1
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Liu Y, Kong J, Liu G, Li Z, Xiao Y. Precise Gene Knock-In Tools with Minimized Risk of DSBs: A Trend for Gene Manipulation. ADVANCED SCIENCE (WEINHEIM, BADEN-WURTTEMBERG, GERMANY) 2024; 11:e2401797. [PMID: 38728624 PMCID: PMC11267366 DOI: 10.1002/advs.202401797] [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: 02/20/2024] [Revised: 04/29/2024] [Indexed: 05/12/2024]
Abstract
Gene knock-in refers to the insertion of exogenous functional genes into a target genome to achieve continuous expression. Currently, most knock-in tools are based on site-directed nucleases, which can induce double-strand breaks (DSBs) at the target, following which the designed donors carrying functional genes can be inserted via the endogenous gene repair pathway. The size of donor genes is limited by the characteristics of gene repair, and the DSBs induce risks like genotoxicity. New generation tools, such as prime editing, transposase, and integrase, can insert larger gene fragments while minimizing or eliminating the risk of DSBs, opening new avenues in the development of animal models and gene therapy. However, the elimination of off-target events and the production of delivery carriers with precise requirements remain challenging, restricting the application of the current knock-in treatments to mainly in vitro settings. Here, a comprehensive review of the knock-in tools that do not/minimally rely on DSBs and use other mechanisms is provided. Moreover, the challenges and recent advances of in vivo knock-in treatments in terms of the therapeutic process is discussed. Collectively, the new generation of DSBs-minimizing and large-fragment knock-in tools has revolutionized the field of gene editing, from basic research to clinical treatment.
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Affiliation(s)
- Yongfeng Liu
- Department of PharmacologySchool of PharmacyChina Pharmaceutical UniversityNanjing210009China
- State Key Laboratory of Natural MedicinesChina Pharmaceutical UniversityNanjing210009China
- Mudi Meng Honors CollegeChina Pharmaceutical UniversityNanjing210009China
| | - Jianping Kong
- Department of PharmacologySchool of PharmacyChina Pharmaceutical UniversityNanjing210009China
- State Key Laboratory of Natural MedicinesChina Pharmaceutical UniversityNanjing210009China
| | - Gongyu Liu
- Department of PharmacologySchool of PharmacyChina Pharmaceutical UniversityNanjing210009China
- State Key Laboratory of Natural MedicinesChina Pharmaceutical UniversityNanjing210009China
| | - Zhaoxing Li
- Department of PharmacologySchool of PharmacyChina Pharmaceutical UniversityNanjing210009China
- State Key Laboratory of Natural MedicinesChina Pharmaceutical UniversityNanjing210009China
- Chongqing Innovation Institute of China Pharmaceutical UniversityChongqing401135China
| | - Yibei Xiao
- Department of PharmacologySchool of PharmacyChina Pharmaceutical UniversityNanjing210009China
- State Key Laboratory of Natural MedicinesChina Pharmaceutical UniversityNanjing210009China
- Chongqing Innovation Institute of China Pharmaceutical UniversityChongqing401135China
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2
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Ryu J, Statz JP, Chan W, Oyama K, Custer M, Wienisch M, Chen R, Hanna CB, Hennebold JD. Generation of Rhesus Macaque Embryos with Expanded CAG Trinucleotide Repeats in the Huntingtin Gene. Cells 2024; 13:829. [PMID: 38786052 PMCID: PMC11119628 DOI: 10.3390/cells13100829] [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: 02/01/2024] [Revised: 05/03/2024] [Accepted: 05/08/2024] [Indexed: 05/25/2024] Open
Abstract
Huntington's disease (HD) arises from expanded CAG repeats in exon 1 of the Huntingtin (HTT) gene. The resultant misfolded HTT protein accumulates within neuronal cells, negatively impacting their function and survival. Ultimately, HTT accumulation results in cell death, causing the development of HD. A nonhuman primate (NHP) HD model would provide important insight into disease development and the generation of novel therapies due to their genetic and physiological similarity to humans. For this purpose, we tested CRISPR/Cas9 and a single-stranded DNA (ssDNA) containing expanded CAG repeats in introducing an expanded CAG repeat into the HTT gene in rhesus macaque embryos. Analyses were conducted on arrested embryos and trophectoderm (TE) cells biopsied from blastocysts to assess the insertion of the ssDNA into the HTT gene. Genotyping results demonstrated that 15% of the embryos carried an expanded CAG repeat. The integration of an expanded CAG repeat region was successfully identified in five blastocysts, which were cryopreserved for NHP HD animal production. Some off-target events were observed in biopsies from the cryopreserved blastocysts. NHP embryos were successfully produced, which will help to establish an NHP HD model and, ultimately, may serve as a vital tool for better understanding HD's pathology and developing novel treatments.
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Affiliation(s)
- Junghyun Ryu
- Division of Reproductive & Developmental Sciences, Oregon National Primate Research Center, Oregon Health & Science University, Beaverton, OR 97006, USA; (J.R.); (J.P.S.); (W.C.); (K.O.); (M.C.); (C.B.H.)
| | - John P. Statz
- Division of Reproductive & Developmental Sciences, Oregon National Primate Research Center, Oregon Health & Science University, Beaverton, OR 97006, USA; (J.R.); (J.P.S.); (W.C.); (K.O.); (M.C.); (C.B.H.)
| | - William Chan
- Division of Reproductive & Developmental Sciences, Oregon National Primate Research Center, Oregon Health & Science University, Beaverton, OR 97006, USA; (J.R.); (J.P.S.); (W.C.); (K.O.); (M.C.); (C.B.H.)
| | - Kiana Oyama
- Division of Reproductive & Developmental Sciences, Oregon National Primate Research Center, Oregon Health & Science University, Beaverton, OR 97006, USA; (J.R.); (J.P.S.); (W.C.); (K.O.); (M.C.); (C.B.H.)
| | - Maggie Custer
- Division of Reproductive & Developmental Sciences, Oregon National Primate Research Center, Oregon Health & Science University, Beaverton, OR 97006, USA; (J.R.); (J.P.S.); (W.C.); (K.O.); (M.C.); (C.B.H.)
| | - Martin Wienisch
- Department of Brain and Cognitive Sciences, McGovern Institute for Brain Research, Massachusetts Institute of Technology (MIT), Cambridge, MA 02139, USA;
| | | | - Carol B. Hanna
- Division of Reproductive & Developmental Sciences, Oregon National Primate Research Center, Oregon Health & Science University, Beaverton, OR 97006, USA; (J.R.); (J.P.S.); (W.C.); (K.O.); (M.C.); (C.B.H.)
- Assisted Reproductive Technologies Core, Oregon National Primate Research Center, Oregon Health & Science University, Beaverton, OR 97006, USA
| | - Jon D. Hennebold
- Division of Reproductive & Developmental Sciences, Oregon National Primate Research Center, Oregon Health & Science University, Beaverton, OR 97006, USA; (J.R.); (J.P.S.); (W.C.); (K.O.); (M.C.); (C.B.H.)
- Department of Obstetrics & Gynecology, Oregon Health & Science University, Portland, OR 97239, USA
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3
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Davison C, Harzman H, Nicholson J, Entriken S, Mobley K, Krull A, Singhal M, Skow C, Matthews N, Kopp L, Gillette B, Weide TJ, Hukvari JR, Stumpf SC, Feldmann OM, McGrail M, Srivastava R, Essner JJ. Tagging the tjp1a Gene in Zebrafish with Monomeric Red Fluorescent Protein Using Biotin Homology Arms. Zebrafish 2024; 21:191-197. [PMID: 38621205 PMCID: PMC11035848 DOI: 10.1089/zeb.2023.0096] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 04/17/2024] Open
Abstract
Tjp1a and other tight junction and adherens proteins play important roles in cell-cell adhesion, scaffolding, and forming seals between cells in epithelial and endothelial tissues. In this study, we labeled Tjp1a of zebrafish with the monomeric red fluorescent protein (mRFP) using CRISPR/Cas9-mediated targeted integration of biotin-labeled polymerase chain reaction (PCR) generated templates. Labeling Tjp1a with RFP allowed us to follow membrane and junctional dynamics of epithelial and endothelial cells throughout zebrafish embryo development. For targeted integration, we used short 35 bp homology arms on each side of the Cas9 genomic target site at the C-terminal of the coding sequence in tjp1a. Through PCR using 5' biotinylated primers containing the homology arms, we generated a double-stranded template for homology directed repair containing a flexible linker followed by RFP. Cas9 protein was complexed with the tjp1a gRNA before mixing with the repair template and microinjected into one-cell zebrafish embryos. We confirmed and recovered a precise integration allele at the desired site at the tjp1a C-terminus. Examination of fluorescence reveals RFP cell-cell junctional labeling using confocal imaging. We are currently using this stable tjp1a-mRFPis86 line to examine the behavior and interactions between cells during vascular formation in zebrafish.
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Affiliation(s)
- Connor Davison
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
| | - Hamelynn Harzman
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
| | - Jessie Nicholson
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
| | - Seth Entriken
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
| | - Kierinn Mobley
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
| | - Abigail Krull
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
| | - Manik Singhal
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
| | - Caleb Skow
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
| | - Nathan Matthews
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
| | - Lindsey Kopp
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
| | - Benjamin Gillette
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
| | - Tyler J. Weide
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
| | - Jana R. Hukvari
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
| | - Sofia C.P. Stumpf
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
| | - Olivia M. Feldmann
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
| | - Maura McGrail
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
| | - Renu Srivastava
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
| | - Jeffrey J. Essner
- Department of Genetics, Development and Cell Biology, Iowa State University, Ames, Iowa, USA
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4
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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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5
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Li Y, Huang B, Chen J, Huang L, Xu J, Wang Y, Cui G, Zhao H, Xin B, Song W, Zhu J, Lai J. Targeted large fragment deletion in plants using paired crRNAs with type I CRISPR system. PLANT BIOTECHNOLOGY JOURNAL 2023; 21:2196-2208. [PMID: 37641539 PMCID: PMC10579709 DOI: 10.1111/pbi.14122] [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: 02/08/2023] [Revised: 05/19/2023] [Accepted: 06/25/2023] [Indexed: 08/31/2023]
Abstract
The CRISPR-Cas systems have been widely used as genome editing tools, with type II and V systems typically introducing small indels, and type I system mediating long-range deletions. However, the precision of type I systems for large fragment deletion is still remained to be optimized. Here, we developed a compact Cascade-Cas3 Dvu I-C system with Cas11c for plant genome editing. The Dvu I-C system was efficient to introduce controllable large fragment deletion up to at least 20 kb using paired crRNAs. The paired-crRNAs design also improved the controllability of deletions for the type I-E system. Dvu I-C system was sensitive to spacer length and mismatch, which was benefit for target specificity. In addition, we showed that the Dvu I-C system was efficient for generating stable transgenic lines in maize and rice with the editing efficiency up to 86.67%. Overall, Dvu I-C system we developed here is powerful for achieving controllable large fragment deletions.
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Affiliation(s)
- Yingnan Li
- State Key Laboratory of Maize Bio‐breeding, National Maize Improvement Center, Department of Plant Genetics and BreedingChina Agricultural UniversityBeijingChina
| | - Boyu Huang
- Shanghai Center for Plant Stress Biology, CAS Center for Excellence in Molecular Plant SciencesChinese Academy of SciencesShanghaiChina
- University of Chinese Academy of SciencesBeijingChina
| | - Jian Chen
- State Key Laboratory of Maize Bio‐breeding, National Maize Improvement Center, Department of Plant Genetics and BreedingChina Agricultural UniversityBeijingChina
| | - Liangliang Huang
- State Key Laboratory of Maize Bio‐breeding, National Maize Improvement Center, Department of Plant Genetics and BreedingChina Agricultural UniversityBeijingChina
| | - Jianghai Xu
- College of Biological SciencesChina Agricultural UniversityBeijingChina
| | - Yingying Wang
- College of Biological SciencesChina Agricultural UniversityBeijingChina
| | - Guanghui Cui
- State Key Laboratory of Maize Bio‐breeding, National Maize Improvement Center, Department of Plant Genetics and BreedingChina Agricultural UniversityBeijingChina
| | - Haiming Zhao
- State Key Laboratory of Maize Bio‐breeding, National Maize Improvement Center, Department of Plant Genetics and BreedingChina Agricultural UniversityBeijingChina
| | - Beibei Xin
- State Key Laboratory of Maize Bio‐breeding, National Maize Improvement Center, Department of Plant Genetics and BreedingChina Agricultural UniversityBeijingChina
| | - Weibin Song
- State Key Laboratory of Maize Bio‐breeding, National Maize Improvement Center, Department of Plant Genetics and BreedingChina Agricultural UniversityBeijingChina
| | - Jian‐Kang Zhu
- Institute of Advanced Biotechnology and School of Life SciencesSouthern University of Science and TechnologyShenzhenChina
- Center for Advanced Bioindustry TechnologiesChinese Academy of Agricultural SciencesBeijingChina
| | - Jinsheng Lai
- State Key Laboratory of Maize Bio‐breeding, National Maize Improvement Center, Department of Plant Genetics and BreedingChina Agricultural UniversityBeijingChina
- Frontiers Science Center for Molecular Design BreedingChina Agricultural UniversityBeijingChina
- Center for Crop Functional Genomics and Molecular BreedingChina Agricultural UniversityBeijingChina
- Sanya Institute of China Agricultural UniversitySanyaChina
- Hainan Yazhou Bay Seed LaboratorySanyaChina
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6
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Clark B, Kuwalekar M, Fischer B, Woltering J, Biran J, Juntti S, Kratochwil CF, Santos ME, Almeida MV. Genome editing in East African cichlids and tilapias: state-of-the-art and future directions. Open Biol 2023; 13:230257. [PMID: 38018094 PMCID: PMC10685126 DOI: 10.1098/rsob.230257] [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: 08/02/2023] [Accepted: 10/27/2023] [Indexed: 11/30/2023] Open
Abstract
African cichlid fishes of the Cichlidae family are a group of teleosts important for aquaculture and research. A thriving research community is particularly interested in the cichlid radiations of the East African Great Lakes. One key goal is to pinpoint genetic variation underlying phenotypic diversification, but the lack of genetic tools has precluded thorough dissection of the genetic basis of relevant traits in cichlids. Genome editing technologies are well established in teleost models like zebrafish and medaka. However, this is not the case for emerging model organisms, such as East African cichlids, where these technologies remain inaccessible to most laboratories, due in part to limited exchange of knowledge and expertise. The Cichlid Science 2022 meeting (Cambridge, UK) hosted for the first time a Genome Editing Workshop, where the community discussed recent advances in genome editing, with an emphasis on CRISPR/Cas9 technologies. Based on the workshop findings and discussions, in this review we define the state-of-the-art of cichlid genome editing, share resources and protocols, and propose new possible avenues to further expand the cichlid genome editing toolkit.
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Affiliation(s)
- Bethan Clark
- Department of Zoology, University of Cambridge, Cambridge, UK
| | - Muktai Kuwalekar
- Helsinki Institute of Life Science (HiLIFE), University of Helsinki, Helsinki, Uusimaa 00014, Finland
- Faculty of Biological and Environmental Sciences, University of Helsinki, Helsinki, Uusimaa 00014, Finland
| | - Bettina Fischer
- Department of Genetics, University of Cambridge, Cambridge, UK
| | - Joost Woltering
- Zoology and Evolutionary Biology, Department of Biology, University of Konstanz, Konstanz, Baden-Württemberg 78457, Germany
| | - Jakob Biran
- Department of Poultry and Aquaculture, Institute of Animal Sciences, Agricultural Research Organization, Volcani Center, Rishon Lezion, Israel
| | - Scott Juntti
- Department of Biology, University of Maryland, College Park, MD, USA
| | - Claudius F. Kratochwil
- Helsinki Institute of Life Science (HiLIFE), University of Helsinki, Helsinki, Uusimaa 00014, Finland
- Faculty of Biological and Environmental Sciences, University of Helsinki, Helsinki, Uusimaa 00014, Finland
| | | | - Miguel Vasconcelos Almeida
- Department of Biochemistry, University of Cambridge, Cambridge, UK
- Wellcome/CRUK Gurdon Institute, University of Cambridge, Cambridge, UK
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7
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Palmiotti A, Lichak MR, Shih PY, Kwon YM, Bendesky A. Genetic manipulation of betta fish. Front Genome Ed 2023; 5:1167093. [PMID: 37545763 PMCID: PMC10401044 DOI: 10.3389/fgeed.2023.1167093] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/15/2023] [Accepted: 07/07/2023] [Indexed: 08/08/2023] Open
Abstract
Betta splendens, also known as Siamese fighting fish or "betta," is a freshwater fish species renowned for its astonishing morphological diversity and extreme aggressive behavior. Despite recent advances in our understanding of the genetics and neurobiology of betta, the lack of tools to manipulate their genome has hindered progress at functional and mechanistic levels. In this study, we outline the use of three genetic manipulation technologies, which we have optimized for use in betta: CRISPR/Cas9-mediated knockout, CRISPR/Cas9-mediated knockin, and Tol2-mediated transgenesis. We knocked out three genes: alkal2l, bco1l, and mitfa, and analyzed their effects on viability and pigmentation. Furthermore, we knocked in a fluorescent protein into the mitfa locus, a proof-of-principle experiment of this powerful technology in betta. Finally, we used Tol2-mediated transgenesis to create fish with ubiquitous expression of GFP, and then developed a bicistronic plasmid with heart-specific expression of a red fluorescent protein to serve as a visible marker of successful transgenesis. Our work highlights the potential for the genetic manipulation of betta, providing valuable resources for the effective use of genetic tools in this animal model.
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Affiliation(s)
- Alec Palmiotti
- Zuckerman Mind Brain Behavior Institute, Columbia University, New York, NY, United States
- Department of Ecology, Evolution and Environmental Biology, Columbia University, New York, NY, United States
| | - Madison R. Lichak
- Zuckerman Mind Brain Behavior Institute, Columbia University, New York, NY, United States
- Department of Ecology, Evolution and Environmental Biology, Columbia University, New York, NY, United States
| | - Pei-Yin Shih
- Zuckerman Mind Brain Behavior Institute, Columbia University, New York, NY, United States
- Department of Ecology, Evolution and Environmental Biology, Columbia University, New York, NY, United States
| | - Young Mi Kwon
- Zuckerman Mind Brain Behavior Institute, Columbia University, New York, NY, United States
- Department of Ecology, Evolution and Environmental Biology, Columbia University, New York, NY, United States
| | - Andres Bendesky
- Zuckerman Mind Brain Behavior Institute, Columbia University, New York, NY, United States
- Department of Ecology, Evolution and Environmental Biology, Columbia University, New York, NY, United States
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8
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Mabuchi A, Hata S, Genova M, Tei C, Ito KK, Hirota M, Komori T, Fukuyama M, Chinen T, Toyoda A, Kitagawa D. ssDNA is not superior to dsDNA as long HDR donors for CRISPR-mediated endogenous gene tagging in human diploid RPE1 and HCT116 cells. BMC Genomics 2023; 24:289. [PMID: 37248464 DOI: 10.1186/s12864-023-09377-3] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/15/2022] [Accepted: 05/14/2023] [Indexed: 05/31/2023] Open
Abstract
BACKGROUND Recent advances in CRISPR technology have enabled us to perform gene knock-in in various species and cell lines. CRISPR-mediated knock-in requires donor DNA which serves as a template for homology-directed repair (HDR). For knock-in of short sequences or base substitutions, ssDNA donors are frequently used among various other forms of HDR donors, such as linear dsDNA. However, partly due to the complexity of long ssDNA preparation, it remains unclear whether ssDNA is the optimal type of HDR donors for insertion of long transgenes such as fluorescent reporters in human cells. RESULTS In this study, we established a nuclease-based simple method for the preparation of long ssDNA with high yield and purity, and comprehensively compared the performance of ssDNA and dsDNA donors with 90 bases of homology arms for endogenous gene tagging with long transgenes in human diploid RPE1 and HCT116 cells. Quantification using flow cytometry revealed lower efficiency of endogenous fluorescent tagging with ssDNA donors than with dsDNA. By analyzing knock-in outcomes using long-read amplicon sequencing and a classification framework, a variety of mis-integration events were detected regardless of the donor type. Importantly, the ratio of precise insertion was lower with ssDNA donors than with dsDNA. Moreover, in off-target integration analyses using donors without homology arms, ssDNA and dsDNA were comparably prone to non-homologous integration. CONCLUSIONS These results indicate that ssDNA is not superior to dsDNA as long HDR donors with relatively short homology arms for gene knock-in in human RPE1 and HCT116 cells.
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Affiliation(s)
- Akira Mabuchi
- Department of Physiological Chemistry, Graduate School of Pharmaceutical Sciences, The University of Tokyo, Bunkyo, Tokyo, Japan
| | - Shoji Hata
- Department of Physiological Chemistry, Graduate School of Pharmaceutical Sciences, The University of Tokyo, Bunkyo, Tokyo, Japan.
- Precursory Research for Embryonic Science and Technology (PRESTO) Program, Japan Science and Technology Agency, Honcho Kawaguchi, Saitama, Japan.
| | - Mariya Genova
- Zentrum Für Molekulare Biologie, Universität Heidelberg, DKFZ-ZMBH Allianz, Heidelberg, Germany
| | - Chiharu Tei
- Department of Physiological Chemistry, Graduate School of Pharmaceutical Sciences, The University of Tokyo, Bunkyo, Tokyo, Japan
| | - Kei K Ito
- Department of Physiological Chemistry, Graduate School of Pharmaceutical Sciences, The University of Tokyo, Bunkyo, Tokyo, Japan
| | - Masayasu Hirota
- Department of Physiological Chemistry, Graduate School of Pharmaceutical Sciences, The University of Tokyo, Bunkyo, Tokyo, Japan
| | - Takuma Komori
- Department of Physiological Chemistry, Graduate School of Pharmaceutical Sciences, The University of Tokyo, Bunkyo, Tokyo, Japan
| | - Masamitsu Fukuyama
- Department of Physiological Chemistry, Graduate School of Pharmaceutical Sciences, The University of Tokyo, Bunkyo, Tokyo, Japan
| | - Takumi Chinen
- Department of Physiological Chemistry, Graduate School of Pharmaceutical Sciences, The University of Tokyo, Bunkyo, Tokyo, Japan
| | - Atsushi Toyoda
- Comparative Genomics Laboratory and Advanced Genomics Center, National Institute of Genetics, Mishima, Shizuoka, Japan
| | - Daiju Kitagawa
- Department of Physiological Chemistry, Graduate School of Pharmaceutical Sciences, The University of Tokyo, Bunkyo, Tokyo, Japan.
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9
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Palmiotti A, Lichak MR, Shih PY, Bendesky A. Genetic manipulation of betta fish. BIORXIV : THE PREPRINT SERVER FOR BIOLOGY 2023:2023.02.16.528733. [PMID: 36824853 PMCID: PMC9948955 DOI: 10.1101/2023.02.16.528733] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Figures] [Subscribe] [Scholar Register] [Indexed: 02/18/2023]
Abstract
Betta splendens , also known as Siamese fighting fish or 'betta', are renowned for their astonishing morphological diversity and extreme aggressive behavior. Despite recent advances in our understanding of the genetics and neurobiology of betta, the lack of tools to manipulate their genome has hindered progress at functional and mechanistic levels. In this study, we outline the use of three genetic manipulation technologies, which we have optimized for use in betta: CRISPR/Cas9-mediated knockout, CRISPR/Cas9-mediated knockin, and Tol2-mediated transgenesis. We knocked out three genes: alkal2l, bco1l , and mitfa , and analyzed their effects on viability and pigmentation. Furthermore, we successfully knocked in a fluorescent protein into the mitfa locus, a proof-of-principle experiment of this powerful technology in betta. Finally, we used Tol2-mediated transgenesis to create fish with ubiquitous expression of GFP, and then developed a bicistronic plasmid with heart-specific expression of a red fluorescent protein to serve as a visible marker of successful transgenesis. Our work highlights the potential for the genetic manipulation of betta, providing valuable resources for the effective use of genetic tools in this animal model.
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Affiliation(s)
- Alec Palmiotti
- Zuckerman Mind Brain Behavior Institute, Columbia University, NY, USA
- Department of Ecology, Evolution and Environmental Biology, Columbia University, NY, USA
| | - Madison R Lichak
- Zuckerman Mind Brain Behavior Institute, Columbia University, NY, USA
- Department of Ecology, Evolution and Environmental Biology, Columbia University, NY, USA
- Present address: Department of Ecology and Evolutionary Biology, Princeton University, NJ, USA
| | - Pei-Yin Shih
- Zuckerman Mind Brain Behavior Institute, Columbia University, NY, USA
- Department of Ecology, Evolution and Environmental Biology, Columbia University, NY, USA
| | - Andres Bendesky
- Zuckerman Mind Brain Behavior Institute, Columbia University, NY, USA
- Department of Ecology, Evolution and Environmental Biology, Columbia University, NY, USA
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10
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Richardson C, Kelsh RN, J. Richardson R. New advances in CRISPR/Cas-mediated precise gene-editing techniques. Dis Model Mech 2023; 16:dmm049874. [PMID: 36847161 PMCID: PMC10003097 DOI: 10.1242/dmm.049874] [Citation(s) in RCA: 7] [Impact Index Per Article: 7.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 03/01/2023] Open
Abstract
Over the past decade, CRISPR/Cas-based gene editing has become a powerful tool for generating mutations in a variety of model organisms, from Escherichia coli to zebrafish, rodents and large mammals. CRISPR/Cas-based gene editing effectively generates insertions or deletions (indels), which allow for rapid gene disruption. However, a large proportion of human genetic diseases are caused by single-base-pair substitutions, which result in more subtle alterations to protein function, and which require more complex and precise editing to recreate in model systems. Precise genome editing (PGE) methods, however, typically have efficiencies of less than a tenth of those that generate less-specific indels, and so there has been a great deal of effort to improve PGE efficiency. Such optimisations include optimal guide RNA and mutation-bearing donor DNA template design, modulation of DNA repair pathways that underpin how edits result from Cas-induced cuts, and the development of Cas9 fusion proteins that introduce edits via alternative mechanisms. In this Review, we provide an overview of the recent progress in optimising PGE methods and their potential for generating models of human genetic disease.
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Affiliation(s)
- Chris Richardson
- School of Physiology, Pharmacology and Neuroscience, Faculty of Biomedical Sciences, University of Bristol, Bristol BS8 1TD, UK
| | - Robert N. Kelsh
- Department of Life Sciences, University of Bath, Bath BA2 7AY, UK
| | - Rebecca J. Richardson
- School of Physiology, Pharmacology and Neuroscience, Faculty of Biomedical Sciences, University of Bristol, Bristol BS8 1TD, UK
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11
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Kamachi Y, Kawahara A. CRISPR-Cas9-Mediated Genome Modifications in Zebrafish. Methods Mol Biol 2023; 2637:313-324. [PMID: 36773157 DOI: 10.1007/978-1-0716-3016-7_24] [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: 04/27/2023]
Abstract
CRISPR-Cas9 genome editing technology has been successfully applied to generate various genetic modifications in zebrafish. The CRISPR-Cas9 system, which originally consisted of three components, CRISPR RNA (crRNA), trans-activating crRNA (tracrRNA), and Cas9, efficiently induces DNA double-strand breaks (DSBs) at targeted genomic loci, often resulting in frameshift-mediated target gene disruption (knockout). However, it remains difficult to perform the targeted integration of exogenous DNA fragments (knock-in) with CRISPR-Cas9. DSBs can be restored through DNA repair mechanisms, such as nonhomologous end joining (NHEJ), microhomology-mediated end joining (MMEJ), and homology-directed repair (HDR). One of our two research groups established a method for the precise MMEJ-mediated targeted integrations of exogenous genes containing homologous microhomology sequences flanking a targeted genomic locus in zebrafish. The other group recently developed a method for knocking in ~200 nt sequences encoding composite tags using long single-stranded DNA (ssDNA) donors. This chapter summarizes the CRISPR-Cas9-mediated genome modification strategy in zebrafish.
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Affiliation(s)
- Yusuke Kamachi
- School of Environmental Science and Engineering, Kochi University of Technology, Tosayamada-cho, Kami, Kochi, Japan
| | - Atsuo Kawahara
- Laboratory for Developmental Biology, Graduate School of Medical Science, University of Yamanashi, Chuo, Yamanashi, Japan.
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12
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Shakirova A, Karpov T, Komarova Y, Lepik K. In search of an ideal template for therapeutic genome editing: A review of current developments for structure optimization. Front Genome Ed 2023; 5:1068637. [PMID: 36911237 PMCID: PMC9992834 DOI: 10.3389/fgeed.2023.1068637] [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: 10/13/2022] [Accepted: 02/08/2023] [Indexed: 02/24/2023] Open
Abstract
Gene therapy is a fast developing field of medicine with hundreds of ongoing early-stage clinical trials and numerous preclinical studies. Genome editing (GE) now is an increasingly important technology for achieving stable therapeutic effect in gene correction, with hematopoietic cells representing a key target cell population for developing novel treatments for a number of hereditary diseases, infections and cancer. By introducing a double strand break (DSB) in the defined locus of genomic DNA, GE tools allow to knockout the desired gene or to knock-in the therapeutic gene if provided with an appropriate repair template. Currently, the efficiency of methods for GE-mediated knock-in is limited. Significant efforts were focused on improving the parameters and interaction of GE nuclease proteins. However, emerging data suggests that optimal characteristics of repair templates may play an important role in the knock-in mechanisms. While viral vectors with notable example of AAVs as a donor template carrier remain the mainstay in many preclinical trials, non-viral templates, including plasmid and linear dsDNA, long ssDNA templates, single and double-stranded ODNs, represent a promising alternative. Furthermore, tuning of editing conditions for the chosen template as well as its structure, length, sequence optimization, homology arm (HA) modifications may have paramount importance for achieving highly efficient knock-in with favorable safety profile. This review outlines the current developments in optimization of templates for the GE mediated therapeutic gene correction.
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Affiliation(s)
- Alena Shakirova
- RM Gorbacheva Research Institute of Pediatric Oncology, Hematology and Transplantation, Pavlov University, Saint Petersburg, Russia
| | - Timofey Karpov
- RM Gorbacheva Research Institute of Pediatric Oncology, Hematology and Transplantation, Pavlov University, Saint Petersburg, Russia.,Peter the Great St. Petersburg Polytechnic University, Saint Petersburg, Russia
| | - Yaroslava Komarova
- RM Gorbacheva Research Institute of Pediatric Oncology, Hematology and Transplantation, Pavlov University, Saint Petersburg, Russia
| | - Kirill Lepik
- RM Gorbacheva Research Institute of Pediatric Oncology, Hematology and Transplantation, Pavlov University, Saint Petersburg, Russia
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13
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Martin SA. A CRISPR/Cas-Based Method for Precise DNA Integration in Xenopus laevis Oocytes Followed by Intracytoplasmic Sperm Injection (ICSI) Fertilization. Methods Mol Biol 2023; 2633:131-143. [PMID: 36853462 DOI: 10.1007/978-1-0716-3004-4_11] [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] [Indexed: 03/01/2023]
Abstract
Xenopus has long had a reputation for being a powerful model organism for use in developmental cell and biochemistry research. With the advent of gene-editing technologies, and the full genome sequencing of Xenopus genomes revealing the extent of the genetic conservation between Xenopus and humans, Xenopus has the potential to become an ideal model for human genetic disease. However, the inability to produce non-mosaic, precise DNA insertions through homology directed repair has limited the strength of Xenopus this field. Furthermore, it has prevented researchers from taking full advantage of fusion tagging, a method for directly tagging genes with either epitope or fluorescent tags, allowing the visualization, quantification, and tracking of proteins without the use of protein-specific antibodies. Here, we describe a method for precise DNA insertion into oocytes using CRISPR/Cas9, followed by in vitro maturation and fertilization by intracytoplasmic sperm injection (ICSI), culminating in the production of embryos carrying a non-mosaic, heterozygous insertion.
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Affiliation(s)
- Sian Angela Martin
- European Xenopus Resource Centre (EXRC), University of Portsmouth, Portsmouth, UK.
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14
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Carrington B, Ramanagoudr-Bhojappa R, Bresciani E, Han TU, Sood R. 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: 3] [Impact Index Per Article: 1.5] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [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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Affiliation(s)
- Blake Carrington
- Translational and Functional Genomics Branch, Zebrafish Core, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD, 20892, USA
| | - Ramanagouda Ramanagoudr-Bhojappa
- Cancer Genetics Unit, Cancer Genetics and Comparative Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD, 20892, USA
| | - Erica Bresciani
- Oncogenesis and Development Section, Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD, 20892, USA
| | - Tae-Un Han
- Molecular Neurogenetics Section, Medical Genetics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD, 20892, USA
| | - Raman Sood
- Translational and Functional Genomics Branch, Zebrafish Core, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD, 20892, USA.
- Oncogenesis and Development Section, Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD, 20892, USA.
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15
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Dudziak K, Nowak M, Sozoniuk M. One Host-Multiple Applications: Zebrafish ( Danio rerio) as Promising Model for Studying Human Cancers and Pathogenic Diseases. Int J Mol Sci 2022; 23:10255. [PMID: 36142160 PMCID: PMC9499349 DOI: 10.3390/ijms231810255] [Citation(s) in RCA: 4] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/30/2022] [Revised: 09/03/2022] [Accepted: 09/03/2022] [Indexed: 11/17/2022] Open
Abstract
In recent years, zebrafish (ZF) has been increasingly applied as a model in human disease studies, with a particular focus on cancer. A number of advantages make it an attractive alternative for mice widely used so far. Due to the many advantages of zebrafish, modifications can be based on different mechanisms and the induction of human disease can take different forms depending on the research goal. Genetic manipulation, tumor transplantation, or injection of the pathogen are only a few examples of using ZF as a model. Most of the studies are conducted in order to understand the disease mechanism, monitor disease progression, test new or alternative therapies, and select the best treatment. The transplantation of cancer cells derived from patients enables the development of personalized medicine. To better mimic a patient's body environment, immune-deficient models (SCID) have been developed. A lower immune response is mostly generated by genetic manipulation but also by irradiation or dexamethasone treatment. For many studies, using SCID provides a better chance to avoid cancer cell rejection. In this review, we describe the main directions of using ZF in research, explain why and how zebrafish can be used as a model, what kind of limitations will be met and how to overcome them. We collected recent achievements in this field, indicating promising perspectives for the future.
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Affiliation(s)
- Karolina Dudziak
- Chair and Department of Biochemistry and Molecular Biology, Medical University of Lublin, 20-059 Lublin, Poland
| | - Michał Nowak
- Institute of Plant Genetics, Breeding and Biotechnology, University of Life Sciences in Lublin, 20-950 Lublin, Poland
| | - Magdalena Sozoniuk
- Institute of Plant Genetics, Breeding and Biotechnology, University of Life Sciences in Lublin, 20-950 Lublin, Poland
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16
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Petel Légaré V, Rampal CJ, Gurberg TJN, Harji ZA, Allard-Chamard X, Rodríguez EC, Armstrong GAB. Development of an endogenously myc-tagged TARDBP (TDP-43) zebrafish model using the CRISPR/Cas9 system and homology directed repair. Comp Biochem Physiol B Biochem Mol Biol 2022; 261:110756. [PMID: 35580804 DOI: 10.1016/j.cbpb.2022.110756] [Citation(s) in RCA: 2] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/12/2022] [Revised: 05/08/2022] [Accepted: 05/11/2022] [Indexed: 11/25/2022]
Abstract
Many of the modern advances in cellular biology have been made by the expression of engineered constructs with epitope tags for subsequent biochemical investigations. While the utility of epitope tags has permitted insights in cellular and animal models, these are often expressed using traditional transgenic approaches. Using the CRISPR/Cas9 system and homology directed repair we recombine a single myc epitope sequence following the start codon of the zebrafish ortholog of TARDBP (TDP-43). TDP-43 is an RNA binding protein that is involved in the neurodegenerative disease amyotrophic lateral sclerosis and frontotemporal dementia. We report that zebrafish expressing the myc-tardbp engendered allele produced a stable protein that was detected by both western blot and immunofluorescence. Furthermore, both heterozygous and homozygous carriers of the myc-tardbp allele developed to sexual maturity. We propose that the methodology used here will be useful for zebrafish researchers and other comparative animal biologists interested in developing animal models expressing endogenously tagged proteins.
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Affiliation(s)
- Virginie Petel Légaré
- Department of Neurology and Neurosurgery, Montreal Neurological Institute, Faculty of Medicine, McGill University. https://twitter.com/virginiepet
| | - Christian J Rampal
- Department of Neurology and Neurosurgery, Montreal Neurological Institute, Faculty of Medicine, McGill University. https://twitter.com/ChristianRampal
| | - Tyler J N Gurberg
- Department of Neurology and Neurosurgery, Montreal Neurological Institute, Faculty of Medicine, McGill University
| | - Ziyaan A Harji
- Department of Neurology and Neurosurgery, Montreal Neurological Institute, Faculty of Medicine, McGill University. https://twitter.com/ziyaanharji
| | - Xavier Allard-Chamard
- Department of Neurology and Neurosurgery, Montreal Neurological Institute, Faculty of Medicine, McGill University
| | - Esteban C Rodríguez
- Department of Neurology and Neurosurgery, Montreal Neurological Institute, Faculty of Medicine, McGill University
| | - Gary A B Armstrong
- Department of Neurology and Neurosurgery, Montreal Neurological Institute, Faculty of Medicine, McGill University.
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17
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Krueger LA, Morris AC. Generation of a zebrafish knock-in line expressing MYC-tagged Sox11a using CRISPR/Cas9 genome editing. Biochem Biophys Res Commun 2022; 608:8-13. [PMID: 35378361 PMCID: PMC9050874 DOI: 10.1016/j.bbrc.2022.03.103] [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: 03/17/2022] [Accepted: 03/21/2022] [Indexed: 11/02/2022]
Abstract
Advances in CRISPR-Cas9 genome editing technology have strengthened the role of zebrafish as a model organism for genetics and developmental biology. These tools have led to a significant increase in the production of loss-of-function mutant zebrafish lines. However, the generation of precisely edited knock-in lines has remained a significant challenge in the field due to the decreased efficiency of homology directed repair (HDR). In this study, we overcame some of these challenges by combining available design tools and synthetic, commercially available CRISPR reagents to generate a knock-in line carrying an in-frame MYC epitope tag at the sox11a locus. Zebrafish Sox11a is a transcription factor with critical roles in organogenesis, neurogenesis, craniofacial, and skeletal development; however, only a few direct molecular targets of Sox11a have been identified. Here, we evaluate the knock-in efficiency of various HDR donor configurations and demonstrate the successful expression and localization of the resulting knock-in allele. Our results provide an efficient, streamlined approach to knock-in experiments in zebrafish, which will enable expansion of downstream experimental applications that have previously been difficult to perform. Moreover, the MYC-Sox11a line we have generated will allow further investigation into the function and direct targets of Sox11a.
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Affiliation(s)
- Laura A Krueger
- Department of Biology, University of Kentucky, Lexington, KY, 40506-0225, USA
| | - Ann C Morris
- Department of Biology, University of Kentucky, Lexington, KY, 40506-0225, USA.
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18
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Hillen AEJ, Hruzova M, Rothgangl T, Breur M, Bugiani M, van der Knaap MS, Schwank G, Heine VM. In vivo targeting of a variant causing vanishing white matter using CRISPR/Cas9. Mol Ther Methods Clin Dev 2022; 25:17-25. [PMID: 35317047 PMCID: PMC8917273 DOI: 10.1016/j.omtm.2022.02.006] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/27/2021] [Accepted: 02/19/2022] [Indexed: 11/17/2022]
Abstract
Vanishing white matter (VWM) is a leukodystrophy caused by recessive variants in subunits of eIF2B. At present, no curative treatment is available and patients often die at young age. Due to its monogenic nature, VWM is a promising candidate for the development of CRISPR/Cas9-mediated gene therapy. Here we tested a dual-AAV approach in VWM mice encoding CRISPR/Cas9 and a DNA donor template to correct a pathogenic variant in Eif2b5. We performed sequencing analysis to assess gene correction rates and examined effects on the VWM phenotype, including motor behavior. Sequence analysis demonstrated that over 90% of CRISPR/Cas9-induced edits at the targeted locus are insertion or deletion (indel) mutations, rather than precise corrections from the DNA donor template by homology-directed repair. Around half of the CRISPR/Cas9-treated animals died prematurely. VWM mice showed no improvement in motor skills, weight, or neurological scores at 7 months of age, and CRISPR/Cas9-treated controls displayed an induced VWM phenotype. In conclusion, CRISPR/Cas9-induced DNA double-strand breaks (DSBs) at the Eif2b5 locus did not lead to sufficient correction of the VWM variant. Moreover, indel formation in Eif2b5 induced an exacerbated VWM phenotype. Therefore, DSB-independent strategies like base- or prime editing might better suited for VWM correction.
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Affiliation(s)
- Anne E J Hillen
- Department of Pediatrics and Child Neurology, Emma Children's Hospital, Amsterdam Neuroscience, Amsterdam UMC, De Boelelaan 1117, 1081 Amsterdam, the Netherlands
| | - Martina Hruzova
- Department of Biology, Institute for Molecular Health Sciences, ETH Zurich, Otto-Stern-Weg 7, 8093 Zurich, Switzerland.,Institute of Pharmacology and Toxicology, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Tanja Rothgangl
- Institute of Pharmacology and Toxicology, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Marjolein Breur
- Department of Pediatrics and Child Neurology, Emma Children's Hospital, Amsterdam Neuroscience, Amsterdam UMC, De Boelelaan 1117, 1081 Amsterdam, the Netherlands
| | - Marianna Bugiani
- Department of Pediatrics and Child Neurology, Emma Children's Hospital, Amsterdam Neuroscience, Amsterdam UMC, De Boelelaan 1117, 1081 Amsterdam, the Netherlands
| | - Marjo S van der Knaap
- Department of Pediatrics and Child Neurology, Emma Children's Hospital, Amsterdam Neuroscience, Amsterdam UMC, De Boelelaan 1117, 1081 Amsterdam, the Netherlands.,Department of Functional Genomics, Center for Neurogenomics and Cognitive Research, Amsterdam Neuroscience, VU University, De Boelelaan 1085, 1081 Amsterdam, the Netherlands
| | - Gerald Schwank
- Department of Biology, Institute for Molecular Health Sciences, ETH Zurich, Otto-Stern-Weg 7, 8093 Zurich, Switzerland.,Institute of Pharmacology and Toxicology, University of Zurich, Winterthurerstrasse 190, 8057 Zurich, Switzerland
| | - Vivi M Heine
- Department of Complex Trait Genetics, Center for Neurogenomics and Cognitive Research, Amsterdam Neuroscience, VU University, De Boelelaan 1085, 1081 Amsterdam, the Netherlands.,Department of Child and Adolescence Psychiatry, Emma Children's Hospital, Amsterdam Neuroscience, Amsterdam UMC, De Boelelaan 1085, 1081 Amsterdam, the Netherlands
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19
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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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20
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Ou-Yang H, Yang SH, Chen W, Yang SH, Cidem A, Sung LY, Chen CM. Cruciform DNA Structures Act as Legible Templates for Accelerating Homologous Recombination in Transgenic Animals. Int J Mol Sci 2022; 23:3973. [PMID: 35409332 PMCID: PMC9000021 DOI: 10.3390/ijms23073973] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/26/2022] [Revised: 03/27/2022] [Accepted: 03/29/2022] [Indexed: 11/16/2022] Open
Abstract
Inverted repeat (IR) DNA sequences compose cruciform structures. Some genetic disorders are the result of genome inversion or translocation by cruciform DNA structures. The present study examined whether exogenous DNA integration into the chromosomes of transgenic animals was related to cruciform DNA structures. Large imperfect cruciform structures were frequently predicted around predestinated transgene integration sites in host genomes of microinjection-based transgenic (Tg) animals (αLA-LPH Tg goat, Akr1A1eGFP/eGFP Tg mouse, and NFκB-Luc Tg mouse) or CRISPR/Cas9 gene-editing (GE) animals (αLA-AP1 GE mouse). Transgene cassettes were imperfectly matched with their predestinated sequences. According to the analyzed data, we proposed a putative model in which the flexible cruciform DNA structures acted as a legible template for DNA integration into linear DNAs or double-strand break (DSB) alleles. To demonstrate this model, artificial inverted repeat knock-in (KI) reporter plasmids were created to analyze the KI rate using the CRISPR/Cas9 system in NIH3T3 cells. Notably, the KI rate of the 5′ homologous arm inverted repeat donor plasmid (5′IR) with the ROSA gRNA group (31.5%) was significantly higher than the knock-in reporter donor plasmid (KIR) with the ROSA gRNA group (21.3%, p < 0.05). However, the KI rate of the 3′ inverted terminal repeat/inverted repeat donor plasmid (3′ITRIR) group was not different from the KIR group (23.0% vs. 22.0%). These results demonstrated that the legibility of the sequence with the cruciform DNA existing in the transgene promoted homologous recombination (HR) with a higher KI rate. Our findings suggest that flexible cruciform DNAs folded by IR sequences improve the legibility and accelerate DNA 3′-overhang integration into the host genome via homologous recombination machinery.
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Affiliation(s)
- Huan Ou-Yang
- Program in Translational Medicine, Department of Life Sciences, National Chung Hsing University, Taichung 402, Taiwan; (H.O.-Y.); (S.-H.Y.); (A.C.)
- Institute of Biotechnology, College of Bioresources and Agriculture, National Taiwan University, Taipei 106, Taiwan
| | - Shiao-Hsuan Yang
- Program in Translational Medicine, Department of Life Sciences, National Chung Hsing University, Taichung 402, Taiwan; (H.O.-Y.); (S.-H.Y.); (A.C.)
- Reproductive Medicine Center, Department of Gynecology, Changhua Christian Hospital, Changhua 515, Taiwan
| | - Wei Chen
- Division of Pulmonary and Critical Care Medicine, Chia-Yi Christian Hospital, Chiayi 600, Taiwan;
| | - Shang-Hsun Yang
- Department of Physiology, National Cheng Kung University, Tainan 701, Taiwan;
- Institute of Basic Medical Sciences, College of Medicine, National Cheng Kung University, Tainan 701, Taiwan
| | - Abdulkadir Cidem
- Program in Translational Medicine, Department of Life Sciences, National Chung Hsing University, Taichung 402, Taiwan; (H.O.-Y.); (S.-H.Y.); (A.C.)
- Department of Molecular Biology and Genetics, Erzurum Technical University, Erzurum 25250, Turkey
| | - Li-Ying Sung
- Institute of Biotechnology, College of Bioresources and Agriculture, National Taiwan University, Taipei 106, Taiwan
| | - Chuan-Mu Chen
- Program in Translational Medicine, Department of Life Sciences, National Chung Hsing University, Taichung 402, Taiwan; (H.O.-Y.); (S.-H.Y.); (A.C.)
- The iEGG and Animal Biotechnology Center, National Chung Hsing University, Taichung 402, Taiwan
- Rong-Hsing Translational Medicine Research Center, Taichung Veterans General Hospital, Taichung 407, Taiwan
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21
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Okada K, Aoki K, Tabei T, Sugio K, Imai K, Bonkohara Y, Kamachi Y. Key sequence features of CRISPR RNA for dual-guide CRISPR-Cas9 ribonucleoprotein complexes assembled with wild-type or HiFi Cas9. Nucleic Acids Res 2022; 50:2854-2871. [PMID: 35166844 PMCID: PMC8934663 DOI: 10.1093/nar/gkac100] [Citation(s) in RCA: 4] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/10/2021] [Revised: 01/28/2022] [Accepted: 02/02/2022] [Indexed: 11/22/2022] Open
Abstract
Specific sequence features of the protospacer and protospacer-adjacent motif (PAM) are critical for efficient cleavage by CRISPR-Cas9, but current knowledge is largely derived from single-guide RNA (sgRNA) systems assessed in cultured cells. In this study, we sought to determine gRNA sequence features of a more native CRISPR-Cas9 ribonucleoprotein (RNP) complex with dual-guide RNAs (dgRNAs) composed of crRNA and tracrRNA, which has been used increasingly in recent CRISPR-Cas9 applications, particularly in zebrafish. Using both wild-type and HiFi SpCas9, we determined on-target cleavage efficiencies of 51 crRNAs in zebrafish embryos by assessing indel occurrence. Statistical analysis of these data identified novel position-specific mononucleotide features relevant to cleavage efficiencies throughout the protospacer sequence that may be unique to CRISPR-Cas9 RNPs pre-assembled with perfectly matched gRNAs. Overall features for wild-type Cas9 resembled those for HiFi Cas9, but specific differences were also observed. Mutational analysis of mononucleotide features confirmed their relevance to cleavage efficiencies. Moreover, the mononucleotide feature-based score, CRISPR-kp, correlated well with efficiencies of gRNAs reported in previous zebrafish RNP injection experiments, as well as independently tested crRNAs only in RNP format, but not with Cas9 mRNA co-injection. These findings will facilitate design of gRNA/crRNAs in genome editing applications, especially when using pre-assembled RNPs.
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Affiliation(s)
- Keita Okada
- School of Environmental Science and Engineering, Kochi University of Technology, Kami, Kochi 782-8502, Japan
| | - Kanae Aoki
- School of Environmental Science and Engineering, Kochi University of Technology, Kami, Kochi 782-8502, Japan
| | - Teruyuki Tabei
- School of Environmental Science and Engineering, Kochi University of Technology, Kami, Kochi 782-8502, Japan
| | - Kota Sugio
- School of Environmental Science and Engineering, Kochi University of Technology, Kami, Kochi 782-8502, Japan
| | - Katsunori Imai
- School of Environmental Science and Engineering, Kochi University of Technology, Kami, Kochi 782-8502, Japan
| | - Yuki Bonkohara
- School of Environmental Science and Engineering, Kochi University of Technology, Kami, Kochi 782-8502, Japan
| | - Yusuke Kamachi
- School of Environmental Science and Engineering, Kochi University of Technology, Kami, Kochi 782-8502, Japan
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22
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Yan Y, Wang Q. BMP Signaling: Lighting up the Way for Embryonic Dorsoventral Patterning. Front Cell Dev Biol 2022; 9:799772. [PMID: 35036406 PMCID: PMC8753366 DOI: 10.3389/fcell.2021.799772] [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: 10/22/2021] [Accepted: 12/06/2021] [Indexed: 11/13/2022] Open
Abstract
One of the most significant events during early embryonic development is the establishment of a basic embryonic body plan, which is defined by anteroposterior, dorsoventral (DV), and left-right axes. It is well-known that the morphogen gradient created by BMP signaling activity is crucial for DV axis patterning across a diverse set of vertebrates. The regulation of BMP signaling during DV patterning has been strongly conserved across evolution. This is a remarkable regulatory and evolutionary feat, as the BMP gradient has been maintained despite the tremendous variation in embryonic size and shape across species. Interestingly, the embryonic DV axis exhibits robust stability, even in face of variations in BMP signaling. Multiple lines of genetic, molecular, and embryological evidence have suggested that numerous BMP signaling components and their attendant regulators act in concert to shape the developing DV axis. In this review, we summarize the current knowledge of the function and regulation of BMP signaling in DV patterning. Throughout, we focus specifically on popular model animals, such as Xenopus and zebrafish, highlighting the similarities and differences of the regulatory networks between species. We also review recent advances regarding the molecular nature of DV patterning, including the initiation of the DV axis, the formation of the BMP gradient, and the regulatory molecular mechanisms behind BMP signaling during the establishment of the DV axis. Collectively, this review will help clarify our current understanding of the molecular nature of DV axis formation.
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Affiliation(s)
- Yifang Yan
- Center for Reproductive Medicine, Department of Obstetrics and Gynecology, Peking University Third Hospital, Beijing, China.,National Clinical Research Center for Obstetrics and Gynecology (Peking University Third Hospital), Beijing, China.,Key Laboratory of Assisted Reproduction (Peking University), Ministry of Education, Beijing, China.,Beijing Key Laboratory of Reproductive Endocrinology and Assisted Reproductive Technology, Beijing, China
| | - Qiang Wang
- State Key Laboratory of Membrane Biology, CAS Center for Excellence in Molecular Cell Science, Institute of Zoology, University of Chinese Academy of Sciences, Chinese Academy of Sciences, Beijing, China.,Institute for Stem Cell and Regeneration, Chinese Academy of Sciences, Beijing, China
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23
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Ittiprasert W, Chatupheeraphat C, Mann VH, Li W, Miller A, Ogunbayo T, Tran K, Alrefaei YN, Mentink-Kane M, Brindley PJ. RNA-Guided AsCas12a- and SpCas9-Catalyzed Knockout and Homology Directed Repair of the Omega-1 Locus of the Human Blood Fluke, Schistosoma mansoni. Int J Mol Sci 2022; 23:631. [PMID: 35054816 PMCID: PMC8775552 DOI: 10.3390/ijms23020631] [Citation(s) in RCA: 8] [Impact Index Per Article: 4.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/07/2021] [Revised: 12/30/2021] [Accepted: 01/01/2022] [Indexed: 12/17/2022] Open
Abstract
The efficiency of the RNA-guided AsCas12a nuclease of Acidaminococcus sp. was compared with SpCas9 from Streptococcus pyogenes, for functional genomics in Schistosoma mansoni. We deployed optimized conditions for the ratio of guide RNAs to the nuclease, donor templates, and electroporation parameters, to target a key schistosome enzyme termed omega-1. Programmed cleavages catalyzed by Cas12a and Cas9 resulted in staggered- and blunt-ended strand breaks, respectively. AsCas12a was more efficient than SpCas9 for gene knockout, as determined by TIDE analysis. CRISPResso2 analysis confirmed that most mutations were deletions. Knockout efficiency of both nucleases markedly increased in the presence of single-stranded oligodeoxynucleotide (ssODN) template. With AsCas12a, ssODNs representative of both the non-CRISPR target (NT) and target (T) strands were tested, resulting in KO efficiencies of 15.67, 28.71, and 21.43% in the SpCas9 plus ssODN, AsCas12a plus NT-ssODN, and AsCas12a plus T-ssODN groups, respectively. Trans-cleavage against the ssODNs by activated AsCas12a was not apparent in vitro. SpCas9 catalyzed more precise transgene insertion, with knock-in efficiencies of 17.07% for the KI_Cas9 group, 14.58% for KI_Cas12a-NT-ssODN, and 12.37% for KI_Cas12a-T-ssODN. Although AsCas12a induced fewer mutations per genome than SpCas9, the phenotypic impact on transcription and expression of omega-1 was similar for both nucleases.
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Affiliation(s)
- Wannaporn Ittiprasert
- Department of Microbiology, Immunology & Tropical Medicine, & Research Center for Neglected Diseases of Poverty, School of Medicine & Health Sciences, George Washington University, Washington, DC 20037, USA; (C.C.); (V.H.M.); (W.L.); (Y.N.A.)
| | - Chawalit Chatupheeraphat
- Department of Microbiology, Immunology & Tropical Medicine, & Research Center for Neglected Diseases of Poverty, School of Medicine & Health Sciences, George Washington University, Washington, DC 20037, USA; (C.C.); (V.H.M.); (W.L.); (Y.N.A.)
- Center for Research and Innovation, Faculty of Medical Technology, Mahidol University, Salaya, Nakhon Pathom 73170, Thailand
| | - Victoria H. Mann
- Department of Microbiology, Immunology & Tropical Medicine, & Research Center for Neglected Diseases of Poverty, School of Medicine & Health Sciences, George Washington University, Washington, DC 20037, USA; (C.C.); (V.H.M.); (W.L.); (Y.N.A.)
| | - Wenhui Li
- Department of Microbiology, Immunology & Tropical Medicine, & Research Center for Neglected Diseases of Poverty, School of Medicine & Health Sciences, George Washington University, Washington, DC 20037, USA; (C.C.); (V.H.M.); (W.L.); (Y.N.A.)
- Lanzhou Veterinary Research Institute, Chinese Academy of Agricultural Sciences, Lanzhou 730050, China
| | - André Miller
- Schistosomiasis Resource Center, Biomedical Research Institute, Rockville, MD 20850, USA; (A.M.); (T.O.); (K.T.); (M.M.-K.)
| | - Taiwo Ogunbayo
- Schistosomiasis Resource Center, Biomedical Research Institute, Rockville, MD 20850, USA; (A.M.); (T.O.); (K.T.); (M.M.-K.)
| | - Kenny Tran
- Schistosomiasis Resource Center, Biomedical Research Institute, Rockville, MD 20850, USA; (A.M.); (T.O.); (K.T.); (M.M.-K.)
| | - Yousef N. Alrefaei
- Department of Microbiology, Immunology & Tropical Medicine, & Research Center for Neglected Diseases of Poverty, School of Medicine & Health Sciences, George Washington University, Washington, DC 20037, USA; (C.C.); (V.H.M.); (W.L.); (Y.N.A.)
- Department of Medical Laboratory Technology, College of Health Sciences, PAEET, Adailiya, Kuwait City 73101, Kuwait
| | - Margaret Mentink-Kane
- Schistosomiasis Resource Center, Biomedical Research Institute, Rockville, MD 20850, USA; (A.M.); (T.O.); (K.T.); (M.M.-K.)
| | - Paul J. Brindley
- Department of Microbiology, Immunology & Tropical Medicine, & Research Center for Neglected Diseases of Poverty, School of Medicine & Health Sciences, George Washington University, Washington, DC 20037, USA; (C.C.); (V.H.M.); (W.L.); (Y.N.A.)
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Levic DS, Yamaguchi N, Wang S, Knaut H, Bagnat M. Knock-in tagging in zebrafish facilitated by insertion into non-coding regions. Development 2021; 148:dev199994. [PMID: 34495314 PMCID: PMC8513609 DOI: 10.1242/dev.199994] [Citation(s) in RCA: 17] [Impact Index Per Article: 5.7] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/08/2021] [Accepted: 08/27/2021] [Indexed: 11/20/2022]
Abstract
Zebrafish provide an excellent model for in vivo cell biology studies because of their amenability to live imaging. Protein visualization in zebrafish has traditionally relied on overexpression of fluorescently tagged proteins from heterologous promoters, making it difficult to recapitulate endogenous expression patterns and protein function. One way to circumvent this problem is to tag the proteins by modifying their endogenous genomic loci. Such an approach is not widely available to zebrafish researchers because of inefficient homologous recombination and the error-prone nature of targeted integration in zebrafish. Here, we report a simple approach for tagging proteins in zebrafish on their N or C termini with fluorescent proteins by inserting PCR-generated donor amplicons into non-coding regions of the corresponding genes. Using this approach, we generated endogenously tagged alleles for several genes that are crucial for epithelial biology and organ development, including the tight junction components ZO-1 and Cldn15la, the trafficking effector Rab11a, the apical polarity protein aPKC and the ECM receptor Integrin β1b. Our approach facilitates the generation of knock-in lines in zebrafish, opening the way for accurate quantitative imaging studies.
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Affiliation(s)
- Daniel S. Levic
- Department of Cell Biology, Duke University, Durham, NC 27710, USA
| | - Naoya Yamaguchi
- Skirball Institute of Biomolecular Medicine, New York University Grossman School of Medicine, Department of Cell Biology, New York, NY 10016, USA
| | - Siyao Wang
- Department of Cell Biology, Duke University, Durham, NC 27710, USA
| | - Holger Knaut
- Skirball Institute of Biomolecular Medicine, New York University Grossman School of Medicine, Department of Cell Biology, New York, NY 10016, USA
| | - Michel Bagnat
- Department of Cell Biology, Duke University, Durham, NC 27710, USA
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