krantz DE. Precise CRISPR-Cas9-mediated mutation of a membrane trafficking domain in the Drosophila vesicular monoamine transporter gene. Curr Res Physiol 2023; 6:100101. [PMID: 37409154 PMCID: PMC10318446 DOI: 10.1016/j.crphys.2023.100101] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/29/2023] [Revised: 05/16/2023] [Accepted: 06/19/2023] [Indexed: 07/07/2023] Open
Abstract
Monoamine neurotransmitters such as noradrenalin are released from both synaptic vesicles (SVs) and large dense-core vesicles (LDCVs), the latter mediating extrasynaptic signaling. The contribution of synaptic versus extrasynaptic signaling to circuit function and behavior remains poorly understood. To address this question, we have previously used transgenes encoding a mutation in the Drosophila Vesicular Monoamine Transporter (dVMAT) that shifts amine release from SVs to LDCVs. To circumvent the use of transgenes with non-endogenous patterns of expression, we have now used CRISPR-Cas9 to generate a trafficking mutant in the endogenous dVMAT gene. To minimize disruption of the dVMAT coding sequence and a nearby RNA splice site, we precisely introduced a point mutation using single-stranded oligonucleotide repair. A predicted decrease in fertility was used as a phenotypic screen to identify founders in lieu of a visible marker. Phenotypic analysis revealed a defect in the ovulation of mature follicles and egg retention in the ovaries. We did not detect defects in the contraction of lateral oviducts following optogenetic stimulation of octopaminergic neurons. Our findings suggest that release of mature eggs from the ovary is disrupted by changing the balance of VMAT trafficking between SVs and LDCVs. Further experiments using this model will help determine the mechanisms that sensitize specific circuits to changes in synaptic versus extrasynaptic signaling.
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Affiliation(s)
- James D. Asuncion
- Medical Scientist Training Program, David Geffen School of Medicine, University of California, Los Angeles, CA, 90095, USA
- Department of Psychiatry and Biobehavioral Sciences, David Geffen School of Medicine, University of California, Los Angeles, CA, 90095, USA
- UCLA Neuroscience Interdepartmental Program, University of California, Los Angeles, CA, 90095, USA
| | - Aditya Eamani
- Department of Psychiatry and Biobehavioral Sciences, David Geffen School of Medicine, University of California, Los Angeles, CA, 90095, USA
| | - Ethan W. Rohrbach
- Department of Psychiatry and Biobehavioral Sciences, David Geffen School of Medicine, University of California, Los Angeles, CA, 90095, USA
- UCLA Neuroscience Interdepartmental Program, University of California, Los Angeles, CA, 90095, USA
| | - Elizabeth M. Knapp
- Department of Psychiatry and Biobehavioral Sciences, David Geffen School of Medicine, University of California, Los Angeles, CA, 90095, USA
| | - Sonali A. Deshpande
- Department of Psychiatry and Biobehavioral Sciences, David Geffen School of Medicine, University of California, Los Angeles, CA, 90095, USA
| | - Shivan L. Bonanno
- Department of Psychiatry and Biobehavioral Sciences, David Geffen School of Medicine, University of California, Los Angeles, CA, 90095, USA
| | - Jeremy E. Murphy
- Department of Biological Sciences, Delaware State University, Dover, DE, USA, 19901, USA
| | - Hakeem O. Lawal
- Department of Biological Sciences, Delaware State University, Dover, DE, USA, 19901, USA
| | - David E. Krantz
- Department of Psychiatry and Biobehavioral Sciences, David Geffen School of Medicine, University of California, Los Angeles, CA, 90095, USA
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6
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Mikkelsen NS, Hernandez SS, Jensen TI, Schneller JL, Bak RO. Enrichment of transgene integrations by transient CRISPR activation of a silent reporter gene. Mol Ther Methods Clin Dev 2023; 29:1-16. [PMID: 36922985 PMCID: PMC10009645 DOI: 10.1016/j.omtm.2023.02.010] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/15/2022] [Accepted: 02/13/2023] [Indexed: 02/18/2023]
Abstract
CRISPR-Cas-mediated site-specific integration of transgenes by homology-directed repair (HDR) is challenging, especially in primary cells, where inferior editing efficiency may impede the development of gene- and cellular therapies. Various strategies for enrichment of cells with transgene integrations have been developed, but most strategies either generate unwanted genomic scars or rely on permanent integration and expression of a reporter gene used for selection. However, stable expression of a reporter gene may perturb cell homeostasis and function. Here we develop a broadly applicable and versatile enrichment strategy by harnessing the capability of CRISPR activation (CRISPRa) to transiently induce expression of a therapeutically relevant reporter gene used for immunomagnetic enrichment. This strategy is readily adaptable to primary human T cells and CD34+ hematopoietic stem and progenitor cells (HSPCs), where enrichment of 1.8- to 3.3-fold and 3.2- to 3.6-fold was achieved, respectively. Furthermore, chimeric antigen receptor (CAR) T cells were enriched 2.5-fold and demonstrated improved cytotoxicity over non-enriched CAR T cells. Analysis of HDR integrations showed a proportion of cells harboring deletions of the transgene cassette arising either from impartial HDR or truncated adeno-associated virus (AAV) vector genomes. Nonetheless, this novel enrichment strategy expands the possibility to enrich for transgene integrations in research settings and in gene and cellular therapies.
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Affiliation(s)
| | | | - Trine I Jensen
- Department of Biomedicine, Aarhus University, Aarhus C, Denmark
| | - Jessica L Schneller
- Department of Biomedicine, Aarhus University, Aarhus C, Denmark.,RNA and Gene Therapies, Novo Nordisk A/S, Maaloev, Denmark
| | - Rasmus O Bak
- Department of Biomedicine, Aarhus University, Aarhus C, Denmark.,Aarhus Institute of Advanced Studies, Aarhus University, Aarhus C, Denmark
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7
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A tapt1 knock-out zebrafish line with aberrant lens development and impaired vision models human early-onset cataract. Hum Genet 2023; 142:457-476. [PMID: 36697720 DOI: 10.1007/s00439-022-02518-w] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/28/2022] [Accepted: 12/19/2022] [Indexed: 01/27/2023]
Abstract
Bi-allelic mutations in the gene coding for human trans-membrane anterior-posterior transformation protein 1 (TAPT1) result in a broad phenotypic spectrum, ranging from syndromic disease with severe skeletal and congenital abnormalities to isolated early-onset cataract. We present here the first patient with a frameshift mutation in the TAPT1 gene, resulting in both bilateral early-onset cataract and skeletal abnormalities, in addition to several dysmorphic features, in this way further expanding the phenotypic spectrum associated with TAPT1 mutations. A tapt1a/tapt1b double knock-out (KO) zebrafish model generated by CRISPR/Cas9 gene editing revealed an early larval phenotype with eye malformations, loss of vision, increased photokinetics and hyperpigmentation, without visible skeletal involvement. Ultrastructural analysis of the eyes showed a smaller condensed lens, loss of integrity of the lens capsule with formation of a secondary lens and hyperplasia of the cells in the ganglion and inner plexiform layers of the retina. Transcriptomic analysis pointed to an impaired lens development with aberrant expression of many of the crystallin and other lens-specific genes. Furthermore, the phototransduction and visual perception pathways were found to be significantly disturbed. Differences in light perception are likely the cause of the increased dark photokinetics and generalized hyperpigmentation observed in this zebrafish model. In conclusion, this study validates TAPT1 as a new gene for early-onset cataract and sheds light on its ultrastructural and molecular characteristics.
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8
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Elworthy S, Rutherford HA, Prajsnar TK, Hamilton NM, Vogt K, Renshaw SA, Condliffe AM. Activated PI3K delta syndrome 1 mutations cause neutrophilia in zebrafish larvae. Dis Model Mech 2023; 16:dmm049841. [PMID: 36805642 PMCID: PMC10655814 DOI: 10.1242/dmm.049841] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/15/2022] [Accepted: 02/14/2023] [Indexed: 02/22/2023] Open
Abstract
People with activated PI3 kinase delta syndrome 1 (APDS1) suffer from immune deficiency and severe bronchiectasis. APDS1 is caused by dominant activating mutations of the PIK3CD gene that encodes the PI3 kinase delta (PI3Kδ) catalytic subunit. Despite the importance of innate immunity defects in bronchiectasis, there has been limited investigation of neutrophils or macrophages in APDS1 patients or mouse models. Zebrafish embryos provide an ideal system to study neutrophils and macrophages. We used CRISPR-Cas9 and CRISPR-Cpf1, with oligonucleotide-directed homologous repair, to engineer zebrafish equivalents of the two most prevalent human APDS1 disease mutations. These zebrafish pik3cd alleles dominantly caused excessive neutrophilic inflammation in a tail-fin injury model. They also resulted in total body neutrophilia in the absence of any inflammatory stimulus but normal numbers of macrophages. Exposure of zebrafish to the PI3Kδ inhibitor CAL-101 reversed the total body neutrophilia. There was no apparent defect in neutrophil maturation or migration, and tail-fin regeneration was unimpaired. Overall, the finding is of enhanced granulopoeisis, in the absence of notable phenotypic change in neutrophils and macrophages.
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Affiliation(s)
- Stone Elworthy
- Department of Infection, Immunity and Cardiovascular Disease, University of Sheffield, Sheffield S10 2TN, UK
| | - Holly A. Rutherford
- Department of Infection, Immunity and Cardiovascular Disease, University of Sheffield, Sheffield S10 2TN, UK
| | - Tomasz K. Prajsnar
- Department of Infection, Immunity and Cardiovascular Disease, University of Sheffield, Sheffield S10 2TN, UK
- Department of Evolutionary Immunology, Institute of Zoology and Biomedical Research, Jagiellonian University, Gronostajowa 9, 30-387 Krakow, Poland
| | - Noémie M. Hamilton
- Department of Infection, Immunity and Cardiovascular Disease, University of Sheffield, Sheffield S10 2TN, UK
| | - Katja Vogt
- Department of Infection, Immunity and Cardiovascular Disease, University of Sheffield, Sheffield S10 2TN, UK
| | - Stephen A. Renshaw
- Department of Infection, Immunity and Cardiovascular Disease, University of Sheffield, Sheffield S10 2TN, UK
| | - Alison M. Condliffe
- Department of Infection, Immunity and Cardiovascular Disease, University of Sheffield, Sheffield S10 2TN, UK
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9
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Henke K, Farmer DT, Niu X, Kraus JM, Galloway JL, Youngstrom DW. Genetically engineered zebrafish as models of skeletal development and regeneration. Bone 2023; 167:116611. [PMID: 36395960 PMCID: PMC11080330 DOI: 10.1016/j.bone.2022.116611] [Citation(s) in RCA: 1] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Submit a Manuscript] [Subscribe] [Scholar Register] [Received: 09/21/2022] [Revised: 11/01/2022] [Accepted: 11/08/2022] [Indexed: 11/16/2022]
Abstract
Zebrafish (Danio rerio) are aquatic vertebrates with significant homology to their terrestrial counterparts. While zebrafish have a centuries-long track record in developmental and regenerative biology, their utility has grown exponentially with the onset of modern genetics. This is exemplified in studies focused on skeletal development and repair. Herein, the numerous contributions of zebrafish to our understanding of the basic science of cartilage, bone, tendon/ligament, and other skeletal tissues are described, with a particular focus on applications to development and regeneration. We summarize the genetic strengths that have made the zebrafish a powerful model to understand skeletal biology. We also highlight the large body of existing tools and techniques available to understand skeletal development and repair in the zebrafish and introduce emerging methods that will aid in novel discoveries in skeletal biology. Finally, we review the unique contributions of zebrafish to our understanding of regeneration and highlight diverse routes of repair in different contexts of injury. We conclude that zebrafish will continue to fill a niche of increasing breadth and depth in the study of basic cellular mechanisms of skeletal biology.
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Affiliation(s)
- Katrin Henke
- Department of Orthopaedics, Department of Human Genetics, Emory University School of Medicine, Atlanta, GA 30322, USA.
| | - D'Juan T Farmer
- Department of Molecular, Cell and Developmental Biology, University of California, Los Angeles, CA 90095, USA; Department of Orthopaedic Surgery, University of California, Los Angeles, CA 90095, USA.
| | - Xubo Niu
- Center for Regenerative Medicine, Department of Orthopaedic Surgery, Massachusetts General Hospital, Harvard Medical School, Boston, MA 02114, USA.
| | - Jessica M Kraus
- Department of Orthopaedic Surgery, University of Connecticut Health Center, Farmington, CT 06030, USA.
| | - Jenna L Galloway
- Center for Regenerative Medicine, Department of Orthopaedic Surgery, Massachusetts General Hospital, Harvard Medical School, Boston, MA 02114, USA.
| | - Daniel W Youngstrom
- Department of Orthopaedic Surgery, University of Connecticut Health Center, Farmington, CT 06030, USA.
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10
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Sieliwonczyk E, Vandendriessche B, Claes C, Mayeur E, Alaerts M, Holmgren P, Canter Cremers T, Snyders D, Loeys B, Schepers D. Improved selection of zebrafish CRISPR editing by early next-generation sequencing based genotyping. Sci Rep 2023; 13:1491. [PMID: 36707549 PMCID: PMC9883431 DOI: 10.1038/s41598-023-27503-9] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/29/2022] [Accepted: 01/03/2023] [Indexed: 01/28/2023] Open
Abstract
Despite numerous prior attempts to improve knock-in (KI) efficiency, the introduction of precise base pair substitutions by the CRISPR-Cas9 technique in zebrafish remains challenging. In our efforts to generate KI zebrafish models of human CACNA1C mutations, we have tested the effect of several CRISPR determinants on KI efficiency across two sites in a single gene and developed a novel method for early selection to ameliorate KI efficiency. We identified optimal KI conditions for Cas9 protein and non-target asymmetric PAM-distal single stranded deoxynucleotide repair templates at both cacna1c sites. An effect of distance to the cut site on the KI efficiency was only observed for a single repair template conformation at one of the two sites. By combining minimally invasive early genotyping with the zebrafish embryo genotyper (ZEG) device and next-generation sequencing, we were able to obtain an almost 17-fold increase in somatic editing efficiency. The added benefit of the early selection procedure was particularly evident for alleles with lower somatic editing efficiencies. We further explored the potential of the ZEG selection procedure for the improvement of germline transmission by demonstrating germline transmission events in three groups of pre-selected embryos.
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Affiliation(s)
- Ewa Sieliwonczyk
- Faculty of Medicine and Health Sciences, Center for Medical Genetics, University of Antwerp and Antwerp University Hospital, Antwerp, Belgium.
| | - Bert Vandendriessche
- Faculty of Medicine and Health Sciences, Center for Medical Genetics, University of Antwerp and Antwerp University Hospital, Antwerp, Belgium
| | - Charlotte Claes
- Faculty of Medicine and Health Sciences, Center for Medical Genetics, University of Antwerp and Antwerp University Hospital, Antwerp, Belgium
| | - Evy Mayeur
- Experimental Neurobiology Unit, Department of Biomedical Sciences, University of Antwerp, Antwerp, Belgium
| | - Maaike Alaerts
- Faculty of Medicine and Health Sciences, Center for Medical Genetics, University of Antwerp and Antwerp University Hospital, Antwerp, Belgium
| | - Philip Holmgren
- Faculty of Medicine and Health Sciences, Center for Medical Genetics, University of Antwerp and Antwerp University Hospital, Antwerp, Belgium
| | - Tycho Canter Cremers
- Faculty of Medicine and Health Sciences, Center for Medical Genetics, University of Antwerp and Antwerp University Hospital, Antwerp, Belgium
| | - Dirk Snyders
- Experimental Neurobiology Unit, Department of Biomedical Sciences, University of Antwerp, Antwerp, Belgium
| | - Bart Loeys
- Faculty of Medicine and Health Sciences, Center for Medical Genetics, University of Antwerp and Antwerp University Hospital, Antwerp, Belgium.,Department of Clinical Genetics, Radboud University Medical Center, Nijmegen, The Netherlands
| | - Dorien Schepers
- Faculty of Medicine and Health Sciences, Center for Medical Genetics, University of Antwerp and Antwerp University Hospital, Antwerp, Belgium.,Experimental Neurobiology Unit, Department of Biomedical Sciences, University of Antwerp, Antwerp, Belgium
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11
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Moreno-Nombela S, Romero-Parra J, Ruiz-Ojeda FJ, Solis-Urra P, Baig AT, Plaza-Diaz J. Genome Editing and Protein Energy Malnutrition. ADVANCES IN EXPERIMENTAL MEDICINE AND BIOLOGY 2023; 1396:215-232. [DOI: 10.1007/978-981-19-5642-3_15] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 12/03/2022]
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12
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A robust pipeline for efficient knock-in of point mutations and epitope tags in zebrafish using fluorescent PCR based screening. BMC Genomics 2022; 23:810. [PMID: 36476416 PMCID: PMC9730659 DOI: 10.1186/s12864-022-08971-1] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/23/2022] [Accepted: 10/26/2022] [Indexed: 12/12/2022] Open
Abstract
BACKGROUND Genome editing using CRISPR/Cas9 has become a powerful tool in zebrafish to generate targeted gene knockouts models. However, its use for targeted knock-in remains challenging due to inefficient homology directed repair (HDR) pathway in zebrafish, highlighting the need for efficient and cost-effective screening methods. RESULTS: Here, we present our fluorescent PCR and capillary electrophoresis based screening approach for knock-in using a single-stranded oligodeoxynucleotide donor (ssODN) as a repair template for the targeted insertion of epitope tags, or single nucleotide changes to recapitulate pathogenic human alleles. For the insertion of epitope tags, we took advantage of the expected change in size of the PCR product. For point mutations, we combined fluorescent PCR with restriction fragment length polymorphism (RFLP) analysis to distinguish the fish with the knock-in allele. As a proof-of-principle, we present our data on the generation of fish lines with insertion of a FLAG tag at the tcnba locus, an HA tag at the gata2b locus, and a point mutation observed in Gaucher disease patients in the gba gene. Despite the low number of germline transmitting founders (1-5%), combining our screening methods with prioritization of founder fish by fin biopsies allowed us to establish stable knock-in lines by screening 12 or less fish per gene. CONCLUSIONS We have established a robust pipeline for the generation of zebrafish models with precise integration of small DNA sequences and point mutations at the desired sites in the genome. Our screening method is very efficient and easy to implement as it is PCR-based and only requires access to a capillary sequencer.
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13
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SpG and SpRY variants expand the CRISPR toolbox for genome editing in zebrafish. Nat Commun 2022; 13:3421. [PMID: 35701400 PMCID: PMC9198057 DOI: 10.1038/s41467-022-31034-8] [Citation(s) in RCA: 9] [Impact Index Per Article: 4.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/06/2021] [Accepted: 05/31/2022] [Indexed: 11/27/2022] Open
Abstract
Precise genetic modifications in model organisms are essential for biomedical research. The recent development of PAM-less base editors makes it possible to assess the functional impact and pathogenicity of nucleotide mutations in animals. Here we first optimize SpG and SpRY systems in zebrafish by purifying protein combined with synthetically modified gRNA. SpG shows high editing efficiency at NGN PAM sites, whereas SpRY efficiently edit PAM-less sites in the zebrafish genome. Then, we generate the SpRY-mediated cytosine base editor SpRY-CBE4max and SpRY-mediated adenine base editor zSpRY-ABE8e. Both target relaxed PAM with up to 96% editing efficiency and high product purity. With these tools, some previously inaccessible disease-relevant genetic variants are generated in zebrafish, supporting the utility of high-resolution targeting across genome-editing applications. Our study significantly improves CRISPR-Cas targeting in the genomic landscape of zebrafish, promoting the application of this model organism in revealing gene function, physiological mechanisms, and disease pathogenesis.
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14
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Abstract
Heart disease is the leading cause of death worldwide. Despite decades of research, most heart pathologies have limited treatments, and often the only curative approach is heart transplantation. Thus, there is an urgent need to develop new therapeutic approaches for treating cardiac diseases. Animal models that reproduce the human pathophysiology are essential to uncovering the biology of diseases and discovering therapies. Traditionally, mammals have been used as models of cardiac disease, but the cost of generating and maintaining new models is exorbitant, and the studies have very low throughput. In the last decade, the zebrafish has emerged as a tractable model for cardiac diseases, owing to several characteristics that made this animal popular among developmental biologists. Zebrafish fertilization and development are external; embryos can be obtained in high numbers, are cheap and easy to maintain, and can be manipulated to create new genetic models. Moreover, zebrafish exhibit an exceptional ability to regenerate their heart after injury. This review summarizes 25 years of research using the zebrafish to study the heart, from the classical forward screenings to the contemporary methods to model mutations found in patients with cardiac disease. We discuss the advantages and limitations of this model organism and introduce the experimental approaches exploited in zebrafish, including forward and reverse genetics and chemical screenings. Last, we review the models used to induce cardiac injury and essential ideas derived from studying natural regeneration. Studies using zebrafish have the potential to accelerate the discovery of new strategies to treat cardiac diseases.
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Affiliation(s)
- Juan Manuel González-Rosa
- Cardiovascular Research Center, Massachusetts General Hospital Research Institute, Harvard Medical School, Charlestown, MA
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15
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Volodina OV, Smirnikhina SA. The Choice of a Donor Molecule in Genome Editing Experiments in Animal Cells. Mol Biol 2022. [DOI: 10.1134/s002689332203013x] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/23/2022]
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16
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Sansbury BM, Hewes AM, Tharp OM, Masciarelli SB, Kaouser S, Kmiec EB. Homology directed correction, a new pathway model for point mutation repair catalyzed by CRISPR-Cas. Sci Rep 2022; 12:8132. [PMID: 35581233 PMCID: PMC9114366 DOI: 10.1038/s41598-022-11808-2] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/10/2021] [Accepted: 04/28/2022] [Indexed: 11/09/2022] Open
Abstract
Gene correction is often referred to as the gold standard for precise gene editing and while CRISPR-Cas systems continue to expand the toolbox for clinically relevant genetic repair, mechanistic hurdles still hinder widespread implementation. One of the most prominent challenges to precise CRISPR-directed point mutation repair centers on the prevalence of on-site mutagenesis, wherein insertions and deletions appear at the targeted site following correction. Here, we introduce a pathway model for Homology Directed Correction, specifically point mutation repair, which enables a foundational analysis of genetic tools and factors influencing precise gene editing. To do this, we modified an in vitro gene editing system which utilizes a cell-free extract, CRISPR-Cas RNP and donor DNA template to catalyze point mutation repair. We successfully direct correction of four unique point mutations which include two unique nucleotide mutations at two separate targeted sites and visualize the repair profiles resulting from these reactions. This extension of the cell-free gene editing system to model point mutation repair may provide insight for understanding the factors influencing precise point mutation correction.
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Affiliation(s)
- Brett M Sansbury
- Gene Editing Institute, ChristianaCare Health System, 550 S College Ave, Suite 100A, 2nd Floor, Newark, DE, 19713, USA
| | - Amanda M Hewes
- Gene Editing Institute, ChristianaCare Health System, 550 S College Ave, Suite 100A, 2nd Floor, Newark, DE, 19713, USA
| | - Olivia M Tharp
- Gene Editing Institute, ChristianaCare Health System, 550 S College Ave, Suite 100A, 2nd Floor, Newark, DE, 19713, USA.,Department of Medical and Molecular Sciences, University of Delaware, Newark, DE, USA
| | - Sophia B Masciarelli
- Gene Editing Institute, ChristianaCare Health System, 550 S College Ave, Suite 100A, 2nd Floor, Newark, DE, 19713, USA.,Department of Medical and Molecular Sciences, University of Delaware, Newark, DE, USA
| | - Salma Kaouser
- Gene Editing Institute, ChristianaCare Health System, 550 S College Ave, Suite 100A, 2nd Floor, Newark, DE, 19713, USA
| | - Eric B Kmiec
- Gene Editing Institute, ChristianaCare Health System, 550 S College Ave, Suite 100A, 2nd Floor, Newark, DE, 19713, USA.
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17
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Clark B, Elkin J, Marconi A, Turner GF, Smith AM, Joyce D, Miska EA, Juntti SA, Santos ME. Oca2 targeting using CRISPR/Cas9 in the Malawi cichlid Astatotilapia calliptera. ROYAL SOCIETY OPEN SCIENCE 2022; 9:220077. [PMID: 35601449 PMCID: PMC9019512 DOI: 10.1098/rsos.220077] [Citation(s) in RCA: 2] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Figures] [Subscribe] [Scholar Register] [Received: 01/20/2022] [Accepted: 03/22/2022] [Indexed: 05/03/2023]
Abstract
Identifying genetic loci underlying trait variation provides insights into the mechanisms of diversification, but demonstrating causality and characterizing the role of genetic loci requires testing candidate gene function, often in non-model species. Here we establish CRISPR/Cas9 editing in Astatotilapia calliptera, a generalist cichlid of the remarkably diverse Lake Malawi radiation. By targeting the gene oca2 required for melanin synthesis in other vertebrate species, we show efficient editing and germline transmission. Gene edits include indels in the coding region, probably a result of non-homologous end joining, and a large deletion in the 3' untranslated region due to homology-directed repair. We find that oca2 knock-out A. calliptera lack melanin, which may be useful for developmental imaging in embryos and studying colour pattern formation in adults. As A. calliptera resembles the presumed generalist ancestor of the Lake Malawi cichlids radiation, establishing genome editing in this species will facilitate investigating speciation, adaptation and trait diversification in this textbook radiation.
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Affiliation(s)
- Bethan Clark
- Department of Zoology, University of Cambridge, UK
| | - Joel Elkin
- Department of Zoology, University of Cambridge, UK
| | | | - George F. Turner
- School of Natural Sciences, Bangor University, Gwynedd LL57 2TH, UK
| | - Alan M. Smith
- Department of Biological and Marine Sciences, University of Hull, UK
| | - Domino Joyce
- Department of Biological and Marine Sciences, University of Hull, UK
| | - Eric A. Miska
- Department of Genetics, University of Cambridge, UK
- Gurdon Institute, University of Cambridge, Cambridge CB2 1QN, UK
- Wellcome Sanger Institute, Wellcome Trust Genome Campus, Cambridge, UK
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18
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Simone BW, Lee HB, Daby CL, Ata H, Restrepo-Castillo S, Martínez-Gálvez G, Kar B, Gendron WA, Clark KJ, Ekker SC. Chimeric RNA: DNA TracrRNA Improves Homology-Directed Repair In Vitro and In Vivo. CRISPR J 2022; 5:40-52. [PMID: 34935462 PMCID: PMC8892967 DOI: 10.1089/crispr.2021.0087] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.5] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/17/2023] Open
Abstract
Nearly 90% of human pathogenic mutations are caused by small genetic variations, and methods to correct these errors efficiently are critically important. One way to make small DNA changes is providing a single-stranded oligo deoxynucleotide (ssODN) containing an alteration coupled with a targeted double-strand break (DSB) at the target locus in the genome. Coupling an ssODN donor with a CRISPR-Cas9-mediated DSB is one of the most streamlined approaches to introduce small changes. However, in many systems, this approach is inefficient and introduces imprecise repair at the genetic junctions. We herein report a technology that uses spatiotemporal localization of an ssODN with CRISPR-Cas9 to improve gene alteration. We show that by fusing an ssODN template to the trans-activating RNA (tracrRNA), we recover precise genetic alterations, with increased integration and precision in vitro and in vivo. Finally, we show that this technology can be used to enhance gene conversion with other gene editing tools such as transcription activator like effector nucleases.
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Affiliation(s)
- Brandon W. Simone
- Department of Biochemistry and Molecular Biology, Biomedical Engineering and Physiology Track, Mayo Clinic, Rochester, Minnesota, USA
| | - Han B. Lee
- Department of Biochemistry and Molecular Biology, Biomedical Engineering and Physiology Track, Mayo Clinic, Rochester, Minnesota, USA
| | - Camden L. Daby
- Department of Biochemistry and Molecular Biology, Biomedical Engineering and Physiology Track, Mayo Clinic, Rochester, Minnesota, USA
| | - Hirotaka Ata
- Department of Clinical and Translational Sciences, Biomedical Engineering and Physiology Track, Mayo Clinic, Rochester, Minnesota, USA
| | - Santiago Restrepo-Castillo
- Mayo Clinic Graduate School of Biomedical Sciences, Virology and Gene Therapy Track, Biomedical Engineering and Physiology Track, Mayo Clinic, Rochester, Minnesota, USA
| | - Gabriel Martínez-Gálvez
- Mayo Clinic Graduate School of Biomedical Sciences, Biomedical Engineering and Physiology Track, Mayo Clinic, Rochester, Minnesota, USA
| | - Bibekananda Kar
- Department of Biochemistry and Molecular Biology, Biomedical Engineering and Physiology Track, Mayo Clinic, Rochester, Minnesota, USA
| | - William A.C. Gendron
- Mayo Clinic Graduate School of Biomedical Sciences, Virology and Gene Therapy Track, Biomedical Engineering and Physiology Track, Mayo Clinic, Rochester, Minnesota, USA
| | - Karl J. Clark
- Department of Biochemistry and Molecular Biology, Biomedical Engineering and Physiology Track, Mayo Clinic, Rochester, Minnesota, USA
| | - Stephen C. Ekker
- Department of Biochemistry and Molecular Biology, Biomedical Engineering and Physiology Track, Mayo Clinic, Rochester, Minnesota, USA
- Address correspondence to: Stephen C. Ekker, PhD, Department of Biochemistry and Molecular Biology, Mayo Clinic, 200 1st Street SW, Rochester, MN 55905, USA,
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19
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Tan J, Wang Y, Chen S, Lin Z, Zhao Y, Xue Y, Luo Y, Liu YG, Zhu Q. An Efficient Marker Gene Excision Strategy Based on CRISPR/Cas9-Mediated Homology-Directed Repair in Rice. Int J Mol Sci 2022; 23:ijms23031588. [PMID: 35163510 PMCID: PMC8835944 DOI: 10.3390/ijms23031588] [Citation(s) in RCA: 2] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/29/2021] [Revised: 01/26/2022] [Accepted: 01/27/2022] [Indexed: 01/05/2023] Open
Abstract
In order to separate transformed cells from non-transformed cells, antibiotic selectable marker genes are usually utilized in genetic transformation. After obtaining transgenic plants, it is often necessary to remove the marker gene from the plant genome in order to avoid regulatory issues. However, many marker-free systems are time-consuming and labor-intensive. Homology-directed repair (HDR) is a process of homologous recombination using homologous arms for efficient and precise repair of DNA double-strand breaks (DSBs). The clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein-9 (Cas9) system is a powerful genome editing tool that can efficiently cause DSBs. Here, we isolated a rice promoter (Pssi) of a gene that highly expressed in stem, shoot tip and inflorescence, and established a high-efficiency sequence-excision strategy by using this Pssi to drive CRISPR/Cas9-mediated HDR for marker free (PssiCHMF). In our study, PssiCHMF-induced marker gene deletion was detected in 73.3% of T0 plants and 83.2% of T1 plants. A high proportion (55.6%) of homozygous marker-excised plants were obtained in T1 progeny. The recombinant GUS reporter-aided analysis and its sequencing of the recombinant products showed precise deletion and repair mediated by the PssiCHMF method. In conclusion, our CRISPR/Cas9-mediated HDR auto-excision method provides a time-saving and efficient strategy for removing the marker genes from transgenic plants.
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Affiliation(s)
- Jiantao Tan
- State Key Laboratory for Conservation and Utilization of Subtropical Agro-Bioresources, South China Agricultural University, Guangzhou 510642, China; (J.T.); (Y.W.); (S.C.); (Z.L.); (Y.Z.); (Y.X.); (Y.L.); (Y.-G.L.)
- Guangdong Laboratory for Lingnan Modern Agriculture, Guangzhou 510642, China
- College of Life Sciences, South China Agricultural University, Guangzhou 510642, China
| | - Yaxi Wang
- State Key Laboratory for Conservation and Utilization of Subtropical Agro-Bioresources, South China Agricultural University, Guangzhou 510642, China; (J.T.); (Y.W.); (S.C.); (Z.L.); (Y.Z.); (Y.X.); (Y.L.); (Y.-G.L.)
- College of Life Sciences, South China Agricultural University, Guangzhou 510642, China
| | - Shuifu Chen
- State Key Laboratory for Conservation and Utilization of Subtropical Agro-Bioresources, South China Agricultural University, Guangzhou 510642, China; (J.T.); (Y.W.); (S.C.); (Z.L.); (Y.Z.); (Y.X.); (Y.L.); (Y.-G.L.)
- College of Life Sciences, South China Agricultural University, Guangzhou 510642, China
| | - Zhansheng Lin
- State Key Laboratory for Conservation and Utilization of Subtropical Agro-Bioresources, South China Agricultural University, Guangzhou 510642, China; (J.T.); (Y.W.); (S.C.); (Z.L.); (Y.Z.); (Y.X.); (Y.L.); (Y.-G.L.)
- College of Life Sciences, South China Agricultural University, Guangzhou 510642, China
| | - Yanchang Zhao
- State Key Laboratory for Conservation and Utilization of Subtropical Agro-Bioresources, South China Agricultural University, Guangzhou 510642, China; (J.T.); (Y.W.); (S.C.); (Z.L.); (Y.Z.); (Y.X.); (Y.L.); (Y.-G.L.)
- College of Life Sciences, South China Agricultural University, Guangzhou 510642, China
| | - Yang Xue
- State Key Laboratory for Conservation and Utilization of Subtropical Agro-Bioresources, South China Agricultural University, Guangzhou 510642, China; (J.T.); (Y.W.); (S.C.); (Z.L.); (Y.Z.); (Y.X.); (Y.L.); (Y.-G.L.)
- College of Life Sciences, South China Agricultural University, Guangzhou 510642, China
| | - Yuyu Luo
- State Key Laboratory for Conservation and Utilization of Subtropical Agro-Bioresources, South China Agricultural University, Guangzhou 510642, China; (J.T.); (Y.W.); (S.C.); (Z.L.); (Y.Z.); (Y.X.); (Y.L.); (Y.-G.L.)
- College of Life Sciences, South China Agricultural University, Guangzhou 510642, China
| | - Yao-Guang Liu
- State Key Laboratory for Conservation and Utilization of Subtropical Agro-Bioresources, South China Agricultural University, Guangzhou 510642, China; (J.T.); (Y.W.); (S.C.); (Z.L.); (Y.Z.); (Y.X.); (Y.L.); (Y.-G.L.)
- Guangdong Laboratory for Lingnan Modern Agriculture, Guangzhou 510642, China
- College of Life Sciences, South China Agricultural University, Guangzhou 510642, China
| | - Qinlong Zhu
- State Key Laboratory for Conservation and Utilization of Subtropical Agro-Bioresources, South China Agricultural University, Guangzhou 510642, China; (J.T.); (Y.W.); (S.C.); (Z.L.); (Y.Z.); (Y.X.); (Y.L.); (Y.-G.L.)
- Guangdong Laboratory for Lingnan Modern Agriculture, Guangzhou 510642, China
- College of Life Sciences, South China Agricultural University, Guangzhou 510642, China
- Correspondence:
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20
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Murakami Y, Kobayashi T. An effective double gene knock‐in strategy using small‐molecule
L755507
in the medaka fish (
Oryzias latipes
). Genesis 2022; 60:e23465. [DOI: 10.1002/dvg.23465] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/24/2021] [Revised: 01/05/2022] [Accepted: 01/06/2022] [Indexed: 11/07/2022]
Affiliation(s)
- Yu Murakami
- Department of Fisheries, Graduate School of Agriculture Kindai University Nara Japan
| | - Toru Kobayashi
- Department of Fisheries, Graduate School of Agriculture Kindai University Nara Japan
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21
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Nambiar TS, Baudrier L, Billon P, Ciccia A. CRISPR-based genome editing through the lens of DNA repair. Mol Cell 2022; 82:348-388. [PMID: 35063100 PMCID: PMC8887926 DOI: 10.1016/j.molcel.2021.12.026] [Citation(s) in RCA: 66] [Impact Index Per Article: 33.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/15/2021] [Revised: 12/18/2021] [Accepted: 12/20/2021] [Indexed: 01/22/2023]
Abstract
Genome editing technologies operate by inducing site-specific DNA perturbations that are resolved by cellular DNA repair pathways. Products of genome editors include DNA breaks generated by CRISPR-associated nucleases, base modifications induced by base editors, DNA flaps created by prime editors, and integration intermediates formed by site-specific recombinases and transposases associated with CRISPR systems. Here, we discuss the cellular processes that repair CRISPR-generated DNA lesions and describe strategies to obtain desirable genomic changes through modulation of DNA repair pathways. Advances in our understanding of the DNA repair circuitry, in conjunction with the rapid development of innovative genome editing technologies, promise to greatly enhance our ability to improve food production, combat environmental pollution, develop cell-based therapies, and cure genetic and infectious diseases.
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Affiliation(s)
- Tarun S. Nambiar
- Department of Genetics and Development, Herbert Irving Comprehensive Cancer Center, Columbia University Irving Medical Center, New York, NY 10032
| | - Lou Baudrier
- Department of Biochemistry and Molecular Biology, Robson DNA Science Centre, Arnie Charbonneau Cancer Institute, Cumming School of Medicine, University of Calgary, 3330 Hospital Drive N. W., Calgary, Alberta T2N 4N1, Canada
| | - Pierre Billon
- Department of Biochemistry and Molecular Biology, Robson DNA Science Centre, Arnie Charbonneau Cancer Institute, Cumming School of Medicine, University of Calgary, 3330 Hospital Drive N. W., Calgary, Alberta T2N 4N1, Canada,Corresponding authors: ,
| | - Alberto Ciccia
- Department of Genetics and Development, Herbert Irving Comprehensive Cancer Center, Columbia University Irving Medical Center, New York, NY 10032,Lead Contact,Corresponding authors: ,
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22
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Papasavva PL, Patsali P, Loucari CC, Kurita R, Nakamura Y, Kleanthous M, Lederer CW. CRISPR Editing Enables Consequential Tag-Activated MicroRNA-Mediated Endogene Deactivation. Int J Mol Sci 2022; 23:1082. [PMID: 35163006 PMCID: PMC8834719 DOI: 10.3390/ijms23031082] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.5] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/19/2021] [Revised: 01/09/2022] [Accepted: 01/12/2022] [Indexed: 02/01/2023] Open
Abstract
Molecular therapies and functional studies greatly benefit from spatial and temporal precision of genetic intervention. We therefore conceived and explored tag-activated microRNA (miRNA)-mediated endogene deactivation (TAMED) as a research tool and potential lineage-specific therapy. For proof of principle, we aimed to deactivate γ-globin repressor BCL11A in erythroid cells by tagging the 3' untranslated region (UTR) of BCL11A with miRNA recognition sites (MRSs) for the abundant erythromiR miR-451a. To this end, we employed nucleofection of CRISPR/Cas9 ribonucleoprotein (RNP) particles alongside double- or single-stranded oligodeoxynucleotides for, respectively, non-homologous-end-joining (NHEJ)- or homology-directed-repair (HDR)-mediated MRS insertion. NHEJ-based tagging was imprecise and inefficient (≤6%) and uniformly produced knock-in- and indel-containing MRS tags, whereas HDR-based tagging was more efficient (≤18%), but toxic for longer donors encoding concatenated and thus potentially more efficient MRS tags. Isolation of clones for robust HEK293T cells tagged with a homozygous quadruple MRS resulted in 25% spontaneous reduction in BCL11A and up to 36% reduction after transfection with an miR-451a mimic. Isolation of clones for human umbilical cord blood-derived erythroid progenitor-2 (HUDEP-2) cells tagged with single or double MRS allowed detection of albeit weak γ-globin induction. Our study demonstrates suitability of TAMED for physiologically relevant modulation of gene expression and its unsuitability for therapeutic application in its current form.
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Affiliation(s)
- Panayiota L. Papasavva
- Department of Molecular Genetics Thalassemia, The Cyprus Institute of Neurology and Genetics, Nicosia 2371, Cyprus; (P.L.P.); (P.P.); (C.C.L.); (M.K.)
- Cyprus School of Molecular Medicine, Nicosia 2371, Cyprus
| | - Petros Patsali
- Department of Molecular Genetics Thalassemia, The Cyprus Institute of Neurology and Genetics, Nicosia 2371, Cyprus; (P.L.P.); (P.P.); (C.C.L.); (M.K.)
- Cyprus School of Molecular Medicine, Nicosia 2371, Cyprus
| | - Constantinos C. Loucari
- Department of Molecular Genetics Thalassemia, The Cyprus Institute of Neurology and Genetics, Nicosia 2371, Cyprus; (P.L.P.); (P.P.); (C.C.L.); (M.K.)
- Cyprus School of Molecular Medicine, Nicosia 2371, Cyprus
| | - Ryo Kurita
- Research and Development Department, Central Blood Institute, Blood Service Headquarters, Japanese Red Cross Society, Koto-ku, Tokyo 135-8521, Japan;
| | - Yukio Nakamura
- Cell Engineering Division, RIKEN BioResource Research Center, Tsukuba 305-0074, Japan;
| | - Marina Kleanthous
- Department of Molecular Genetics Thalassemia, The Cyprus Institute of Neurology and Genetics, Nicosia 2371, Cyprus; (P.L.P.); (P.P.); (C.C.L.); (M.K.)
- Cyprus School of Molecular Medicine, Nicosia 2371, Cyprus
| | - Carsten W. Lederer
- Department of Molecular Genetics Thalassemia, The Cyprus Institute of Neurology and Genetics, Nicosia 2371, Cyprus; (P.L.P.); (P.P.); (C.C.L.); (M.K.)
- Cyprus School of Molecular Medicine, Nicosia 2371, Cyprus
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23
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Kumari P, Sturgeon M, Bonde G, Cornell RA. Generating Zebrafish RNA-Less Mutant Alleles by Deleting Gene Promoters with CRISPR/Cas9. Methods Mol Biol 2022; 2403:91-106. [PMID: 34913119 PMCID: PMC10136374 DOI: 10.1007/978-1-0716-1847-9_8] [Citation(s) in RCA: 2] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 10/19/2022]
Abstract
Danio rerio (zebrafish), traditionally used in forward genetic screens, has in the last decade become a popular model for reverse genetic studies with the introduction of TALENS, zinc finger nucleases, and CRISPR/Cas9. Unexpectedly, homozygous frameshift mutations generated by these tools frequently result in phenotypes that are less penetrant than those seen in embryos injected with antisense morpholino oligonucleotides targeting the same gene. One explanation for the difference is that some frameshift mutations result in nonsense-mediated decay of the gene transcript, a process which can induce expression of homologous genes. This form of genetic compensation, called transcriptional adaptation, does not occur when the mutant allele results in no RNA transcripts being produced from the targeted gene. Such RNA-less mutants can be generated by deleting a gene's promoter using a pair of guide RNAs and Cas9 protein. Here, we present a protocol and use it to generate alleles of arhgap29b and slc41a1 that lack detectable zygotic transcription. In the case of the arhgap29b mutant, an emerging phenotype did not segregate with the promoter deletion mutation, highlighting the potential for off-target mutagenesis with these tools. In summary, this chapter describes a method to generate zebrafish mutants that avoid a form of genetic compensation that occurs in many frameshift mutants.
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Affiliation(s)
- Priyanka Kumari
- Department of Anatomy and Cell Biology, University of Iowa, Iowa City, IA, USA
| | - Morgan Sturgeon
- Department of Anatomy and Cell Biology, University of Iowa, Iowa City, IA, USA
- Integrated DNA Technologies, Coralville, IA, USA
| | - Gregory Bonde
- Department of Anatomy and Cell Biology, University of Iowa, Iowa City, IA, USA
| | - Robert A Cornell
- Department of Anatomy and Cell Biology, University of Iowa, Iowa City, IA, USA.
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24
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Smirnov A, Battulin N. Concatenation of Transgenic DNA: Random or Orchestrated? Genes (Basel) 2021; 12:genes12121969. [PMID: 34946918 PMCID: PMC8701086 DOI: 10.3390/genes12121969] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.7] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/10/2021] [Revised: 12/08/2021] [Accepted: 12/09/2021] [Indexed: 12/18/2022] Open
Abstract
Generation of transgenic organisms by pronuclear microinjection has become a routine procedure. However, while the process of DNA integration in the genome is well understood, we still do not know much about the recombination between transgene molecules that happens in the first moments after DNA injection. Most of the time, injected molecules are joined together in head-to-tail tandem repeats-the so-called concatemers. In this review, we focused on the possible concatenation mechanisms and how they could be studied with genetic reporters tracking individual copies in concatemers. We also discuss various features of concatemers, including palindromic junctions and repeat-induced gene silencing (RIGS). Finally, we speculate how cooperation of DNA repair pathways creates a multicopy concatenated insert.
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Affiliation(s)
- Alexander Smirnov
- Laboratory of Developmental Genetics, Institute of Cytology and Genetics SB RAS, 630090 Novosibirsk, Russia;
| | - Nariman Battulin
- Laboratory of Developmental Genetics, Institute of Cytology and Genetics SB RAS, 630090 Novosibirsk, Russia;
- Institute of Genetic Technologies, Novosibirsk State University, 630090 Novosibirsk, Russia
- Correspondence:
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Eini O, Schumann N, Niessen M, Varrelmann M. Targeted mutagenesis in plants using Beet curly top virus for efficient delivery of CRISPR/Cas12a components. N Biotechnol 2021; 67:1-11. [PMID: 34896246 DOI: 10.1016/j.nbt.2021.12.002] [Citation(s) in RCA: 8] [Impact Index Per Article: 2.7] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/19/2021] [Revised: 11/29/2021] [Accepted: 12/05/2021] [Indexed: 10/19/2022]
Abstract
Genome editing using CRISPR/Cas is rapidly being developed for gene targeting in eukaryotes including plants. However, gene targeting by homology-directed DNA recombination (HDR) is an infrequent event compared to the dominant DNA repair by non-homologous end-joining. Another bottleneck is the ineffective delivery of CRISPR/Cas components into plant cells. To overcome these constraints, here a geminiviral replicon from Beet curly top virus (BCTV) has been produced with a wide host range and high DNA accumulation capacity for efficient delivery of CRISPR/Cas12a components into plant cells. Initially, a BCTV replicon was prepared after removing the virion sense genes from an infectious full-length clone for agrobacterium mediated infection. This replicon expressed a green fluorescent protein (GFP) marker gene at a high level compared to T-DNA binary vector. In transient assay, the BCTV replicon produced a higher rate of mutagenesis and HDR in the GFP transgene in Nicotiana benthamiana through efficient delivery of CRISPR/Cas12a components compared to the cognate T-DNA control. This was through a range of complete or partial HDR for conversion of GFP into YFP after exchange of a single amino acid (Thr224Tyr) in the target gene. In addition, induced mutagenesis and HDR in the target gene were heritable. Thus, the BCTV replicon provides a new tool for efficient delivery of CRISPR/Cas12a components that could be used in a wide range of dicotyledonous plants. The established GFP to YFP system and the GFP mutant line produced also enable further optimization and understanding of HDR in plants via CRISPR/Cas12a system using geminiviral replicons.
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Affiliation(s)
- Omid Eini
- Department of Plant Protection, University of Zanjan, 313, Zanjan, Iran; Department of Phytopathology, Institute for Sugar Beet Research, 37079, Göttingen, Germany.
| | | | | | - Mark Varrelmann
- Department of Phytopathology, Institute for Sugar Beet Research, 37079, Göttingen, Germany
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Akella S, Ma X, Bacova R, Harmer ZP, Kolackova M, Wen X, Wright DA, Spalding MH, Weeks DP, Cerutti H. Co-targeting strategy for precise, scarless gene editing with CRISPR/Cas9 and donor ssODNs in Chlamydomonas. PLANT PHYSIOLOGY 2021; 187:2637-2655. [PMID: 34618092 PMCID: PMC8644747 DOI: 10.1093/plphys/kiab418] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 03/26/2021] [Accepted: 07/30/2021] [Indexed: 05/20/2023]
Abstract
Programmable site-specific nucleases, such as the clustered regularly interspaced short palindromic repeat (CRISPR)/ CRISPR-associated protein 9 (Cas9) ribonucleoproteins (RNPs), have allowed creation of valuable knockout mutations and targeted gene modifications in Chlamydomonas (Chlamydomonas reinhardtii). However, in walled strains, present methods for editing genes lacking a selectable phenotype involve co-transfection of RNPs and exogenous double-stranded DNA (dsDNA) encoding a selectable marker gene. Repair of the dsDNA breaks induced by the RNPs is usually accompanied by genomic insertion of exogenous dsDNA fragments, hindering the recovery of precise, scarless mutations in target genes of interest. Here, we tested whether co-targeting two genes by electroporation of pairs of CRISPR/Cas9 RNPs and single-stranded oligodeoxynucleotides (ssODNs) would facilitate the recovery of precise edits in a gene of interest (lacking a selectable phenotype) by selection for precise editing of another gene (creating a selectable marker)-in a process completely lacking exogenous dsDNA. We used PPX1 (encoding protoporphyrinogen IX oxidase) as the generated selectable marker, conferring resistance to oxyfluorfen, and identified precise edits in the homolog of bacterial ftsY or the WD and TetratriCopeptide repeats protein 1 genes in ∼1% of the oxyfluorfen resistant colonies. Analysis of the target site sequences in edited mutants suggested that ssODNs were used as templates for DNA synthesis during homology directed repair, a process prone to replicative errors. The Chlamydomonas acetolactate synthase gene could also be efficiently edited to serve as an alternative selectable marker. This transgene-free strategy may allow creation of individual strains containing precise mutations in multiple target genes, to study complex cellular processes, pathways, or structures.
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Affiliation(s)
- Soujanya Akella
- School of Biological Sciences and Center for Plant Science Innovation, University of Nebraska–Lincoln, Lincoln, Nebraska 68588, USA
| | - Xinrong Ma
- School of Biological Sciences and Center for Plant Science Innovation, University of Nebraska–Lincoln, Lincoln, Nebraska 68588, USA
| | - Romana Bacova
- Department of Chemistry and Biochemistry, Mendel University in Brno, Zemedelska 1, CZ-613 00, Brno, Czech Republic
| | - Zachary P Harmer
- School of Biological Sciences and Center for Plant Science Innovation, University of Nebraska–Lincoln, Lincoln, Nebraska 68588, USA
| | - Martina Kolackova
- Department of Chemistry and Biochemistry, Mendel University in Brno, Zemedelska 1, CZ-613 00, Brno, Czech Republic
| | - Xiaoxue Wen
- School of Biological Sciences and Center for Plant Science Innovation, University of Nebraska–Lincoln, Lincoln, Nebraska 68588, USA
| | - David A Wright
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa 50011, USA
| | - Martin H Spalding
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa 50011, USA
| | - Donald P Weeks
- Department of Biochemistry, University of Nebraska-Lincoln, Lincoln, Nebraska 68588, USA
| | - Heriberto Cerutti
- School of Biological Sciences and Center for Plant Science Innovation, University of Nebraska–Lincoln, Lincoln, Nebraska 68588, USA
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Progress in Gene-Editing Technology of Zebrafish. Biomolecules 2021; 11:biom11091300. [PMID: 34572513 PMCID: PMC8468279 DOI: 10.3390/biom11091300] [Citation(s) in RCA: 12] [Impact Index Per Article: 4.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/07/2021] [Revised: 08/25/2021] [Accepted: 08/30/2021] [Indexed: 12/26/2022] Open
Abstract
As a vertebrate model, zebrafish (Danio rerio) plays a vital role in the field of life sciences. Recently, gene-editing technology has become increasingly innovative, significantly promoting scientific research on zebrafish. However, the implementation of these methods in a reasonable and accurate manner to achieve efficient gene-editing remains challenging. In this review, we systematically summarize the development and latest progress in zebrafish gene-editing technology. Specifically, we outline trends in double-strand break-free genome modification and the prospective applications of fixed-point orientation transformation of any base at any location through a multi-method approach.
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de Vrieze E, de Bruijn SE, Reurink J, Broekman S, van de Riet V, Aben M, Kremer H, van Wijk E. Efficient Generation of Knock-In Zebrafish Models for Inherited Disorders Using CRISPR-Cas9 Ribonucleoprotein Complexes. Int J Mol Sci 2021; 22:9429. [PMID: 34502338 PMCID: PMC8431507 DOI: 10.3390/ijms22179429] [Citation(s) in RCA: 11] [Impact Index Per Article: 3.7] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/30/2021] [Revised: 08/26/2021] [Accepted: 08/27/2021] [Indexed: 12/17/2022] Open
Abstract
CRISPR-Cas9-based genome-editing is a highly efficient and cost-effective method to generate zebrafish loss-of-function alleles. However, introducing patient-specific variants into the zebrafish genome with CRISPR-Cas9 remains challenging. Targeting options can be limited by the predetermined genetic context, and the efficiency of the homology-directed DNA repair pathway is relatively low. Here, we illustrate our efficient approach to develop knock-in zebrafish models using two previously variants associated with hereditary sensory deficits. We employ sgRNA-Cas9 ribonucleoprotein (RNP) complexes that are micro-injected into the first cell of fertilized zebrafish eggs together with an asymmetric, single-stranded DNA template containing the variant of interest. The introduction of knock-in events was confirmed by massive parallel sequencing of genomic DNA extracted from a pool of injected embryos. Simultaneous morpholino-induced blocking of a key component of the non-homologous end joining DNA repair pathway, Ku70, improved the knock-in efficiency for one of the targets. Our use of RNP complexes provides an improved knock-in efficiency as compared to previously published studies. Correct knock-in events were identified in 3-8% of alleles, and 30-45% of injected animals had the target variant in their germline. The detailed technical and procedural insights described here provide a valuable framework for the efficient development of knock-in zebrafish models.
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Affiliation(s)
- Erik de Vrieze
- Department of Otorhinolaryngology, Radboud University Medical Center, 6525 GA Nijmegen, The Netherlands; (S.B.); (V.v.d.R.); (H.K.); (E.v.W.)
- Donders Institute for Brain Cognition and Behaviour, 6500 GL Nijmegen, The Netherlands; (S.E.d.B.); (J.R.); (M.A.)
| | - Suzanne E. de Bruijn
- Donders Institute for Brain Cognition and Behaviour, 6500 GL Nijmegen, The Netherlands; (S.E.d.B.); (J.R.); (M.A.)
- Department of Human Genetics, Radboud University Medical Center, 6525 GA Nijmegen, The Netherlands
| | - Janine Reurink
- Donders Institute for Brain Cognition and Behaviour, 6500 GL Nijmegen, The Netherlands; (S.E.d.B.); (J.R.); (M.A.)
- Department of Human Genetics, Radboud University Medical Center, 6525 GA Nijmegen, The Netherlands
| | - Sanne Broekman
- Department of Otorhinolaryngology, Radboud University Medical Center, 6525 GA Nijmegen, The Netherlands; (S.B.); (V.v.d.R.); (H.K.); (E.v.W.)
- Donders Institute for Brain Cognition and Behaviour, 6500 GL Nijmegen, The Netherlands; (S.E.d.B.); (J.R.); (M.A.)
| | - Vince van de Riet
- Department of Otorhinolaryngology, Radboud University Medical Center, 6525 GA Nijmegen, The Netherlands; (S.B.); (V.v.d.R.); (H.K.); (E.v.W.)
- Donders Institute for Brain Cognition and Behaviour, 6500 GL Nijmegen, The Netherlands; (S.E.d.B.); (J.R.); (M.A.)
| | - Marco Aben
- Donders Institute for Brain Cognition and Behaviour, 6500 GL Nijmegen, The Netherlands; (S.E.d.B.); (J.R.); (M.A.)
- Department of Human Genetics, Radboud University Medical Center, 6525 GA Nijmegen, The Netherlands
| | - Hannie Kremer
- Department of Otorhinolaryngology, Radboud University Medical Center, 6525 GA Nijmegen, The Netherlands; (S.B.); (V.v.d.R.); (H.K.); (E.v.W.)
- Donders Institute for Brain Cognition and Behaviour, 6500 GL Nijmegen, The Netherlands; (S.E.d.B.); (J.R.); (M.A.)
- Department of Human Genetics, Radboud University Medical Center, 6525 GA Nijmegen, The Netherlands
| | - Erwin van Wijk
- Department of Otorhinolaryngology, Radboud University Medical Center, 6525 GA Nijmegen, The Netherlands; (S.B.); (V.v.d.R.); (H.K.); (E.v.W.)
- Donders Institute for Brain Cognition and Behaviour, 6500 GL Nijmegen, The Netherlands; (S.E.d.B.); (J.R.); (M.A.)
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Liu Y, Liang F, Dong Z, Li S, Ye J, Qin W. Genome Editing in Zebrafish by ScCas9 Recognizing NNG PAM. Cells 2021; 10:2099. [PMID: 34440868 PMCID: PMC8392876 DOI: 10.3390/cells10082099] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/14/2021] [Revised: 08/11/2021] [Accepted: 08/13/2021] [Indexed: 11/16/2022] Open
Abstract
The CRISPR/Cas9 system has been widely used for gene editing in zebrafish. However, the required NGG protospacer adjacent motif (PAM) of Streptococcus pyogenes Cas9 (SpCas9) notably restricts the editable range of the zebrafish genome. Recently, Cas9 from S. canis (ScCas9), which has a more relaxed 5'-NNG-3' PAM, was reported to have activities in human cells and plants. However, the editing ability of ScCas9 has not been tested in zebrafish. Here we characterized and optimized the activity of ScCas9 in zebrafish. Delivered as a ribonucleoprotein complex, ScCas9 can induce mutations in zebrafish. Using the synthetic modified crRNA:tracrRNA duplex instead of in vitro-transcribed single guide RNA, the low activity at some loci were dramatically improved in zebrafish. As far as we know, our work is the first report on the evaluation of ScCas9 in animals. Our work optimized ScCas9 as a new nuclease for targeting relaxed NNG PAMs for zebrafish genome editing, which will further improve genome editing in zebrafish.
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Affiliation(s)
- Yunxing Liu
- State Key Laboratory of Chemical Oncogenomics, Key Laboratory of Chemical Genomics, Peking University Shenzhen Graduate School, Shenzhen 518055, China;
| | - Fang Liang
- Guangdong Provincial Key Laboratory for Healthy and Safe Aquaculture, Guangdong Provincial Engineering Technology Research Center for Environmentally-Friendly Aquaculture, Institute of Modern Aquaculture Science and Engineering, School of Life Sciences, South China Normal University, Guangzhou 510631, China; (F.L.); (Z.D.)
| | - Zijiong Dong
- Guangdong Provincial Key Laboratory for Healthy and Safe Aquaculture, Guangdong Provincial Engineering Technology Research Center for Environmentally-Friendly Aquaculture, Institute of Modern Aquaculture Science and Engineering, School of Life Sciences, South China Normal University, Guangzhou 510631, China; (F.L.); (Z.D.)
| | - Song Li
- State Key Laboratory of Chemical Oncogenomics, Key Laboratory of Chemical Genomics, Peking University Shenzhen Graduate School, Shenzhen 518055, China;
| | - Jianmin Ye
- Guangdong Provincial Key Laboratory for Healthy and Safe Aquaculture, Guangdong Provincial Engineering Technology Research Center for Environmentally-Friendly Aquaculture, Institute of Modern Aquaculture Science and Engineering, School of Life Sciences, South China Normal University, Guangzhou 510631, China; (F.L.); (Z.D.)
| | - Wei Qin
- Key Laboratory of Brain, Cognition and Education Sciences, Ministry of Education, Institute for Brain Research and Rehabilitation, South China Normal University, Guangzhou 510631, China
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30
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Formosa MM, Bergen DJM, Gregson CL, Maurizi A, Kämpe A, Garcia-Giralt N, Zhou W, Grinberg D, Ovejero Crespo D, Zillikens MC, Williams GR, Bassett JHD, Brandi ML, Sangiorgi L, Balcells S, Högler W, Van Hul W, Mäkitie O. A Roadmap to Gene Discoveries and Novel Therapies in Monogenic Low and High Bone Mass Disorders. Front Endocrinol (Lausanne) 2021; 12:709711. [PMID: 34539568 PMCID: PMC8444146 DOI: 10.3389/fendo.2021.709711] [Citation(s) in RCA: 7] [Impact Index Per Article: 2.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Submit a Manuscript] [Subscribe] [Scholar Register] [Received: 05/14/2021] [Accepted: 07/12/2021] [Indexed: 12/24/2022] Open
Abstract
Genetic disorders of the skeleton encompass a diverse group of bone diseases differing in clinical characteristics, severity, incidence and molecular etiology. Of particular interest are the monogenic rare bone mass disorders, with the underlying genetic defect contributing to either low or high bone mass phenotype. Extensive, deep phenotyping coupled with high-throughput, cost-effective genotyping is crucial in the characterization and diagnosis of affected individuals. Massive parallel sequencing efforts have been instrumental in the discovery of novel causal genes that merit functional validation using in vitro and ex vivo cell-based techniques, and in vivo models, mainly mice and zebrafish. These translational models also serve as an excellent platform for therapeutic discovery, bridging the gap between basic science research and the clinic. Altogether, genetic studies of monogenic rare bone mass disorders have broadened our knowledge on molecular signaling pathways coordinating bone development and metabolism, disease inheritance patterns, development of new and improved bone biomarkers, and identification of novel drug targets. In this comprehensive review we describe approaches to further enhance the innovative processes taking discoveries from clinic to bench, and then back to clinic in rare bone mass disorders. We highlight the importance of cross laboratory collaboration to perform functional validation in multiple model systems after identification of a novel disease gene. We describe the monogenic forms of rare low and high rare bone mass disorders known to date, provide a roadmap to unravel the genetic determinants of monogenic rare bone mass disorders using proper phenotyping and genotyping methods, and describe different genetic validation approaches paving the way for future treatments.
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Affiliation(s)
- Melissa M. Formosa
- Department of Applied Biomedical Science, Faculty of Health Sciences, University of Malta, Msida, Malta
- Centre for Molecular Medicine and Biobanking, University of Malta, Msida, Malta
| | - Dylan J. M. Bergen
- School of Physiology, Pharmacology, and Neuroscience, Faculty of Life Sciences, University of Bristol, Bristol, United Kingdom
- The Musculoskeletal Research Unit, Translational Health Sciences, Bristol Medical School, Faculty of Health Sciences, University of Bristol, Bristol, United Kingdom
| | - Celia L. Gregson
- The Musculoskeletal Research Unit, Translational Health Sciences, Bristol Medical School, Faculty of Health Sciences, University of Bristol, Bristol, United Kingdom
| | - Antonio Maurizi
- Department of Applied Clinical Sciences and Biotechnological, University of L’Aquila, L’Aquila, Italy
| | - Anders Kämpe
- Department of Molecular Medicine and Surgery, Karolinska Institutet, Stockholm, Sweden
- Department of Clinical Genetics, Karolinska University Hospital, Stockholm, Sweden
| | - Natalia Garcia-Giralt
- IMIM (Hospital del Mar Research Institute), Centro de Investigación Biomédica en Red de Fragilidad y Envejecimiento Saludable (CIBERFES), Barcelona, Spain
| | - Wei Zhou
- Department of Internal Medicine, Erasmus University Medical Center, Rotterdam, Netherlands
| | - Daniel Grinberg
- Department of Genetics, Microbiology and Statistics, Faculty of Biology, Universitat de Barcelona, Centro de Investigación Biomédica en Red de Enfermedades Raras (CIBERER), Institut de Biomedicina de la Universitat de Barcelona (IBUB), Institut de Recerca Sant Joan de Déu (IRSJD), Barcelona, Spain
| | - Diana Ovejero Crespo
- IMIM (Hospital del Mar Research Institute), Centro de Investigación Biomédica en Red de Fragilidad y Envejecimiento Saludable (CIBERFES), Barcelona, Spain
| | - M. Carola Zillikens
- Department of Internal Medicine, Erasmus University Medical Center, Rotterdam, Netherlands
| | - Graham R. Williams
- Molecular Endocrinology Laboratory, Department of Metabolism, Digestion and Reproduction, Imperial College London, London, United Kingdom
| | - J. H. Duncan Bassett
- Molecular Endocrinology Laboratory, Department of Metabolism, Digestion and Reproduction, Imperial College London, London, United Kingdom
| | - Maria Luisa Brandi
- Department of Surgery and Translational Medicine (M.L.B.), University of Florence, Florence, Italy
| | - Luca Sangiorgi
- Department of Medical Genetics and Skeletal Rare Diseases, IRCCS Rizzoli Orthopaedic Institute, Bologna, Italy
| | - Susanna Balcells
- Department of Genetics, Microbiology and Statistics, Faculty of Biology, Universitat de Barcelona, Centro de Investigación Biomédica en Red de Enfermedades Raras (CIBERER), Institut de Biomedicina de la Universitat de Barcelona (IBUB), Institut de Recerca Sant Joan de Déu (IRSJD), Barcelona, Spain
| | - Wolfgang Högler
- Department of Paediatrics and Adolescent Medicine, Johannes Kepler University Linz, Linz, Austria
- Institute of Metabolism and Systems Research, University of Birmingham, Birmingham, United Kingdom
| | - Wim Van Hul
- Department of Medical Genetics, University of Antwerp, Antwerp, Belgium
| | - Outi Mäkitie
- Children’s Hospital, University of Helsinki and Helsinki University Hospital, Helsinki, Finland
- Research Program for Clinical and Molecular Metabolism, Faculty of Medicine, University of Helsinki, Helsinki, Finland
- Folkhälsan Research Centre, Folkhälsan Institute of Genetics, Helsinki, Finland
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Straume AH, Kjærner-Semb E, Skaftnesmo KO, Güralp H, Lillico S, Wargelius A, Edvardsen RB. Single nucleotide replacement in the Atlantic salmon genome using CRISPR/Cas9 and asymmetrical oligonucleotide donors. BMC Genomics 2021; 22:563. [PMID: 34294050 PMCID: PMC8296724 DOI: 10.1186/s12864-021-07823-8] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/23/2021] [Accepted: 06/17/2021] [Indexed: 12/12/2022] Open
Abstract
Background New breeding technologies (NBT) using CRISPR/Cas9-induced homology directed repair (HDR) has the potential to expedite genetic improvement in aquaculture. The long generation time in Atlantic salmon makes breeding an unattractive solution to obtain homozygous mutants and improving the rates of perfect HDR in founder (F0) fish is thus required. Genome editing can represent small DNA changes down to single nucleotide replacements (SNR). This enables edits such as premature stop codons or single amino acid changes and may be used to obtain fish with traits favorable to aquaculture, e.g. disease resistance. A method for SNR has not yet been demonstrated in salmon. Results Using CRISPR/Cas9 and asymmetrical ODNs, we were able to perform precise SNR and introduce a premature stop codon in dnd in F0 salmon. Deep sequencing demonstrated up to 59.2% efficiency in single embryos. In addition, using the same asymmetrical ODN design, we inserted a FLAG element into slc45a2 and dnd, showing high individual perfect HDR efficiencies (up to 36.7 and 32.7%, respectively). Conclusions In this work, we demonstrate that precise SNR and knock-in (KI) can be performed in F0 salmon embryos using asymmetrical oligonucleotide (ODN) donors. We suggest that HDR-induced SNR can be applied as a powerful NBT, allowing efficient introgression of favorable alleles and bypassing challenges associated with traditional selective breeding. Supplementary Information The online version contains supplementary material available at 10.1186/s12864-021-07823-8.
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Affiliation(s)
- Anne Hege Straume
- Institute of Marine Research, P.O. Box 1870, Nordnes, NO-5817, Bergen, Norway
| | - Erik Kjærner-Semb
- Institute of Marine Research, P.O. Box 1870, Nordnes, NO-5817, Bergen, Norway
| | - Kai Ove Skaftnesmo
- Institute of Marine Research, P.O. Box 1870, Nordnes, NO-5817, Bergen, Norway
| | - Hilal Güralp
- Institute of Marine Research, P.O. Box 1870, Nordnes, NO-5817, Bergen, Norway
| | - Simon Lillico
- The Roslin Institute and R(D)SVS, University of Edinburgh, Easter Bush Campus, Midlothian, EH25 9RG, UK
| | - Anna Wargelius
- Institute of Marine Research, P.O. Box 1870, Nordnes, NO-5817, Bergen, Norway
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Sizova I, Kelterborn S, Verbenko V, Kateriya S, Hegemann P. Chlamydomonas POLQ is necessary for CRISPR/Cas9-mediated gene targeting. G3 (BETHESDA, MD.) 2021; 11:jkab114. [PMID: 33836052 PMCID: PMC8495919 DOI: 10.1093/g3journal/jkab114] [Citation(s) in RCA: 9] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Subscribe] [Scholar Register] [Received: 02/09/2021] [Accepted: 03/18/2021] [Indexed: 02/06/2023]
Abstract
The use of CRISPR/Cas endonucleases has revolutionized gene editing techniques for research on Chlamydomonas reinhardtii. To better utilize the CRISPR/Cas system, it is essential to develop a more comprehensive understanding of the DNA repair pathways involved in genome editing. In this study, we have analyzed contributions from canonical KU80/KU70-dependent nonhomologous end-joining (cNHEJ) and DNA polymerase theta (POLQ)-mediated end joining on SpCas9-mediated untemplated mutagenesis and homology-directed repair (HDR)/gene inactivation in Chlamydomonas. Using CRISPR/SpCas9 technology, we generated DNA repair-defective mutants ku80, ku70, polQ for gene targeting experiments. Our results show that untemplated repair of SpCas9-induced double strand breaks results in mutation spectra consistent with an involvement of both KU80/KU70 and POLQ. In addition, the inactivation of POLQ was found to negatively affect HDR of the inactivated paromomycin-resistant mut-aphVIII gene when donor single-stranded oligos were used. Nevertheless, mut-aphVIII was still repaired by homologous recombination in these mutants. POLQ inactivation suppressed random integration of transgenes co-transformed with the donor ssDNA. KU80 deficiency did not affect these events but instead was surprisingly found to stimulate HDR/gene inactivation. Our data suggest that in Chlamydomonas, POLQ is the main contributor to CRISPR/Cas-induced HDR and random integration of transgenes, whereas KU80/KU70 potentially plays a secondary role. We expect our results will lead to improvement of genome editing in C. reinhardtii and can be used for future development of algal biotechnology.
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Affiliation(s)
- Irina Sizova
- Experimental Biophysics, Institute of Biology, Humboldt University of Berlin, Berlin D-10099, Germany
- Petersburg Nuclear Physics Institute named by B.P. Konstantinov of National Research Centre “Kurchatov Institute”, Gatchina 188300, Russia
- Kurchatov Genome Center - PNPI, Gatchina 188300, Russia
| | - Simon Kelterborn
- Experimental Biophysics, Institute of Biology, Humboldt University of Berlin, Berlin D-10099, Germany
| | - Valeriy Verbenko
- Petersburg Nuclear Physics Institute named by B.P. Konstantinov of National Research Centre “Kurchatov Institute”, Gatchina 188300, Russia
- Kurchatov Genome Center - PNPI, Gatchina 188300, Russia
| | - Suneel Kateriya
- Laboratory of Optobiology School of Biotechnology, Jawaharlal Nehru University, New Delhi 110067, India
| | - Peter Hegemann
- Experimental Biophysics, Institute of Biology, Humboldt University of Berlin, Berlin D-10099, Germany
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Jayarajan V, Kounatidou E, Qasim W, Di W. Ex vivo gene modification therapy for genetic skin diseases-recent advances in gene modification technologies and delivery. Exp Dermatol 2021; 30:887-896. [PMID: 33657662 PMCID: PMC8432139 DOI: 10.1111/exd.14314] [Citation(s) in RCA: 11] [Impact Index Per Article: 3.7] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/22/2020] [Revised: 02/19/2021] [Accepted: 02/25/2021] [Indexed: 12/19/2022]
Abstract
Genetic skin diseases, also known as genodermatoses, are inherited disorders affecting skin and constitute a large and heterogeneous group of diseases. While genodermatoses are rare with the prevalence rate of less than 1 in 50,000 - 200,000, they frequently occur at birth or early in life and are generally chronic, severe, and could be life-threatening. The quality of life of patients and their families are severely compromised by the negative psychosocial impact of disease, physical manifestations, and the lack or loss of autonomy. Currently, there are no curative treatments for these conditions. Ex vivo gene modification therapy that involves modification or correction of mutant genes in patients' cells in vitro and then transplanted back to patients to restore functional gene expression has being developed for genodermatoses. In this review, the ex vivo gene modification therapy strategies for genodermatoses are reviewed, focusing on current advances in gene modification and correction in patients' cells and delivery of genetically modified cells to patients with discussions on gene therapy trials which have been performed in this area.
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Affiliation(s)
- Vignesh Jayarajan
- Infection, Immunity and Inflammation Research & Teaching Department, Immunobiology SectionUCL Great Ormond Street Institute of Child HealthLondonUK
| | - Evangelia Kounatidou
- Infection, Immunity and Inflammation Research & Teaching Department, Immunobiology SectionUCL Great Ormond Street Institute of Child HealthLondonUK
| | - Waseem Qasim
- Infection, Immunity and Inflammation Research & Teaching Department, Molecular and Cellular Immunology SectionUCL Great Ormond Street Institute of Child HealthLondonUK
| | - Wei‐Li Di
- Infection, Immunity and Inflammation Research & Teaching Department, Immunobiology SectionUCL Great Ormond Street Institute of Child HealthLondonUK
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Zhang M, Yang C, Tasan I, Zhao H. Expanding the Potential of Mammalian Genome Engineering via Targeted DNA Integration. ACS Synth Biol 2021; 10:429-446. [PMID: 33596056 DOI: 10.1021/acssynbio.0c00576] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/08/2023]
Abstract
Inserting custom designed DNA sequences into the mammalian genome plays an essential role in synthetic biology. In particular, the ability to introduce foreign DNA in a site-specific manner offers numerous advantages over random DNA integration. In this review, we focus on two mechanistically distinct systems that have been widely adopted for targeted DNA insertion in mammalian cells, the CRISPR/Cas9 system and site-specific recombinases. The CRISPR/Cas9 system has revolutionized the genome engineering field thanks to its high programmability and ease of use. However, due to its dependence on linearized DNA donor and endogenous cellular pathways to repair the induced double-strand break, CRISPR/Cas9-mediated DNA insertion still faces limitations such as small insert size, and undesired editing outcomes via error-prone repair pathways. In contrast, site-specific recombinases, in particular the Serine integrases, demonstrate large-cargo capability and no dependence on cellular repair pathways for DNA integration. Here we first describe recent advances in improving the overall efficacy of CRISPR/Cas9-based methods for DNA insertion. Moreover, we highlight the advantages of site-specific recombinases over CRISPR/Cas9 in the context of targeted DNA integration, with a special focus on the recent development of programmable recombinases. We conclude by discussing the importance of protein engineering to further expand the current toolkit for targeted DNA insertion in mammalian cells.
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Affiliation(s)
- Meng Zhang
- Department of Chemical and Biomolecular Engineering, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801, United States
- Carl R. Woese Institute for Genomic Biology, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801, United States
| | - Che Yang
- Carl R. Woese Institute for Genomic Biology, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801, United States
- Department of Biochemistry, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801, United States
| | - Ipek Tasan
- Carl R. Woese Institute for Genomic Biology, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801, United States
- Department of Biochemistry, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801, United States
| | - Huimin Zhao
- Department of Chemical and Biomolecular Engineering, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801, United States
- Carl R. Woese Institute for Genomic Biology, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801, United States
- Department of Biochemistry, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801, United States
- Department of Chemistry, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801, United States
- Department of Bioengineering, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801, United States
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Ranawakage DC, Okada K, Sugio K, Kawaguchi Y, Kuninobu-Bonkohara Y, Takada T, Kamachi Y. Efficient CRISPR-Cas9-Mediated Knock-In of Composite Tags in Zebrafish Using Long ssDNA as a Donor. Front Cell Dev Biol 2021; 8:598634. [PMID: 33681181 PMCID: PMC7928300 DOI: 10.3389/fcell.2020.598634] [Citation(s) in RCA: 13] [Impact Index Per Article: 4.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/25/2020] [Accepted: 12/31/2020] [Indexed: 12/12/2022] Open
Abstract
Despite the unprecedented gene editing capability of CRISPR-Cas9-mediated targeted knock-in, the efficiency and precision of this technology still require further optimization, particularly for multicellular model organisms, such as the zebrafish (Danio rerio). Our study demonstrated that an ∼200 base-pair sequence encoding a composite tag can be efficiently "knocked-in" into the zebrafish genome using a combination of the CRISPR-Cas9 ribonucleoprotein complex and a long single-stranded DNA (lssDNA) as a donor template. Here, we targeted the sox3, sox11a, and pax6a genes to evaluate the knock-in efficiency of lssDNA donors with different structures in somatic cells of injected embryos and for their germline transmission. The structures and sequence characteristics of the lssDNA donor templates were found to be crucial to achieve a high rate of precise and heritable knock-ins. The following were our key findings: (1) lssDNA donor strand selection is important; however, strand preference and its dependency appear to vary among the target loci or their sequences. (2) The length of the 3' homology arm of the lssDNA donor affects knock-in efficiency in a site-specific manner; particularly, a shorter 50-nt arm length leads to a higher knock-in efficiency than a longer 300-nt arm for the sox3 and pax6a knock-ins. (3) Some DNA sequence characteristics of the knock-in donors and the distance between the CRISPR-Cas9 cleavage site and the tag insertion site appear to adversely affect the repair process, resulting in imprecise editing. By implementing the proposed method, we successfully obtained precisely edited sox3, sox11a, and pax6a knock-in alleles that contained a composite tag composed of FLAGx3 (or PAx3), Bio tag, and HiBiT tag (or His tag) with moderate to high germline transmission rates as high as 21%. Furthermore, the knock-in allele-specific quantitative polymerase chain reaction (qPCR) for both the 5' and 3' junctions indicated that knock-in allele frequencies were higher at the 3' side of the lssDNAs, suggesting that the lssDNA-templated knock-in was mediated by unidirectional single-strand template repair (SSTR) in zebrafish embryos.
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Affiliation(s)
| | | | | | | | | | | | - Yusuke Kamachi
- School of Environmental Science and Engineering, Kochi University of Technology, Kochi, Japan
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DGK and DZHK position paper on genome editing: basic science applications and future perspective. Basic Res Cardiol 2021; 116:2. [PMID: 33449167 PMCID: PMC7810637 DOI: 10.1007/s00395-020-00839-3] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Submit a Manuscript] [Subscribe] [Scholar Register] [Received: 09/17/2020] [Accepted: 12/09/2020] [Indexed: 12/18/2022]
Abstract
For a long time, gene editing had been a scientific concept, which was limited to a few applications. With recent developments, following the discovery of TALEN zinc-finger endonucleases and in particular the CRISPR/Cas system, gene editing has become a technique applicable in most laboratories. The current gain- and loss-of function models in basic science are revolutionary as they allow unbiased screens of unprecedented depth and complexity and rapid development of transgenic animals. Modifications of CRISPR/Cas have been developed to precisely interrogate epigenetic regulation or to visualize DNA complexes. Moreover, gene editing as a clinical treatment option is rapidly developing with first trials on the way. This article reviews the most recent progress in the field, covering expert opinions gathered during joint conferences on genome editing of the German Cardiac Society (DGK) and the German Center for Cardiovascular Research (DZHK). Particularly focusing on the translational aspect and the combination of cellular and animal applications, the authors aim to provide direction for the development of the field and the most frequent applications with their problems.
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Prill K, Dawson JF. Homology-Directed Repair in Zebrafish: Witchcraft and Wizardry? Front Mol Biosci 2021; 7:595474. [PMID: 33425990 PMCID: PMC7793982 DOI: 10.3389/fmolb.2020.595474] [Citation(s) in RCA: 7] [Impact Index Per Article: 2.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/16/2020] [Accepted: 10/20/2020] [Indexed: 12/12/2022] Open
Abstract
Introducing desired mutations into the genome of model organisms is a priority for all research focusing on protein function and disease modeling. The need to create stable mutant lines has resulted in the rapid advancement of genetic techniques over the last few decades from chemical mutagenesis and zinc finger nucleases to clustered regularly interspaced short palindromic repeats (CRISPR) and homology-directed repair (HDR). However, achieving consistently high success rates for direct mutagenesis in zebrafish remains one of the most sought-after techniques in the field. Several genes have been modified using HDR in zebrafish, but published success rates range widely, suggesting that an optimal protocol is required. In this review, we compare target genes, techniques, and protocols from 50 genes that were successfully modified in zebrafish using HDR to find the statistically best variables for efficient HDR rates.
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Affiliation(s)
- Kendal Prill
- Department of Molecular and Cellular Biology, University of Guelph, Guelph, ON, Canada
| | - John F Dawson
- Department of Molecular and Cellular Biology, University of Guelph, Guelph, ON, Canada
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Sieliwonczyk E, Matchkov VV, Vandendriessche B, Alaerts M, Bakkers J, Loeys B, Schepers D. Inherited Ventricular Arrhythmia in Zebrafish: Genetic Models and Phenotyping Tools. Rev Physiol Biochem Pharmacol 2021; 184:33-68. [PMID: 34533615 DOI: 10.1007/112_2021_65] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/18/2023]
Abstract
In the last years, the field of inheritable ventricular arrhythmia disease modelling has changed significantly with a push towards the use of novel cellular cardiomyocyte based models. However, there is a growing need for new in vivo models to study the disease pathology at the tissue and organ level. Zebrafish provide an excellent opportunity for in vivo modelling of inheritable ventricular arrhythmia syndromes due to the remarkable similarity between their cardiac electrophysiology and that of humans. Additionally, many state-of-the-art methods in gene editing and electrophysiological phenotyping are available for zebrafish research. In this review, we give a comprehensive overview of the published zebrafish genetic models for primary electrical disorders and arrhythmogenic cardiomyopathy. We summarise and discuss the strengths and weaknesses of the different technical approaches for the generation of genetically modified zebrafish disease models, as well as the electrophysiological approaches in zebrafish phenotyping. By providing this detailed overview, we aim to draw attention to the potential of the zebrafish model for studying arrhythmia syndromes at the organ level and as a platform for personalised medicine and drug testing.
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Affiliation(s)
- Ewa Sieliwonczyk
- Center of Medical Genetics, Faculty of Medicine and Health Sciences, University of Antwerp and Antwerp University Hospital, Antwerp, Belgium.
| | - Vladimir V Matchkov
- Department of Biomedicine, Pulmonary and Cardiovascular Pharmacology, Aarhus University, Aarhus, Denmark
| | - Bert Vandendriessche
- Center of Medical Genetics, Faculty of Medicine and Health Sciences, University of Antwerp and Antwerp University Hospital, Antwerp, Belgium
| | - Maaike Alaerts
- Center of Medical Genetics, Faculty of Medicine and Health Sciences, University of Antwerp and Antwerp University Hospital, Antwerp, Belgium
| | - Jeroen Bakkers
- Hubrecht Institute for Developmental and Stem Cell Biology, Utrecht, The Netherlands
| | - Bart Loeys
- Center of Medical Genetics, Faculty of Medicine and Health Sciences, University of Antwerp and Antwerp University Hospital, Antwerp, Belgium
| | - Dorien Schepers
- Center of Medical Genetics, Faculty of Medicine and Health Sciences, University of Antwerp and Antwerp University Hospital, Antwerp, Belgium.,Laboratory for Molecular, Cellular and Network Excitability, Department of Biomedical Sciences, University of Antwerp, Antwerp, Belgium
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Potekhina ES, Bass DY, Kelmanson IV, Fetisova ES, Ivanenko AV, Belousov VV, Bilan DS. Drug Screening with Genetically Encoded Fluorescent Sensors: Today and Tomorrow. Int J Mol Sci 2020; 22:E148. [PMID: 33375682 PMCID: PMC7794770 DOI: 10.3390/ijms22010148] [Citation(s) in RCA: 5] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/01/2020] [Revised: 12/18/2020] [Accepted: 12/24/2020] [Indexed: 02/07/2023] Open
Abstract
Genetically-encoded fluorescent sensors have been actively developed over the last few decades and used in live imaging and drug screening. Real-time monitoring of drug action in a specific cellular compartment, organ, or tissue type; the ability to screen at the single-cell resolution; and the elimination of false-positive results caused by low drug bioavailability that is not detected by in vitro testing methods are a few of the obvious benefits of using genetically-encoded fluorescent sensors in drug screening. In combination with high-throughput screening (HTS), some genetically-encoded fluorescent sensors may provide high reproducibility and robustness to assays. We provide a brief overview of successful, perspective, and hopeful attempts at using genetically encoded fluorescent sensors in HTS of modulators of ion channels, Ca2+ homeostasis, GPCR activity, and for screening cytotoxic, anticancer, and anti-parasitic compounds. We discuss the advantages of sensors in whole organism drug screening models and the perspectives of the combination of human disease modeling by CRISPR techniques with genetically encoded fluorescent sensors for drug screening.
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Affiliation(s)
- Ekaterina S. Potekhina
- Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, 117997 Moscow, Russia; (D.Y.B.); (I.V.K.); (E.S.F.); (A.V.I.); (V.V.B.)
- Center for Precision Genome Editing and Genetic Technologies for Biomedicine, Pirogov Russian National Research Medical University, 117997 Moscow, Russia
| | - Dina Y. Bass
- Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, 117997 Moscow, Russia; (D.Y.B.); (I.V.K.); (E.S.F.); (A.V.I.); (V.V.B.)
- Center for Precision Genome Editing and Genetic Technologies for Biomedicine, Pirogov Russian National Research Medical University, 117997 Moscow, Russia
| | - Ilya V. Kelmanson
- Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, 117997 Moscow, Russia; (D.Y.B.); (I.V.K.); (E.S.F.); (A.V.I.); (V.V.B.)
- Center for Precision Genome Editing and Genetic Technologies for Biomedicine, Pirogov Russian National Research Medical University, 117997 Moscow, Russia
| | - Elena S. Fetisova
- Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, 117997 Moscow, Russia; (D.Y.B.); (I.V.K.); (E.S.F.); (A.V.I.); (V.V.B.)
| | - Alexander V. Ivanenko
- Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, 117997 Moscow, Russia; (D.Y.B.); (I.V.K.); (E.S.F.); (A.V.I.); (V.V.B.)
- Center for Precision Genome Editing and Genetic Technologies for Biomedicine, Pirogov Russian National Research Medical University, 117997 Moscow, Russia
| | - Vsevolod V. Belousov
- Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, 117997 Moscow, Russia; (D.Y.B.); (I.V.K.); (E.S.F.); (A.V.I.); (V.V.B.)
- Center for Precision Genome Editing and Genetic Technologies for Biomedicine, Pirogov Russian National Research Medical University, 117997 Moscow, Russia
- Federal Center of Brain Research and Neurotechnologies of the Federal Medical Biological Agency, 117997 Moscow, Russia
| | - Dmitry S. Bilan
- Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, 117997 Moscow, Russia; (D.Y.B.); (I.V.K.); (E.S.F.); (A.V.I.); (V.V.B.)
- Center for Precision Genome Editing and Genetic Technologies for Biomedicine, Pirogov Russian National Research Medical University, 117997 Moscow, Russia
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Hewes AM, Sansbury BM, Kmiec EB. The Diversity of Genetic Outcomes from CRISPR/Cas Gene Editing is Regulated by the Length of the Symmetrical Donor DNA Template. Genes (Basel) 2020; 11:genes11101160. [PMID: 33008045 PMCID: PMC7599521 DOI: 10.3390/genes11101160] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/25/2020] [Revised: 09/22/2020] [Accepted: 09/28/2020] [Indexed: 12/27/2022] Open
Abstract
Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas gene editing systems have enabled molecular geneticists to manipulate prokaryotic and eukaryotic genomes with greater efficiency and precision. CRISPR/Cas provides adaptive immunity in bacterial cells by degrading invading viral genomes. By democratizing this activity into human cells, it is possible to knock out specific genes to disable their function and repair errors. The latter of these activities requires the participation of a single-stranded donor DNA template that provides the genetic information to execute correction in a process referred to as homology directed repair (HDR). Here, we utilized an established cell-free extract system to determine the influence that the donor DNA template length has on the diversity of products from CRISPR-directed gene editing. This model system enables us to view all outcomes of this reaction and reveals that donor template length can influence the efficiency of the reaction and the categories of error-prone products that accompany it. A careful measurement of the products revealed a category of error-prone events that contained the corrected template along with insertions and deletions (indels). Our data provides foundational information for those whose aim is to translate CRISPR/Cas from bench to bedside.
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Affiliation(s)
- Amanda M. Hewes
- Gene Editing Institute, Helen F. Graham Cancer Center & Research Institute, Christiana Care Health System, Newark, DE 19713, USA; (A.M.H.); (B.M.S.)
| | - Brett M. Sansbury
- Gene Editing Institute, Helen F. Graham Cancer Center & Research Institute, Christiana Care Health System, Newark, DE 19713, USA; (A.M.H.); (B.M.S.)
- Department of Medical and Molecular Sciences, University of Delaware, Newark, DE 19716, USA
| | - Eric B. Kmiec
- Gene Editing Institute, Helen F. Graham Cancer Center & Research Institute, Christiana Care Health System, Newark, DE 19713, USA; (A.M.H.); (B.M.S.)
- Department of Medical and Molecular Sciences, University of Delaware, Newark, DE 19716, USA
- Correspondence: ; Tel.: +1-(0)302-623-0628
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41
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Modulation of DNA double-strand break repair as a strategy to improve precise genome editing. Oncogene 2020; 39:6393-6405. [PMID: 32884115 DOI: 10.1038/s41388-020-01445-2] [Citation(s) in RCA: 24] [Impact Index Per Article: 6.0] [Reference Citation Analysis] [Abstract] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/06/2020] [Revised: 08/07/2020] [Accepted: 08/21/2020] [Indexed: 12/11/2022]
Abstract
In the present day, it is possible to incorporate targeted mutations or replace a gene using genome editing techniques such as customisable CRISPR/Cas9 system. Although induction of DNA double-strand breaks (DSBs) by genome editing tools can be repaired by both non-homologous end joining (NHEJ) and homologous recombination (HR), the skewness of the former pathway in human and other mammals normally result in imprecise repair. Scientists working at the crossroads of DNA repair and genome editing have devised new strategies for using a specific pathway to their advantage. Refinement in the efficiency of precise gene editing was witnessed upon downregulation of NHEJ by knockdown or using small molecule inhibitors on one hand, and upregulation of HR proteins and addition of HR stimulators, other hand. The exploitation of cell cycle phase differences together with appropriate donor DNA length/sequence and small molecules has provided further improvement in precise genome editing. The present article reviews the mechanisms of improving the efficiency of precise genome editing in several model organisms and in clinics.
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Carrington B, Weinstein RN, Sood R. BE4max and AncBE4max Are Efficient in Germline Conversion of C:G to T:A Base Pairs in Zebrafish. Cells 2020; 9:cells9071690. [PMID: 32674364 PMCID: PMC7407168 DOI: 10.3390/cells9071690] [Citation(s) in RCA: 13] [Impact Index Per Article: 3.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/15/2020] [Revised: 07/06/2020] [Accepted: 07/10/2020] [Indexed: 12/12/2022] Open
Abstract
The ease of use and robustness of genome editing by CRISPR/Cas9 has led to successful use of gene knockout zebrafish for disease modeling. However, it still remains a challenge to precisely edit the zebrafish genome to create single-nucleotide substitutions, which account for ~60% of human disease-causing mutations. Recently developed base editing nucleases provide an excellent alternate to CRISPR/Cas9-mediated homology dependent repair for generation of zebrafish with point mutations. A new set of cytosine base editors, termed BE4max and AncBE4max, demonstrated improved base editing efficiency in mammalian cells but have not been evaluated in zebrafish. Therefore, we undertook this study to evaluate their efficiency in converting C:G to T:A base pairs in zebrafish by somatic and germline analysis using highly active sgRNAs to twist and ntl genes. Our data demonstrated that these improved BE4max set of plasmids provide desired base substitutions at similar efficiency and without any indels compared to the previously reported BE3 and Target-AID plasmids in zebrafish. Our data also showed that AncBE4max produces fewer incorrect and bystander edits, suggesting that it can be further improved by codon optimization of its components for use in zebrafish.
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Hewes AM, Sansbury BM, Barth S, Tarcic G, Kmiec EB. gRNA Sequence Heterology Tolerance Catalyzed by CRISPR/Cas in an In Vitro Homology-Directed Repair Reaction. MOLECULAR THERAPY. NUCLEIC ACIDS 2020; 20:568-579. [PMID: 32330873 PMCID: PMC7177190 DOI: 10.1016/j.omtn.2020.03.012] [Citation(s) in RCA: 7] [Impact Index Per Article: 1.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Subscribe] [Scholar Register] [Received: 02/27/2020] [Revised: 03/17/2020] [Accepted: 03/24/2020] [Indexed: 12/17/2022]
Abstract
CRISPR and associated Cas nucleases are genetic engineering tools revolutionizing innovative approaches to cancer and inherited diseases. CRISPR-directed gene editing relies heavily on proper DNA sequence alignment between the guide RNA (gRNA)/CRISPR complex and its genomic target. Accurate hybridization of complementary DNA initiates gene editing in human cells, but inherent gRNA sequence variation that could influence the gene editing reaction has been clearly established among diverse genetic populations. As this technology advances toward clinical implementation, it will be essential to assess what degree of gRNA variation generates unwanted and erroneous CRISPR activity. With the use of a system in which a cell-free extract catalyzes nonhomologous end joining (NHEJ) and homology-directed repair (HDR), it is possible to observe a more representative population of all forms of gene editing outcomes. In this manuscript, we demonstrate CRISPR/Cas complexation at heterologous binding sites that facilitate precise and error-prone HDR. The tolerance of mispairing between the gRNA and target site of the DNA to enable HDR is surprisingly high and greatly influenced by polarity of the donor DNA strand in the reaction. These results suggest that some collateral genomic activity could occur at unintended sites in CRISPR-directed gene editing in human cells.
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Affiliation(s)
- Amanda M Hewes
- Gene Editing Institute, Helen F. Graham Cancer Center & Research Institute, Christiana Care Health System, Newark, DE, USA
| | - Brett M Sansbury
- Gene Editing Institute, Helen F. Graham Cancer Center & Research Institute, Christiana Care Health System, Newark, DE, USA; Department of Medical and Molecular Sciences, University of Delaware, Newark, DE, USA
| | - Shaul Barth
- Novellus, Jerusalem Bio-Park, 1(st) Kiryat Hadassah, Hadassah Ein-Kerem Medical Center Campus, Jerusalem, Israel, 9112001
| | - Gabi Tarcic
- Novellus, Jerusalem Bio-Park, 1(st) Kiryat Hadassah, Hadassah Ein-Kerem Medical Center Campus, Jerusalem, Israel, 9112001
| | - Eric B Kmiec
- Gene Editing Institute, Helen F. Graham Cancer Center & Research Institute, Christiana Care Health System, Newark, DE, USA; Department of Medical and Molecular Sciences, University of Delaware, Newark, DE, USA.
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Erdogan M, Fabritius A, Basquin J, Griesbeck O. Targeted In Situ Protein Diversification and Intra-organelle Validation in Mammalian Cells. Cell Chem Biol 2020; 27:610-621.e5. [PMID: 32142629 DOI: 10.1016/j.chembiol.2020.02.004] [Citation(s) in RCA: 16] [Impact Index Per Article: 4.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/29/2019] [Revised: 12/22/2019] [Accepted: 02/14/2020] [Indexed: 02/08/2023]
Abstract
Engineered proteins must be phenotypically selected for function in the appropriate physiological context. Here, we present a versatile approach that allows generating panels of mammalian cells that express diversified heterologous protein libraries in the cytosol or subcellular compartments under stable conditions and in a single-variant-per-cell manner. To this end we adapt CRISPR/Cas9 editing technology to diversify targeted stretches of a protein of interest in situ. We demonstrate the utility of the approach by in situ engineering and intra-lysosome specific selection of an extremely pH-resistant long Stokes shift red fluorescent protein variant. Tailoring properties to specific conditions of cellular sub-compartments or organelles of mammalian cells can be an important asset to optimize various proteins, protein-based tools, and biosensors for distinct functions.
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Affiliation(s)
- Mutlu Erdogan
- Tools for Bio-Imaging, Max-Planck-Institut für Neurobiologie, Am Klopferspitz 18, Martinsried 82152, Germany
| | - Arne Fabritius
- Tools for Bio-Imaging, Max-Planck-Institut für Neurobiologie, Am Klopferspitz 18, Martinsried 82152, Germany
| | - Jérome Basquin
- Structural Cell Biology, Max-Planck-Institut für Biochemie, Am Klopferspitz 18, Martinsried 82152, Germany
| | - Oliver Griesbeck
- Tools for Bio-Imaging, Max-Planck-Institut für Neurobiologie, Am Klopferspitz 18, Martinsried 82152, Germany.
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Straume AH, Kjærner-Semb E, Ove Skaftnesmo K, Güralp H, Kleppe L, Wargelius A, Edvardsen RB. Indel locations are determined by template polarity in highly efficient in vivo CRISPR/Cas9-mediated HDR in Atlantic salmon. Sci Rep 2020; 10:409. [PMID: 31941961 PMCID: PMC6962318 DOI: 10.1038/s41598-019-57295-w] [Citation(s) in RCA: 14] [Impact Index Per Article: 3.5] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/22/2019] [Accepted: 12/19/2019] [Indexed: 01/20/2023] Open
Abstract
Precise gene editing such as CRISPR/Cas9-mediated homology directed repair (HDR) can increase our understanding of gene function and improve traits of importance for aquaculture. This fine-tuned technology has not been developed for farmed fish including Atlantic salmon. We performed knock-in (KI) of a FLAG element in the slc45a2 gene in salmon using sense (S), anti-sense (AS) and double-stranded (ds) oligodeoxynucleotide (ODN) templates with short (24/48/84 bp) homology arms. We show in vivo ODN integration in almost all the gene edited animals, and demonstrate perfect HDR rates up to 27% in individual F0 embryos, much higher than reported previously in any fish. HDR efficiency was dependent on template concentration, but not homology arm length. Analysis of imperfect HDR variants suggest that repair occurs by synthesis-dependent strand annealing (SDSA), as we show for the first time in any species that indel location is dependent on template polarity. Correct ODN polarity can be used to avoid 5'-indels interrupting the reading frame of an inserted sequence and be of importance for HDR template design in general.
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Affiliation(s)
- Anne Hege Straume
- Institute of Marine Research, P.O. Box 1870, Nordnes, NO-5817, Bergen, Norway
| | - Erik Kjærner-Semb
- Institute of Marine Research, P.O. Box 1870, Nordnes, NO-5817, Bergen, Norway
| | - Kai Ove Skaftnesmo
- Institute of Marine Research, P.O. Box 1870, Nordnes, NO-5817, Bergen, Norway
| | - Hilal Güralp
- Institute of Marine Research, P.O. Box 1870, Nordnes, NO-5817, Bergen, Norway
| | - Lene Kleppe
- Institute of Marine Research, P.O. Box 1870, Nordnes, NO-5817, Bergen, Norway
| | - Anna Wargelius
- Institute of Marine Research, P.O. Box 1870, Nordnes, NO-5817, Bergen, Norway
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Tonelli F, Bek JW, Besio R, De Clercq A, Leoni L, Salmon P, Coucke PJ, Willaert A, Forlino A. Zebrafish: A Resourceful Vertebrate Model to Investigate Skeletal Disorders. Front Endocrinol (Lausanne) 2020; 11:489. [PMID: 32849280 PMCID: PMC7416647 DOI: 10.3389/fendo.2020.00489] [Citation(s) in RCA: 68] [Impact Index Per Article: 17.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Submit a Manuscript] [Subscribe] [Scholar Register] [Received: 04/25/2020] [Accepted: 06/22/2020] [Indexed: 12/11/2022] Open
Abstract
Animal models are essential tools for addressing fundamental scientific questions about skeletal diseases and for the development of new therapeutic approaches. Traditionally, mice have been the most common model organism in biomedical research, but their use is hampered by several limitations including complex generation, demanding investigation of early developmental stages, regulatory restrictions on breeding, and high maintenance cost. The zebrafish has been used as an efficient alternative vertebrate model for the study of human skeletal diseases, thanks to its easy genetic manipulation, high fecundity, external fertilization, transparency of rapidly developing embryos, and low maintenance cost. Furthermore, zebrafish share similar skeletal cells and ossification types with mammals. In the last decades, the use of both forward and new reverse genetics techniques has resulted in the generation of many mutant lines carrying skeletal phenotypes associated with human diseases. In addition, transgenic lines expressing fluorescent proteins under bone cell- or pathway- specific promoters enable in vivo imaging of differentiation and signaling at the cellular level. Despite the small size of the zebrafish, many traditional techniques for skeletal phenotyping, such as x-ray and microCT imaging and histological approaches, can be applied using the appropriate equipment and custom protocols. The ability of adult zebrafish to remodel skeletal tissues can be exploited as a unique tool to investigate bone formation and repair. Finally, the permeability of embryos to chemicals dissolved in water, together with the availability of large numbers of small-sized animals makes zebrafish a perfect model for high-throughput bone anabolic drug screening. This review aims to discuss the techniques that make zebrafish a powerful model to investigate the molecular and physiological basis of skeletal disorders.
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Affiliation(s)
- Francesca Tonelli
- Biochemistry Unit, Department of Molecular Medicine, University of Pavia, Pavia, Italy
| | - Jan Willem Bek
- Department of Biomolecular Medicine, Center of Medical Genetics, Ghent University-University Hospital, Ghent, Belgium
| | - Roberta Besio
- Biochemistry Unit, Department of Molecular Medicine, University of Pavia, Pavia, Italy
| | - Adelbert De Clercq
- Department of Biomolecular Medicine, Center of Medical Genetics, Ghent University-University Hospital, Ghent, Belgium
| | - Laura Leoni
- Biochemistry Unit, Department of Molecular Medicine, University of Pavia, Pavia, Italy
| | | | - Paul J. Coucke
- Department of Biomolecular Medicine, Center of Medical Genetics, Ghent University-University Hospital, Ghent, Belgium
| | - Andy Willaert
- Department of Biomolecular Medicine, Center of Medical Genetics, Ghent University-University Hospital, Ghent, Belgium
| | - Antonella Forlino
- Biochemistry Unit, Department of Molecular Medicine, University of Pavia, Pavia, Italy
- *Correspondence: Antonella Forlino
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Hypomorphic zebrafish models mimic the musculoskeletal phenotype of β4GalT7-deficient Ehlers-Danlos syndrome. Matrix Biol 2019; 89:59-75. [PMID: 31862401 DOI: 10.1016/j.matbio.2019.12.002] [Citation(s) in RCA: 13] [Impact Index Per Article: 2.6] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/20/2019] [Revised: 11/06/2019] [Accepted: 12/12/2019] [Indexed: 02/02/2023]
Abstract
β4GalT7 is a transmembrane Golgi enzyme, encoded by B4GALT7, that plays a pivotal role in the proteoglycan linker region formation during proteoglycan biosynthesis. Defects in this enzyme give rise to a rare autosomal recessive form of Ehlers-Danlos syndrome (EDS), currently known as 'spondylodysplastic EDS (spEDS-B4GALT7)'. This EDS subtype is mainly characterized by short stature, hypotonia and skeletal abnormalities, thereby illustrating its pleiotropic importance during human development. Insights into the pathogenic mechanisms underlying this disabling disease are very limited, in part due to the lack of a relevant in vivo model. As the majority of mutations identified in patients with spEDS-B4GALT7 are hypomorphic, we generated zebrafish models with partial loss of B4galt7 function, including different knockdown (morphant) and mosaic knockout (crispant) b4galt7 zebrafish models and studied the morphologic, functional and molecular aspects in embryonic and larval stages. Morphant and crispant zebrafish show highly similar morphological abnormalities in early development including a small, round head, bowed pectoral fins, short body-axis and mild developmental delay. Several craniofacial cartilage and bone structures are absent or strongly misshapen. In addition, the total amount of sulfated glycosaminoglycans is significantly diminished and particularly heparan and chondroitin sulfate proteoglycan levels are greatly reduced. We also show impaired cartilage patterning and loss of chondrocyte organization in a cartilage-specific Tg(Col2a1aBAC:mcherry) zebrafish reporter line. The occurrence of the same abnormalities in the different models confirms these are specifically caused by B4galt7 deficiency. A disturbed actin pattern, along with a lack of muscle tone, was only noted in morphants in which translation of b4galt7 was blocked. In conclusion, we generated the first viable animal models for spEDS-B4GALT7, and show that in early development the human spEDS-B4GALT7 phenotype is faithfully mimicked in these zebrafish models. Our findings underscore a key role for β4GalT7 in early development of cartilage, bone and muscle. These models will lead to a better understanding of spEDS-B4GALT7 and can be used in future efforts focusing on therapeutic applications.
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Sansbury BM, Hewes AM, Kmiec EB. Understanding the diversity of genetic outcomes from CRISPR-Cas generated homology-directed repair. Commun Biol 2019; 2:458. [PMID: 31840103 PMCID: PMC6898364 DOI: 10.1038/s42003-019-0705-y] [Citation(s) in RCA: 26] [Impact Index Per Article: 5.2] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/29/2019] [Accepted: 11/11/2019] [Indexed: 02/06/2023] Open
Abstract
As CRISPR-Cas systems advance toward clinical application, it is essential to identify all the outcomes of gene-editing activity in human cells. Reports highlighting the remarkable success of homology-directed repair (HDR) in the treatment of inherited diseases may inadvertently underreport the collateral activity of this remarkable technology. We are utilizing an in vitro gene-editing system in which a CRISPR-Cas complex provides the double-stranded cleavage and a mammalian cell-free extract provides the enzymatic activity to promote non-homologous end joining, micro-homology mediated end joining, and homology-directed repair. Here, we detail the broad spectrum of gene-editing reaction outcomes utilizing Cas9 and Cas12a in combination with single-stranded donor templates of the sense and nonsense polarity. This system offers the opportunity to see the range of outcomes of gene-editing reactions in an unbiased fashion, detailing the distribution of DNA repair outcomes as a function of a set of genetic tools.
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Affiliation(s)
- Brett M. Sansbury
- Gene Editing Institute, Helen F. Graham Cancer Center & Research Institute, Christiana Care Health System, Newark, DE USA
- Department of Medical and Molecular Sciences, University of Delaware, Newark, DE USA
| | - Amanda M. Hewes
- Department of Medical and Molecular Sciences, University of Delaware, Newark, DE USA
| | - Eric B. Kmiec
- Gene Editing Institute, Helen F. Graham Cancer Center & Research Institute, Christiana Care Health System, Newark, DE USA
- Department of Medical and Molecular Sciences, University of Delaware, Newark, DE USA
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Konantz M, Schürch C, Hanns P, Müller JS, Sauteur L, Lengerke C. Modeling hematopoietic disorders in zebrafish. Dis Model Mech 2019; 12:12/9/dmm040360. [PMID: 31519693 PMCID: PMC6765189 DOI: 10.1242/dmm.040360] [Citation(s) in RCA: 18] [Impact Index Per Article: 3.6] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/11/2022] Open
Abstract
Zebrafish offer a powerful vertebrate model for studies of development and disease. The major advantages of this model include the possibilities of conducting reverse and forward genetic screens and of observing cellular processes by in vivo imaging of single cells. Moreover, pathways regulating blood development are highly conserved between zebrafish and mammals, and several discoveries made in fish were later translated to murine and human models. This review and accompanying poster provide an overview of zebrafish hematopoiesis and discuss the existing zebrafish models of blood disorders, such as myeloid and lymphoid malignancies, bone marrow failure syndromes and immunodeficiencies, with a focus on how these models were generated and how they can be applied for translational research. Summary: This At A Glance article and poster summarize the last 20 years of research in zebrafish models for hematopoietic disorders, highlighting how these models were created and are being applied for translational research.
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Affiliation(s)
- Martina Konantz
- Department of Biomedicine, University of Basel and University Hospital Basel, Basel 4031, Switzerland
| | - Christoph Schürch
- Department of Biomedicine, University of Basel and University Hospital Basel, Basel 4031, Switzerland
| | - Pauline Hanns
- Department of Biomedicine, University of Basel and University Hospital Basel, Basel 4031, Switzerland
| | - Joëlle S Müller
- Department of Biomedicine, University of Basel and University Hospital Basel, Basel 4031, Switzerland
| | - Loïc Sauteur
- Department of Biomedicine, University of Basel and University Hospital Basel, Basel 4031, Switzerland
| | - Claudia Lengerke
- Department of Biomedicine, University of Basel and University Hospital Basel, Basel 4031, Switzerland.,Division of Hematology, University of Basel and University Hospital Basel, Basel 4031, Switzerland
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DNA repair and neurological disease: From molecular understanding to the development of diagnostics and model organisms. DNA Repair (Amst) 2019; 81:102669. [PMID: 31331820 DOI: 10.1016/j.dnarep.2019.102669] [Citation(s) in RCA: 24] [Impact Index Per Article: 4.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/12/2022]
Abstract
In both replicating and non-replicating cells, the maintenance of genomic stability is of utmost importance. Dividing cells can repair DNA damage during cell division, tolerate the damage by employing potentially mutagenic DNA polymerases or die via apoptosis. However, the options for accurate DNA repair are more limited in non-replicating neuronal cells. If DNA damage is left unresolved, neuronal cells die causing neurodegenerative disorders. A number of pathogenic variants of DNA repair proteins have been linked to multiple neurological diseases. The current challenge is to harness our knowledge of fundamental properties of DNA repair to improve diagnosis, prognosis and treatment of such debilitating disorders. In this perspective, we will focus on recent efforts in identifying novel DNA repair biomarkers for the diagnosis of neurological disorders and their use in monitoring the patient response to therapy. These efforts are greatly facilitated by the development of model organisms such as zebrafish, which will also be summarised.
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