301
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Prykhozhij SV, Steele SL, Razaghi B, Berman JN. A rapid and effective method for screening, sequencing and reporter verification of engineered frameshift mutations in zebrafish. Dis Model Mech 2017; 10:811-822. [PMID: 28280001 PMCID: PMC5483001 DOI: 10.1242/dmm.026765] [Citation(s) in RCA: 40] [Impact Index Per Article: 5.7] [Reference Citation Analysis] [Abstract] [Key Words] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/28/2016] [Accepted: 03/03/2017] [Indexed: 12/30/2022] Open
Abstract
Clustered regularly interspaced palindromic repeats (CRISPR)/Cas-based adaptive immunity against pathogens in bacteria has been adapted for genome editing and applied in zebrafish (Danio rerio) to generate frameshift mutations in protein-coding genes. Although there are methods to detect, quantify and sequence CRISPR/Cas9-induced mutations, identifying mutations in F1 heterozygous fish remains challenging. Additionally, sequencing a mutation and assuming that it causes a frameshift does not prove causality because of possible alternative translation start sites and potential effects of mutations on splicing. This problem is compounded by the relatively few antibodies available for zebrafish proteins, limiting validation at the protein level. To address these issues, we developed a detailed protocol to screen F1 mutation carriers, and clone and sequence identified mutations. In order to verify that mutations actually cause frameshifts, we created a fluorescent reporter system that can detect frameshift efficiency based on the cloning of wild-type and mutant cDNA fragments and their expression levels. As proof of principle, we applied this strategy to three CRISPR/Cas9-induced mutations in pycr1a, chd7 and hace1 genes. An insertion of seven nucleotides in pycr1a resulted in the first reported observation of exon skipping by CRISPR/Cas9-induced mutations in zebrafish. However, of these three mutant genes, the fluorescent reporter revealed effective frameshifting exclusively in the case of a two-nucleotide deletion in chd7, suggesting activity of alternative translation sites in the other two mutants even though pycr1a exon-skipping deletion is likely to be deleterious. This article provides a protocol for characterizing frameshift mutations in zebrafish, and highlights the importance of checking mutations at the mRNA level and verifying their effects on translation by fluorescent reporters when antibody detection of protein loss is not possible.
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Affiliation(s)
| | - Shelby L Steele
- Department of Pediatrics, Dalhousie University, Halifax, NS, Canada B3K 6R8
| | - Babak Razaghi
- Department of Pediatrics, Dalhousie University, Halifax, NS, Canada B3K 6R8
| | - Jason N Berman
- Department of Pediatrics, Dalhousie University, Halifax, NS, Canada B3K 6R8 .,Department of Microbiology and Immunology, Dalhousie University, Halifax, NS, Canada B3H 4R2.,Department of Pathology, Dalhousie University, Halifax, NS, Canada B3H4R2
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302
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Jiang T, Kindt K, Wu DK. Transcription factor Emx2 controls stereociliary bundle orientation of sensory hair cells. eLife 2017; 6. [PMID: 28266911 PMCID: PMC5388538 DOI: 10.7554/elife.23661] [Citation(s) in RCA: 70] [Impact Index Per Article: 10.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/26/2016] [Accepted: 03/04/2017] [Indexed: 12/13/2022] Open
Abstract
The asymmetric location of stereociliary bundle (hair bundle) on the apical surface of mechanosensory hair cells (HCs) dictates the direction in which a given HC can respond to cues such as sound, head movements, and water pressure. Notably, vestibular sensory organs of the inner ear, the maculae, exhibit a line of polarity reversal (LPR) across which, hair bundles are polarized in a mirror-image pattern. Similarly, HCs in neuromasts of the zebrafish lateral line system are generated as pairs, and two sibling HCs develop opposite hair bundle orientations. Within these sensory organs, expression of the transcription factor Emx2 is restricted to only one side of the LPR in the maculae or one of the two sibling HCs in neuromasts. Emx2 mediates hair bundle polarity reversal in these restricted subsets of HCs and generates the mirror-image pattern of the sensory organs. Downstream effectors of Emx2 control bundle polarity cell-autonomously via heterotrimeric G proteins.
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Affiliation(s)
- Tao Jiang
- Program in Neuroscience and Cognitive Science, University of Maryland, College Park, United States.,Laboratory of Molecular Biology, National Institute on Deafness and Other Communication Disorders, National Institutes of Health, Bethesda, United States
| | - Katie Kindt
- Section on Sensory Cell Development, National Institute on Deafness and Other Communication Disorders, National Institutes of Health, Bethesda, United States
| | - Doris K Wu
- Laboratory of Molecular Biology, National Institute on Deafness and Other Communication Disorders, National Institutes of Health, Bethesda, United States
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303
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High-resolution 3D imaging of whole organ after clearing: taking a new look at the zebrafish testis. Sci Rep 2017; 7:43012. [PMID: 28211501 PMCID: PMC5314416 DOI: 10.1038/srep43012] [Citation(s) in RCA: 26] [Impact Index Per Article: 3.7] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/18/2016] [Accepted: 01/17/2017] [Indexed: 12/17/2022] Open
Abstract
Zebrafish testis has become a powerful model for reproductive biology of teleostean fishes and other vertebrates and encompasses multiple applications in applied and basic research. Many studies have focused on 2D images, which is time consuming and implies extrapolation of results. Three-dimensional imaging of whole organs recently became an important challenge to better understand their architecture and allow cell enumeration. Several protocols have thus been developed to enhance sample transparency, a limiting step for imaging large biological samples. However, none of these methods has been applied to the zebrafish testis. We tested five clearing protocols to determine if some of them could be applied with only small modifications to the testis. We compared clearing efficiency at both macroscopic and microscopic levels. CUBIC and PACT were suitable for an efficient transparency, an optimal optical penetration, the GFP fluorescence preservation and avoiding meaningful tissue deformation. Finally, we succeeded in whole testis 3D capture at a cellular resolution with both CUBIC and PACT, which will be valuable in a standard workflow to investigate the 3D architecture of the testis and its cellular content. This paves the way for further development of high content phenotyping studies in several fields including development, genetic or toxicology.
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304
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Watkins-Chow DE, Varshney GK, Garrett LJ, Chen Z, Jimenez EA, Rivas C, Bishop KS, Sood R, Harper UL, Pavan WJ, Burgess SM. Highly Efficient Cpf1-Mediated Gene Targeting in Mice Following High Concentration Pronuclear Injection. G3 (BETHESDA, MD.) 2017; 7:719-722. [PMID: 28040780 PMCID: PMC5295614 DOI: 10.1534/g3.116.038091] [Citation(s) in RCA: 21] [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: 09/28/2016] [Accepted: 12/20/2016] [Indexed: 01/12/2023]
Abstract
Cpf1 has emerged as an alternative to the Cas9 RNA-guided nuclease. Here we show that gene targeting rates in mice using Cpf1 can meet, or even surpass, Cas9 targeting rates (approaching 100% targeting), but require higher concentrations of mRNA and guide. We also demonstrate that coinjecting two guides with close targeting sites can result in synergistic genomic cutting, even if one of the guides has minimal cutting activity.
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Affiliation(s)
- Dawn E Watkins-Chow
- Genetic Disease Research Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, Maryland 20892
| | - Gaurav K Varshney
- Functional and Chemical Genomics Program, Oklahoma Medical Research Foundation, Oklahoma City, Oklahoma 73104
| | - Lisa J Garrett
- Embryonic Stem Cell and Transgenic Mouse Core, National Human Genome Research Institute, National Institutes of Health, Bethesda, Maryland 20892
| | - Zelin Chen
- Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, Maryland 20892
| | - Erin A Jimenez
- Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, Maryland 20892
| | - Cecilia Rivas
- Embryonic Stem Cell and Transgenic Mouse Core, National Human Genome Research Institute, National Institutes of Health, Bethesda, Maryland 20892
| | - Kevin S Bishop
- Zebrafish Core, Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, Maryland 20892
| | - Raman Sood
- Zebrafish Core, Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, Maryland 20892
| | - Ursula L Harper
- Genomics Core, National Human Genome Research Institute, National Institutes of Health, Bethesda, Maryland 20892
| | - William J Pavan
- Genetic Disease Research Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, Maryland 20892
| | - Shawn M Burgess
- Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, Maryland 20892
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305
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Horstick EJ, Bayleyen Y, Sinclair JL, Burgess HA. Search strategy is regulated by somatostatin signaling and deep brain photoreceptors in zebrafish. BMC Biol 2017; 15:4. [PMID: 28122559 PMCID: PMC5267475 DOI: 10.1186/s12915-016-0346-2] [Citation(s) in RCA: 31] [Impact Index Per Article: 4.4] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/13/2016] [Accepted: 12/22/2016] [Indexed: 11/10/2022] Open
Abstract
BACKGROUND Animals use sensory cues to efficiently locate resources, but when sensory information is insufficient, they may rely on internally coded search strategies. Despite the importance of search behavior, there is limited understanding of the underlying neural mechanisms in vertebrates. RESULTS Here, we report that loss of illumination initiates sophisticated light-search behavior in larval zebrafish. Using three-dimensional tracking, we show that at the onset of darkness larvae swim in a helical trajectory that is spatially restricted in the horizontal plane, before gradually transitioning to an outward movement profile. Local and outward swim patterns display characteristic features of area-restricted and roaming search strategies, differentially enhancing phototaxis to nearby and remote sources of light. Retinal signaling is only required to initiate area-restricted search, implying that photoreceptors within the brain drive the transition to the roaming search state. Supporting this, orthopediaA mutant larvae manifest impaired transition to roaming search, a phenotype which is recapitulated by loss of the non-visual opsin opn4a and somatostatin signaling. CONCLUSION These findings define distinct neuronal pathways for area-restricted and roaming search behaviors and clarify how internal drives promote goal-directed activity.
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Affiliation(s)
- Eric J Horstick
- Division of Developmental Biology, Eunice Kennedy Shriver National Institute of Child Health and Human Development, Bethesda, MD, 20892, USA.
| | - Yared Bayleyen
- Division of Developmental Biology, Eunice Kennedy Shriver National Institute of Child Health and Human Development, Bethesda, MD, 20892, USA
| | - Jennifer L Sinclair
- Division of Developmental Biology, Eunice Kennedy Shriver National Institute of Child Health and Human Development, Bethesda, MD, 20892, USA
| | - Harold A Burgess
- Division of Developmental Biology, Eunice Kennedy Shriver National Institute of Child Health and Human Development, Bethesda, MD, 20892, USA.
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306
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Capon SJ, Baillie GJ, Bower NI, da Silva JA, Paterson S, Hogan BM, Simons C, Smith KA. Utilising polymorphisms to achieve allele-specific genome editing in zebrafish. Biol Open 2017; 6:125-131. [PMID: 27895053 PMCID: PMC5278422 DOI: 10.1242/bio.020974] [Citation(s) in RCA: 15] [Impact Index Per Article: 2.1] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/26/2022] Open
Abstract
The advent of genome editing has significantly altered genetic research, including research using the zebrafish model. To better understand the selectivity of the commonly used CRISPR/Cas9 system, we investigated single base pair mismatches in target sites and examined how they affect genome editing in the zebrafish model. Using two different zebrafish strains that have been deep sequenced, CRISPR/Cas9 target sites containing polymorphisms between the two strains were identified. These strains were crossed (creating heterozygotes at polymorphic sites) and CRISPR/Cas9 complexes that perfectly complement one strain injected. Sequencing of targeted sites showed biased, allele-specific editing for the perfectly complementary sequence in the majority of cases (14/19). To test utility, we examined whether phenotypes generated by F0 injection could be internally controlled with such polymorphisms. Targeting of genes bmp7a and chordin showed reduction in the frequency of phenotypes in injected ‘heterozygotes’ compared with injecting the strain with perfect complementarity. Next, injecting CRISPR/Cas9 complexes targeting two separate sites created deletions, but deletions were biased to selected chromosomes when one CRISPR/Cas9 target contained a polymorphism. Finally, integration of loxP sequences occurred preferentially in alleles with perfect complementarity. These experiments demonstrate that single nucleotide polymorphisms (SNPs) present throughout the genome can be utilised to increase the efficiency of in cis genome editing using CRISPR/Cas9 in the zebrafish model. Summary: Heterozygous single nucleotide polymorphisms in CRISPR/Cas9 target sites bias genome editing in favour of alleles with perfect complementarity to gRNAs, a feature which can be exploited for chromosome-specific editing.
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Affiliation(s)
- Samuel J Capon
- Division of Genomics of Development and Disease, Institute for Molecular Bioscience, The University of Queensland, Brisbane, Queensland 4072, Australia
| | - Gregory J Baillie
- Division of Genomics of Development and Disease, Institute for Molecular Bioscience, The University of Queensland, Brisbane, Queensland 4072, Australia
| | - Neil I Bower
- Division of Genomics of Development and Disease, Institute for Molecular Bioscience, The University of Queensland, Brisbane, Queensland 4072, Australia
| | - Jason A da Silva
- Division of Genomics of Development and Disease, Institute for Molecular Bioscience, The University of Queensland, Brisbane, Queensland 4072, Australia
| | - Scott Paterson
- Division of Genomics of Development and Disease, Institute for Molecular Bioscience, The University of Queensland, Brisbane, Queensland 4072, Australia
| | - Benjamin M Hogan
- Division of Genomics of Development and Disease, Institute for Molecular Bioscience, The University of Queensland, Brisbane, Queensland 4072, Australia
| | - Cas Simons
- Division of Genomics of Development and Disease, Institute for Molecular Bioscience, The University of Queensland, Brisbane, Queensland 4072, Australia
| | - Kelly A Smith
- Division of Genomics of Development and Disease, Institute for Molecular Bioscience, The University of Queensland, Brisbane, Queensland 4072, Australia
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307
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Delker RK, Mann RS. From Reductionism to Holism: Toward a More Complete View of Development Through Genome Engineering. ADVANCES IN EXPERIMENTAL MEDICINE AND BIOLOGY 2017; 1016:45-74. [PMID: 29130153 PMCID: PMC6935049 DOI: 10.1007/978-3-319-63904-8_3] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 12/22/2022]
Abstract
Paradigm shifts in science are often coupled to technological advances. New techniques offer new roads of discovery; but, more than this, they shape the way scientists approach questions. Developmental biology exemplifies this idea both in its past and present. The rise of molecular biology and genetics in the late twentieth century shifted the focus from the anatomical to the molecular, nudging the underlying philosophy from holism to reductionism. Developmental biology is currently experiencing yet another transformation triggered by '-omics' technology and propelled forward by CRISPR genome engineering (GE). Together, these technologies are helping to reawaken a holistic approach to development. Herein, we focus on CRISPR GE and its potential to reveal principles of development at the level of the genome, the epigenome, and the cell. Within each stage we illustrate how GE can move past pure reductionism and embrace holism, ultimately delivering a more complete view of development.
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Affiliation(s)
- Rebecca K Delker
- Department of Biochemistry and Molecular Biophysics and Systems Biology, Mortimer B. Zuckerman Mind Brain Behavior Institute, Columbia University, 612 West 130th Street, 9th Floor, New York, NY, 10027, USA.
| | - Richard S Mann
- Department of Biochemistry and Molecular Biophysics and Systems Biology, Mortimer B. Zuckerman Mind Brain Behavior Institute, Columbia University, 612 West 130th Street, 9th Floor, New York, NY, 10027, USA
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308
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Noyes PD, Garcia GR, Tanguay RL. ZEBRAFISH AS AN IN VIVO MODEL FOR SUSTAINABLE CHEMICAL DESIGN. GREEN CHEMISTRY : AN INTERNATIONAL JOURNAL AND GREEN CHEMISTRY RESOURCE : GC 2016; 18:6410-6430. [PMID: 28461781 PMCID: PMC5408959 DOI: 10.1039/c6gc02061e] [Citation(s) in RCA: 7] [Impact Index Per Article: 0.9] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 05/11/2023]
Abstract
Heightened public awareness about the many thousands of chemicals in use and present as persistent contaminants in the environment has increased the demand for safer chemicals and more rigorous toxicity testing. There is a growing recognition that the use of traditional test models and empirical approaches is impractical for screening for toxicity the many thousands of chemicals in the environment and the hundreds of new chemistries introduced each year. These realities coupled with the green chemistry movement have prompted efforts to implement more predictive-based approaches to evaluate chemical toxicity early in product development. While used for many years in environmental toxicology and biomedicine, zebrafish use has accelerated more recently in genetic toxicology, high throughput screening (HTS), and behavioral testing. This review describes major advances in these testing methods that have positioned the zebrafish as a highly applicable model in chemical safety evaluations and sustainable chemistry efforts. Many toxic responses have been shown to be shared among fish and mammals owing to their generally well-conserved development, cellular networks, and organ systems. These shared responses have been observed for chemicals that impair endocrine functioning, development, and reproduction, as well as those that elicit cardiotoxicity and carcinogenicity, among other diseases. HTS technologies with zebrafish enable screening large chemical libraries for bioactivity that provide opportunities for testing early in product development. A compelling attribute of the zebrafish centers on being able to characterize toxicity mechanisms across multiple levels of biological organization from the genome to receptor interactions and cellular processes leading to phenotypic changes such as developmental malformations. Finally, there is a growing recognition of the links between human and wildlife health and the need for approaches that allow for assessment of real world multi-chemical exposures. The zebrafish is poised to be an important model in bridging these two conventionally separate areas of toxicology and characterizing the biological effects of chemical mixtures that could augment its role in sustainable chemistry.
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Affiliation(s)
- Pamela D. Noyes
- Department of Environmental & Molecular Toxicology, Oregon State University, Corvallis, OR 97331
| | - Gloria R. Garcia
- Department of Environmental & Molecular Toxicology, Oregon State University, Corvallis, OR 97331
| | - Robert L. Tanguay
- Department of Environmental & Molecular Toxicology, Oregon State University, Corvallis, OR 97331
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309
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Bortesi L, Zhu C, Zischewski J, Perez L, Bassié L, Nadi R, Forni G, Lade SB, Soto E, Jin X, Medina V, Villorbina G, Muñoz P, Farré G, Fischer R, Twyman RM, Capell T, Christou P, Schillberg S. Patterns of CRISPR/Cas9 activity in plants, animals and microbes. PLANT BIOTECHNOLOGY JOURNAL 2016; 14:2203-2216. [PMID: 27614091 PMCID: PMC5103219 DOI: 10.1111/pbi.12634] [Citation(s) in RCA: 89] [Impact Index Per Article: 11.1] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Received: 07/11/2016] [Revised: 09/05/2016] [Accepted: 09/07/2016] [Indexed: 05/19/2023]
Abstract
The CRISPR/Cas9 system and related RNA-guided endonucleases can introduce double-strand breaks (DSBs) at specific sites in the genome, allowing the generation of targeted mutations in one or more genes as well as more complex genomic rearrangements. Modifications of the canonical CRISPR/Cas9 system from Streptococcus pyogenes and the introduction of related systems from other bacteria have increased the diversity of genomic sites that can be targeted, providing greater control over the resolution of DSBs, the targeting efficiency (frequency of on-target mutations), the targeting accuracy (likelihood of off-target mutations) and the type of mutations that are induced. Although much is now known about the principles of CRISPR/Cas9 genome editing, the likelihood of different outcomes is species-dependent and there have been few comparative studies looking at the basis of such diversity. Here we critically analyse the activity of CRISPR/Cas9 and related systems in different plant species and compare the outcomes in animals and microbes to draw broad conclusions about the design principles required for effective genome editing in different organisms. These principles will be important for the commercial development of crops, farm animals, animal disease models and novel microbial strains using CRISPR/Cas9 and other genome-editing tools.
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Affiliation(s)
- Luisa Bortesi
- Institute for Molecular BiotechnologyRWTH Aachen UniversityAachenGermany
| | - Changfu Zhu
- Department of Plant Production and Forestry ScienceSchool of Agrifood and Forestry Science and Engineering (ETSEA)University of Lleida‐Agrotecnio CenterLleidaSpain
| | - Julia Zischewski
- Institute for Molecular BiotechnologyRWTH Aachen UniversityAachenGermany
| | - Lucia Perez
- Department of Plant Production and Forestry ScienceSchool of Agrifood and Forestry Science and Engineering (ETSEA)University of Lleida‐Agrotecnio CenterLleidaSpain
| | - Ludovic Bassié
- Department of Plant Production and Forestry ScienceSchool of Agrifood and Forestry Science and Engineering (ETSEA)University of Lleida‐Agrotecnio CenterLleidaSpain
| | - Riad Nadi
- Department of Plant Production and Forestry ScienceSchool of Agrifood and Forestry Science and Engineering (ETSEA)University of Lleida‐Agrotecnio CenterLleidaSpain
| | - Giobbe Forni
- Department of Plant Production and Forestry ScienceSchool of Agrifood and Forestry Science and Engineering (ETSEA)University of Lleida‐Agrotecnio CenterLleidaSpain
| | - Sarah Boyd Lade
- Department of Plant Production and Forestry ScienceSchool of Agrifood and Forestry Science and Engineering (ETSEA)University of Lleida‐Agrotecnio CenterLleidaSpain
| | - Erika Soto
- Department of Plant Production and Forestry ScienceSchool of Agrifood and Forestry Science and Engineering (ETSEA)University of Lleida‐Agrotecnio CenterLleidaSpain
| | - Xin Jin
- Department of Plant Production and Forestry ScienceSchool of Agrifood and Forestry Science and Engineering (ETSEA)University of Lleida‐Agrotecnio CenterLleidaSpain
| | - Vicente Medina
- Department of Plant Production and Forestry ScienceSchool of Agrifood and Forestry Science and Engineering (ETSEA)University of Lleida‐Agrotecnio CenterLleidaSpain
| | - Gemma Villorbina
- Department of Plant Production and Forestry ScienceSchool of Agrifood and Forestry Science and Engineering (ETSEA)University of Lleida‐Agrotecnio CenterLleidaSpain
| | - Pilar Muñoz
- Department of Plant Production and Forestry ScienceSchool of Agrifood and Forestry Science and Engineering (ETSEA)University of Lleida‐Agrotecnio CenterLleidaSpain
| | - Gemma Farré
- Department of Plant Production and Forestry ScienceSchool of Agrifood and Forestry Science and Engineering (ETSEA)University of Lleida‐Agrotecnio CenterLleidaSpain
| | - Rainer Fischer
- Institute for Molecular BiotechnologyRWTH Aachen UniversityAachenGermany
- Fraunhofer Institute for Molecular Biology and Applied Ecology IMEAachenGermany
| | | | - Teresa Capell
- Department of Plant Production and Forestry ScienceSchool of Agrifood and Forestry Science and Engineering (ETSEA)University of Lleida‐Agrotecnio CenterLleidaSpain
| | - Paul Christou
- Department of Plant Production and Forestry ScienceSchool of Agrifood and Forestry Science and Engineering (ETSEA)University of Lleida‐Agrotecnio CenterLleidaSpain
- ICREACatalan Institute for Research and Advanced StudiesBarcelonaSpain
| | - Stefan Schillberg
- Fraunhofer Institute for Molecular Biology and Applied Ecology IMEAachenGermany
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310
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Blanco-Sánchez B, Clément A, Phillips JB, Westerfield M. Zebrafish models of human eye and inner ear diseases. Methods Cell Biol 2016; 138:415-467. [PMID: 28129854 DOI: 10.1016/bs.mcb.2016.10.006] [Citation(s) in RCA: 35] [Impact Index Per Article: 4.4] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/27/2022]
Abstract
Eye and inner ear diseases are the most common sensory impairments that greatly impact quality of life. Zebrafish have been intensively employed to understand the fundamental mechanisms underlying eye and inner ear development. The zebrafish visual and vestibulo-acoustic systems are very similar to these in humans, and although not yet mature, they are functional by 5days post-fertilization (dpf). In this chapter, we show how the zebrafish has significantly contributed to the field of biomedical research and how researchers, by establishing disease models and meticulously characterizing their phenotypes, have taken the first steps toward therapies. We review here models for (1) eye diseases, (2) ear diseases, and (3) syndromes affecting eye and/or ear. The use of new genome editing technologies and high-throughput screening systems should increase considerably the speed at which knowledge from zebrafish disease models is acquired, opening avenues for better diagnostics, treatments, and therapies.
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Affiliation(s)
| | - A Clément
- University of Oregon, Eugene, OR, United States
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311
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Li M, Zhao L, Page-McCaw PS, Chen W. Zebrafish Genome Engineering Using the CRISPR-Cas9 System. Trends Genet 2016; 32:815-827. [PMID: 27836208 DOI: 10.1016/j.tig.2016.10.005] [Citation(s) in RCA: 105] [Impact Index Per Article: 13.1] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 04/24/2016] [Revised: 10/05/2016] [Accepted: 10/10/2016] [Indexed: 12/21/2022]
Abstract
Geneticists have long sought the ability to manipulate vertebrate genomes by directly altering the information encoded in specific genes. The recently discovered clustered regularly interspaced short palindromic repeats (CRISPR)-Cas9 endonuclease has the ability to bind single loci within vertebrate genomes and generate double-strand breaks (DSBs) at those sites. These DSBs induce an endogenous DSB repair response that results in small insertions or deletions at the targeted site. Alternatively, a template can be supplied, in which case homology-directed repair results in the generation of engineered alleles at the break site. These changes alter the function of the targeted gene facilitating the analysis of gene function. This tool has been widely adopted in the zebrafish model; we discuss the development of this system in the zebrafish and how it can be manipulated to facilitate genome engineering.
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Affiliation(s)
- Mingyu Li
- School of Pharmaceutical Sciences, Xiamen University, Xiamen 361102, China
| | - Liyuan Zhao
- Third Institute of Oceanography, State Oceanic Administration, Xiamen 361005, China
| | - Patrick S Page-McCaw
- Department of Molecular Physiology and Biophysics, Vanderbilt University School of Medicine, Nashville, TN 37232, USA; Division of Nephrology and Hypertension, Vanderbilt Center for Kidney Disease, Vanderbilt University Medical Center, Nashville, TN.
| | - Wenbiao Chen
- Department of Molecular Physiology and Biophysics, Vanderbilt University School of Medicine, Nashville, TN 37232, USA.
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312
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Liu J, Zhou Y, Qi X, Chen J, Chen W, Qiu G, Wu Z, Wu N. CRISPR/Cas9 in zebrafish: an efficient combination for human genetic diseases modeling. Hum Genet 2016; 136:1-12. [PMID: 27807677 PMCID: PMC5214880 DOI: 10.1007/s00439-016-1739-6] [Citation(s) in RCA: 66] [Impact Index Per Article: 8.3] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/25/2016] [Accepted: 10/17/2016] [Indexed: 12/26/2022]
Abstract
The next-generation sequencing identifies a growing number of candidate genes associated with human genetic diseases, which inevitably requires efficient methods to validate the causal links between genotype and phenotype. Recently, zebrafish, with sufficiently high-throughput capabilities, has become a favored option to study human pathogenesis. In addition, CRISPR/Cas9-based approaches have radically reduced the efforts to introduce targeted genome engineering in various organisms. Here, we systemically review the basic considerations in the design of gene editing in zebrafish with CRISPR/Cas9, and explore the potential of the combination of these two to support efficient functional analysis of human genetic variants.
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Affiliation(s)
- Jiaqi Liu
- Department of Orthopaedic Surgery, Peking Union Medical College Hospital, Peking Union Medical College and Chinese Academy of Medical Sciences, No. 1 Shuaifuyuan, Beijing, 100730, China.,Beijing Key Laboratory for Genetic Research of Skeletal Deformity, Beijing, China.,Department of Breast Surgical Oncology, National Cancer Center/Cancer Hospital, Chinese Academy of Medical Sciences and Peking Union Medical College, Beijing, China
| | - Yangzhong Zhou
- Beijing Key Laboratory for Genetic Research of Skeletal Deformity, Beijing, China.,Medical Research Center of Orthopaedics, Chinese Academy of Medical Sciences, Beijing, China.,Tsinghua University Medical School, Beijing, China
| | - Xiaolong Qi
- Department of General Surgery, Nanfang Hospital, Southern Medical University, Guangzhou, China
| | - Jia Chen
- Department of Orthopaedic Surgery, Peking Union Medical College Hospital, Peking Union Medical College and Chinese Academy of Medical Sciences, No. 1 Shuaifuyuan, Beijing, 100730, China.,Beijing Key Laboratory for Genetic Research of Skeletal Deformity, Beijing, China.,Medical Research Center of Orthopaedics, Chinese Academy of Medical Sciences, Beijing, China
| | - Weisheng Chen
- Department of Orthopaedic Surgery, Peking Union Medical College Hospital, Peking Union Medical College and Chinese Academy of Medical Sciences, No. 1 Shuaifuyuan, Beijing, 100730, China.,Beijing Key Laboratory for Genetic Research of Skeletal Deformity, Beijing, China.,Medical Research Center of Orthopaedics, Chinese Academy of Medical Sciences, Beijing, China
| | - Guixing Qiu
- Department of Orthopaedic Surgery, Peking Union Medical College Hospital, Peking Union Medical College and Chinese Academy of Medical Sciences, No. 1 Shuaifuyuan, Beijing, 100730, China.,Beijing Key Laboratory for Genetic Research of Skeletal Deformity, Beijing, China.,Medical Research Center of Orthopaedics, Chinese Academy of Medical Sciences, Beijing, China
| | - Zhihong Wu
- Beijing Key Laboratory for Genetic Research of Skeletal Deformity, Beijing, China. .,Medical Research Center of Orthopaedics, Chinese Academy of Medical Sciences, Beijing, China. .,Department of Central Laboratory, Peking Union Medical College Hospital, Peking Union Medical College and Chinese Academy of Medical Sciences, No. 1 Shuaifuyuan, Beijing, 100730, China.
| | - Nan Wu
- Department of Orthopaedic Surgery, Peking Union Medical College Hospital, Peking Union Medical College and Chinese Academy of Medical Sciences, No. 1 Shuaifuyuan, Beijing, 100730, China. .,Beijing Key Laboratory for Genetic Research of Skeletal Deformity, Beijing, China. .,Medical Research Center of Orthopaedics, Chinese Academy of Medical Sciences, Beijing, China.
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313
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A high-throughput functional genomics workflow based on CRISPR/Cas9-mediated targeted mutagenesis in zebrafish. Nat Protoc 2016; 11:2357-2375. [PMID: 27809318 DOI: 10.1038/nprot.2016.141] [Citation(s) in RCA: 128] [Impact Index Per Article: 16.0] [Reference Citation Analysis] [Abstract] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/20/2022]
Abstract
The zebrafish is a popular model organism for studying development and disease, and genetically modified zebrafish provide an essential tool for functional genomic studies. Numerous publications have demonstrated the efficacy of gene targeting in zebrafish using CRISPR/Cas9, and they have included descriptions of a variety of tools and methods for guide RNA synthesis and mutant identification. However, most of the published techniques are not readily scalable to increase throughput. We recently described a CRISPR/Cas9-based high-throughput mutagenesis and phenotyping pipeline in zebrafish. Here, we present a complete workflow for this pipeline, including target selection; cloning-free single-guide RNA (sgRNA) synthesis; microinjection; validation of the target-specific activity of the sgRNAs; founder screening to identify germline-transmitting mutations by fluorescence PCR; determination of the exact lesion by Sanger or next-generation sequencing (including software for analysis); and genotyping in the F1 or subsequent generations. Using these methods, sgRNAs can be evaluated in 3 d, zebrafish germline-transmitting mutations can be identified within 3 months and stable lines can be established within 6 months. Realistically, two researchers can target tens to hundreds of genes per year using this protocol.
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314
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Pei W, Huang SC, Xu L, Pettie K, Ceci ML, Sánchez M, Allende ML, Burgess SM. Loss of Mgat5a-mediated N-glycosylation stimulates regeneration in zebrafish. ACTA ACUST UNITED AC 2016; 5:3. [PMID: 27795824 PMCID: PMC5072312 DOI: 10.1186/s13619-016-0031-5] [Citation(s) in RCA: 4] [Impact Index Per Article: 0.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/21/2016] [Accepted: 09/20/2016] [Indexed: 12/27/2022]
Abstract
Background We are using genetics to identify genes specifically involved in hearing regeneration. In a large-scale genetic screening, we identified mgat5a, a gene in the N-glycosylation biosynthesis pathway whose activity negatively impacts hair cell regeneration. Methods We used a combination of mutant analysis in zebrafish and a hair cell regeneration assay to phenotype the loss of Mgat5a activity in zebrafish. We used pharmacological inhibition of N-glycosylation by swansonine. We also used over-expression analysis by mRNA injections to demonstrate how changes in N-glycosylation can alter cell signaling. Results We found that mgat5a was expressed in multiple tissues during zebrafish embryo development, particularly enriched in neural tissues including the brain, retina, and lateral line neuromasts. An mgat5a insertional mutation and a CRISPR/Cas9-generated truncation mutation both caused an enhancement of hair cell regeneration which could be phenocopied by pharmacological inhibition with swansonine. In addition to hair cell regeneration, inhibition of the N-glycosylation pathway also enhanced the regeneration of lateral line axon and caudal fins. Further analysis showed that N-glycosylation altered the responsiveness of TGF-beta signaling. Conclusions The findings from this study provide experimental evidence for the involvement of N-glycosylation in tissue regeneration and cell signaling. Electronic supplementary material The online version of this article (doi:10.1186/s13619-016-0031-5) contains supplementary material, which is available to authorized users.
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Affiliation(s)
- Wuhong Pei
- Functional and Translation Genome Branch, National Human Genome Research Institute, 9000 Rockville Pike, Building 50, Room 5537, Bethesda, MD 20892 USA
| | - Sunny C Huang
- Functional and Translation Genome Branch, National Human Genome Research Institute, 9000 Rockville Pike, Building 50, Room 5537, Bethesda, MD 20892 USA
| | - Lisha Xu
- Functional and Translation Genome Branch, National Human Genome Research Institute, 9000 Rockville Pike, Building 50, Room 5537, Bethesda, MD 20892 USA
| | - Kade Pettie
- Functional and Translation Genome Branch, National Human Genome Research Institute, 9000 Rockville Pike, Building 50, Room 5537, Bethesda, MD 20892 USA
| | - María Laura Ceci
- Center for Genome Regulation, Facultad de Ciencias, Universidad de Chile, Casilla 653, Santiago, Chile
| | - Mario Sánchez
- Center for Genome Regulation, Facultad de Ciencias, Universidad de Chile, Casilla 653, Santiago, Chile
| | - Miguel L Allende
- Center for Genome Regulation, Facultad de Ciencias, Universidad de Chile, Casilla 653, Santiago, Chile
| | - Shawn M Burgess
- Functional and Translation Genome Branch, National Human Genome Research Institute, 9000 Rockville Pike, Building 50, Room 5537, Bethesda, MD 20892 USA
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315
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D'Agostino Y, Locascio A, Ristoratore F, Sordino P, Spagnuolo A, Borra M, D'Aniello S. A Rapid and Cheap Methodology for CRISPR/Cas9 Zebrafish Mutant Screening. Mol Biotechnol 2016; 58:73-8. [PMID: 26676479 PMCID: PMC4709366 DOI: 10.1007/s12033-015-9905-y] [Citation(s) in RCA: 22] [Impact Index Per Article: 2.8] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/26/2022]
Abstract
The introduction of new genome editing tools such as ZFNs, TALENs and, more recently, the CRISPR/Cas9 system, has greatly expanded the ability to knock-out genes in different animal models, including zebrafish. However, time and costs required for the screening of a huge number of animals, aimed to identify first founder fishes (F0), and then carriers (F1) are still a bottleneck. Currently, high-resolution melting (HRM) analysis is the most efficient technology for large-scale InDels detection, but the very expensive equipment demanded for its application may represent a limitation for research laboratories. Here, we propose a rapid and cheap method for high-throughput genotyping that displays efficiency rate similar to the HRM. In fact, using a common ViiA™7 real-time PCR system and optimizing the parameters of the melting analysis, we demonstrated that it is possible to discriminate between the mutant and the wild type melting curves. Due to its simplicity, rapidity and cheapness, our method can be used as a preliminary one-step approach for massive screening, in order to restrict the scope at a limited number of embryos and to focus merely on them for the next sequencing step, necessary for the exact sequence identification of the induced mutation. Moreover, thanks to its versatility, this simple approach can be readily adapted to the detection of any kind of genome editing approach directed to genes or regulatory regions and can be applied to many other animal models.
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Affiliation(s)
- Ylenia D'Agostino
- Biology and Evolution of Marine Organisms (BEOM), Stazione Zoologica Anton Dohrn, Villa Comunale, 80121, Naples, Italy
| | - Annamaria Locascio
- Biology and Evolution of Marine Organisms (BEOM), Stazione Zoologica Anton Dohrn, Villa Comunale, 80121, Naples, Italy
| | - Filomena Ristoratore
- Biology and Evolution of Marine Organisms (BEOM), Stazione Zoologica Anton Dohrn, Villa Comunale, 80121, Naples, Italy
| | - Paolo Sordino
- Biology and Evolution of Marine Organisms (BEOM), Stazione Zoologica Anton Dohrn, Villa Comunale, 80121, Naples, Italy
| | - Antonietta Spagnuolo
- Biology and Evolution of Marine Organisms (BEOM), Stazione Zoologica Anton Dohrn, Villa Comunale, 80121, Naples, Italy
| | - Marco Borra
- Research Infrastructures for Marine Biological Resources (RIMAR), Stazione Zoologica Anton Dohrn, Villa Comunale, 80121, Naples, Italy.
| | - Salvatore D'Aniello
- Biology and Evolution of Marine Organisms (BEOM), Stazione Zoologica Anton Dohrn, Villa Comunale, 80121, Naples, Italy.
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316
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A novel technique based on in vitro oocyte injection to improve CRISPR/Cas9 gene editing in zebrafish. Sci Rep 2016; 6:34555. [PMID: 27680290 PMCID: PMC5041118 DOI: 10.1038/srep34555] [Citation(s) in RCA: 17] [Impact Index Per Article: 2.1] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/17/2016] [Accepted: 09/15/2016] [Indexed: 12/31/2022] Open
Abstract
Contemporary improvements in the type II clustered regularly interspaced short palindromic repeats/CRISPR-associated protein 9 (CRISPR/Cas9) system offer a convenient way for genome editing in zebrafish. However, the low efficiencies of genome editing and germline transmission require a time-intensive and laborious screening work. Here, we reported a method based on in vitro oocyte storage by injecting oocytes in advance and incubating them in oocyte storage medium to significantly improve the efficiencies of genome editing and germline transmission by in vitro fertilization (IVF) in zebrafish. Compared to conventional methods, the prior micro-injection of zebrafish oocytes improved the efficiency of genome editing, especially for the sgRNAs with low targeting efficiency. Due to high throughputs, simplicity and flexible design, this novel strategy will provide an efficient alternative to increase the speed of generating heritable mutants in zebrafish by using CRISPR/Cas9 system.
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317
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Ceasar SA, Rajan V, Prykhozhij SV, Berman JN, Ignacimuthu S. Insert, remove or replace: A highly advanced genome editing system using CRISPR/Cas9. BIOCHIMICA ET BIOPHYSICA ACTA 2016; 1863:2333-44. [PMID: 27350235 DOI: 10.1016/j.bbamcr.2016.06.009] [Citation(s) in RCA: 69] [Impact Index Per Article: 8.6] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Received: 05/09/2016] [Revised: 06/21/2016] [Accepted: 06/22/2016] [Indexed: 12/26/2022]
Abstract
The clustered, regularly interspaced, short palindromic repeat (CRISPR) and CRISPR associated protein 9 (Cas9) system discovered as an adaptive immunity mechanism in prokaryotes has emerged as the most popular tool for the precise alterations of the genomes of diverse species. CRISPR/Cas9 system has taken the world of genome editing by storm in recent years. Its popularity as a tool for altering genomes is due to the ability of Cas9 protein to cause double-stranded breaks in DNA after binding with short guide RNA molecules, which can be produced with dramatically less effort and expense than required for production of transcription-activator like effector nucleases (TALEN) and zinc-finger nucleases (ZFN). This system has been exploited in many species from prokaryotes to higher animals including human cells as evidenced by the literature showing increasing sophistication and ease of CRISPR/Cas9 as well as increasing species variety where it is applicable. This technology is poised to solve several complex molecular biology problems faced in life science research including cancer research. In this review, we highlight the recent advancements in CRISPR/Cas9 system in editing genomes of prokaryotes, fungi, plants and animals and provide details on software tools available for convenient design of CRISPR/Cas9 targeting plasmids. We also discuss the future prospects of this advanced molecular technology.
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Affiliation(s)
- S Antony Ceasar
- Division of Plant Biotechnology, Entomology Research Institute, Loyola College, Chennai, India; Centre for Plant Sciences and School of Molecular and Cellular Biology, Faculty of Biological Sciences, University of Leeds, Leeds LS2 9JT, UK
| | - Vinothkumar Rajan
- Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada
| | - Sergey V Prykhozhij
- Department of Pediatrics, Dalhousie University, Halifax, Nova Scotia, Canada
| | - Jason N Berman
- Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada; Department of Pediatrics, Dalhousie University, Halifax, Nova Scotia, Canada.
| | - S Ignacimuthu
- Division of Plant Biotechnology, Entomology Research Institute, Loyola College, Chennai, India; International Scientific Partnership Program, Deanship of Scientific Research, College of Science, King Saud University, Riyadh, Saudi Arabia.
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318
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Narayanan A, Hill-Teran G, Moro A, Ristori E, Kasper DM, A. Roden C, Lu J, Nicoli S. In vivo mutagenesis of miRNA gene families using a scalable multiplexed CRISPR/Cas9 nuclease system. Sci Rep 2016; 6:32386. [PMID: 27572667 PMCID: PMC5004112 DOI: 10.1038/srep32386] [Citation(s) in RCA: 27] [Impact Index Per Article: 3.4] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/31/2016] [Accepted: 08/08/2016] [Indexed: 01/06/2023] Open
Abstract
A large number of microRNAs (miRNAs) are grouped into families derived from the same phylogenetic ancestors. miRNAs within a family often share the same physiological functions despite differences in their primary sequences, secondary structures, or chromosomal locations. Consequently, the generation of animal models to analyze the activity of miRNA families is extremely challenging. Using zebrafish as a model system, we successfully provide experimental evidence that a large number of miRNAs can be simultaneously mutated to abrogate the activity of an entire miRNA family. We show that injection of the Cas9 nuclease and two, four, ten, and up to twenty-four multiplexed single guide RNAs (sgRNAs) can induce mutations in 90% of the miRNA genomic sequences analyzed. We performed a survey of these 45 mutations in 10 miRNA genes, analyzing the impact of our mutagenesis strategy on the processing of each miRNA both computationally and in vivo. Our results offer an effective approach to mutate and study the activity of miRNA families and pave the way for further analysis on the function of complex miRNA families in higher multicellular organisms.
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Affiliation(s)
- Anand Narayanan
- Yale Cardiovascular Research Center, Section of Cardiology, Department of Internal Medicine, Yale University School of Medicine, New Haven, CT 06510, USA
| | - Guillermina Hill-Teran
- Yale Cardiovascular Research Center, Section of Cardiology, Department of Internal Medicine, Yale University School of Medicine, New Haven, CT 06510, USA
| | - Albertomaria Moro
- Yale Cardiovascular Research Center, Section of Cardiology, Department of Internal Medicine, Yale University School of Medicine, New Haven, CT 06510, USA
| | - Emma Ristori
- Yale Cardiovascular Research Center, Section of Cardiology, Department of Internal Medicine, Yale University School of Medicine, New Haven, CT 06510, USA
| | - Dionna M. Kasper
- Yale Cardiovascular Research Center, Section of Cardiology, Department of Internal Medicine, Yale University School of Medicine, New Haven, CT 06510, USA
| | - Christine A. Roden
- Department of Genetics, Yale University School of Medicine, New Haven, CT 06510, USA
- Yale Stem Cell Center and Yale Cancer Center, Yale University, New Haven, CT, 06520, USA
| | - Jun Lu
- Department of Genetics, Yale University School of Medicine, New Haven, CT 06510, USA
- Yale Stem Cell Center and Yale Cancer Center, Yale University, New Haven, CT, 06520, USA
| | - Stefania Nicoli
- Yale Cardiovascular Research Center, Section of Cardiology, Department of Internal Medicine, Yale University School of Medicine, New Haven, CT 06510, USA
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319
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Elks PM, Renshaw SA, Meijer AH, Walmsley SR, van Eeden FJ. Exploring the HIFs, buts and maybes of hypoxia signalling in disease: lessons from zebrafish models. Dis Model Mech 2016; 8:1349-60. [PMID: 26512123 PMCID: PMC4631790 DOI: 10.1242/dmm.021865] [Citation(s) in RCA: 42] [Impact Index Per Article: 5.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/31/2022] Open
Abstract
A low level of tissue oxygen (hypoxia) is a physiological feature of a wide range of diseases, from cancer to infection. Cellular hypoxia is sensed by oxygen-sensitive hydroxylase enzymes, which regulate the protein stability of hypoxia-inducible factor α (HIF-α) transcription factors. When stabilised, HIF-α binds with its cofactors to HIF-responsive elements (HREs) in the promoters of target genes to coordinate a wide-ranging transcriptional programme in response to the hypoxic environment. This year marks the 20th anniversary of the discovery of the HIF-1α transcription factor, and in recent years the HIF-mediated hypoxia response is being increasingly recognised as an important process in determining the outcome of diseases such as cancer, inflammatory disease and bacterial infections. Animal models have shed light on the roles of HIF in disease and have uncovered intricate control mechanisms that involve multiple cell types, observations that might have been missed in simpler in vitro systems. These findings highlight the need for new whole-organism models of disease to elucidate these complex regulatory mechanisms. In this Review, we discuss recent advances in our understanding of hypoxia and HIFs in disease that have emerged from studies of zebrafish disease models. Findings from such models identify HIF as an integral player in the disease processes. They also highlight HIF pathway components and their targets as potential therapeutic targets against conditions that range from cancers to infectious disease. Summary: Hypoxia signalling, mediated by HIF, is a crucial pathway in many disease processes. Here, we review current knowledge of HIF signalling and disease, focusing on recent findings from zebrafish models.
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Affiliation(s)
- Philip M Elks
- Department of Infection and Immunity, Medical School, The University of Sheffield, Sheffield, S10 2RX, UK The Bateson Centre, The University of Sheffield, Sheffield, S10 2TN, UK
| | - Stephen A Renshaw
- Department of Infection and Immunity, Medical School, The University of Sheffield, Sheffield, S10 2RX, UK The Bateson Centre, The University of Sheffield, Sheffield, S10 2TN, UK
| | - Annemarie H Meijer
- Institute of Biology, Leiden University, 2333 CC Leiden, The Netherlands
| | - Sarah R Walmsley
- MRC Centre for Inflammation Research, University of Edinburgh, Edinburgh, EH16 4TJ, UK
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320
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Santos DP, Kiskinis E, Eggan K, Merkle FT. Comprehensive Protocols for CRISPR/Cas9-based Gene Editing in Human Pluripotent Stem Cells. CURRENT PROTOCOLS IN STEM CELL BIOLOGY 2016; 38:5B.6.1-5B.6.60. [PMID: 27532820 PMCID: PMC4988528 DOI: 10.1002/cpsc.15] [Citation(s) in RCA: 25] [Impact Index Per Article: 3.1] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 12/17/2022]
Abstract
Genome editing of human pluripotent stem cells (hPSCs) with the CRISPR/Cas9 system has the potential to revolutionize hPSC-based disease modeling, drug screening, and transplantation therapy. Here, we aim to provide a single resource to enable groups, even those with limited experience with hPSC culture or the CRISPR/Cas9 system, to successfully perform genome editing. The methods are presented in detail and are supported by a theoretical framework to allow for the incorporation of inevitable improvements in the rapidly evolving gene-editing field. We describe protocols to generate hPSC lines with gene-specific knock-outs, small targeted mutations, or knock-in reporters. © 2016 by John Wiley & Sons, Inc.
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Affiliation(s)
- David P. Santos
- The Ken & Ruth Davee Department of Neurology & Clinical Neurological Sciences, Department of Physiology, Feinberg School of Medicine, Northwestern University, Chicago, IL, 60611, USA
| | - Evangelos Kiskinis
- The Ken & Ruth Davee Department of Neurology & Clinical Neurological Sciences, Department of Physiology, Feinberg School of Medicine, Northwestern University, Chicago, IL, 60611, USA
| | - Kevin Eggan
- Department of Stem Cell and Regenerative Biology, and Harvard Stem Cell Institute, Harvard University, Cambridge, MA 02138, USA
- Stanley Center for Psychiatric Research, Broad Institute of Harvard and MIT, Cambridge, MA 02142, USA
| | - Florian T. Merkle
- Metabolic Research Laboratories and MRC Metabolic Diseases Unit, Wellcome Trust-MRC Institute of Metabolic Science, and Wellcome Trust-MRC Cambridge Stem Cell Institute, University of Cambridge, Cambridge CB2 0QQ, UK
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321
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Abstract
Programmable nucleases have revolutionized zebrafish genetics by enabling targeted genome modifications. In this issue of Developmental Cell, Hoshijima et al. (2016) take genome modification in the zebrafish to the next level, demonstrating the efficient use of homologous recombination to make genetic tools for a range of applications.
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Affiliation(s)
- Arish N Shah
- Division of Basic Sciences, Fred Hutchinson Cancer Research Center, B2-152, 1100 Fairview Avenue North, Seattle, WA 98109-1024, USA
| | - Cecilia B Moens
- Division of Basic Sciences, Fred Hutchinson Cancer Research Center, B2-152, 1100 Fairview Avenue North, Seattle, WA 98109-1024, USA.
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322
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Markossian S, Flamant F. CRISPR/Cas9: a breakthrough in generating mouse models for endocrinologists. J Mol Endocrinol 2016; 57:R81-92. [PMID: 27272521 DOI: 10.1530/jme-15-0305] [Citation(s) in RCA: 8] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Submit a Manuscript] [Subscribe] [Scholar Register] [Received: 06/03/2016] [Accepted: 06/07/2016] [Indexed: 12/26/2022]
Abstract
CRISPR/Cas9 is a recent development in genome editing which is becoming an indispensable element of the genetic toolbox in mice. It provides outstanding possibilities for targeted modification of the genome, and is often extremely efficient. There are currently two main limitations to in ovo genome editing in mice: the first is mosaicism, which is frequent in founder mice. The second is the difficulty to evaluate the advent of off-target mutations, which often imposes to wait for germline transmission to ensure genetic segregation between wanted and unwanted genetic mutations. However rapid progresses are made, suggesting that these difficulties can be overcome in the near future.
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Affiliation(s)
- Suzy Markossian
- Institut de Génomique Fonctionnelle de LyonUniversité de Lyon, CNRS, INRA, École Normale Supérieure de Lyon, Lyon Cedex 07, France
| | - Frédéric Flamant
- Institut de Génomique Fonctionnelle de LyonUniversité de Lyon, CNRS, INRA, École Normale Supérieure de Lyon, Lyon Cedex 07, France
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323
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Blitz IL, Fish MB, Cho KWY. Leapfrogging: primordial germ cell transplantation permits recovery of CRISPR/Cas9-induced mutations in essential genes. Development 2016; 143:2868-75. [PMID: 27385011 PMCID: PMC5004912 DOI: 10.1242/dev.138057] [Citation(s) in RCA: 24] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/29/2016] [Accepted: 06/15/2016] [Indexed: 01/07/2023]
Abstract
CRISPR/Cas9 genome editing is revolutionizing genetic loss-of-function analysis but technical limitations remain that slow progress when creating mutant lines. First, in conventional genetic breeding schemes, mosaic founder animals carrying mutant alleles are outcrossed to produce F1 heterozygotes. Phenotypic analysis occurs in the F2 generation following F1 intercrosses. Thus, mutant analyses will require multi-generational studies. Second, when targeting essential genes, efficient mutagenesis of founders is often lethal, preventing the acquisition of mature animals. Reducing mutagenesis levels may improve founder survival, but results in lower, more variable rates of germline transmission. Therefore, an efficient approach to study lethal mutations would be useful. To overcome these shortfalls, we introduce 'leapfrogging', a method combining efficient CRISPR mutagenesis with transplantation of mutated primordial germ cells into a wild-type host. Tested using Xenopus tropicalis, we show that founders containing transplants transmit mutant alleles with high efficiency. F1 offspring from intercrosses between F0 animals that carry embryonic lethal alleles recapitulate loss-of-function phenotypes, circumventing an entire generation of breeding. We anticipate that leapfrogging will be transferable to other species.
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Affiliation(s)
- Ira L Blitz
- 4410 Natural Sciences Building 2, Department of Developmental and Cell Biology, University of California, Irvine, CA 92697, USA
| | - Margaret B Fish
- 4410 Natural Sciences Building 2, Department of Developmental and Cell Biology, University of California, Irvine, CA 92697, USA
| | - Ken W Y Cho
- 4410 Natural Sciences Building 2, Department of Developmental and Cell Biology, University of California, Irvine, CA 92697, USA
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324
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BATCH-GE: Batch analysis of Next-Generation Sequencing data for genome editing assessment. Sci Rep 2016; 6:30330. [PMID: 27461955 PMCID: PMC4962088 DOI: 10.1038/srep30330] [Citation(s) in RCA: 63] [Impact Index Per Article: 7.9] [Reference Citation Analysis] [Abstract] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/07/2016] [Accepted: 07/04/2016] [Indexed: 02/02/2023] Open
Abstract
Targeted mutagenesis by the CRISPR/Cas9 system is currently revolutionizing genetics. The ease of this technique has enabled genome engineering in-vitro and in a range of model organisms and has pushed experimental dimensions to unprecedented proportions. Due to its tremendous progress in terms of speed, read length, throughput and cost, Next-Generation Sequencing (NGS) has been increasingly used for the analysis of CRISPR/Cas9 genome editing experiments. However, the current tools for genome editing assessment lack flexibility and fall short in the analysis of large amounts of NGS data. Therefore, we designed BATCH-GE, an easy-to-use bioinformatics tool for batch analysis of NGS-generated genome editing data, available from https://github.com/WouterSteyaert/BATCH-GE.git. BATCH-GE detects and reports indel mutations and other precise genome editing events and calculates the corresponding mutagenesis efficiencies for a large number of samples in parallel. Furthermore, this new tool provides flexibility by allowing the user to adapt a number of input variables. The performance of BATCH-GE was evaluated in two genome editing experiments, aiming to generate knock-out and knock-in zebrafish mutants. This tool will not only contribute to the evaluation of CRISPR/Cas9-based experiments, but will be of use in any genome editing experiment and has the ability to analyze data from every organism with a sequenced genome.
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325
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Shao M, Xu TR, Chen CS. The big bang of genome editing technology: development and application of the CRISPR/Cas9 system in disease animal models. DONG WU XUE YAN JIU = ZOOLOGICAL RESEARCH 2016; 37:191-204. [PMID: 27469250 PMCID: PMC4980067 DOI: 10.13918/j.issn.2095-8137.2016.4.191] [Citation(s) in RCA: 12] [Impact Index Per Article: 1.5] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Subscribe] [Scholar Register] [Received: 11/10/2015] [Accepted: 01/20/2016] [Indexed: 12/17/2022]
Abstract
Targeted genome editing technology has been widely used in biomedical studies. The CRISPR-associated RNA-guided endonuclease Cas9 has become a versatile genome editing tool. The CRISPR/Cas9 system is useful for studying gene function through efficient knock-out, knock-in or chromatin modification of the targeted gene loci in various cell types and organisms. It can be applied in a number of fields, such as genetic breeding, disease treatment and gene functional investigation. In this review, we introduce the most recent developments and applications, the challenges, and future directions of Cas9 in generating disease animal model. Derived from the CRISPR adaptive immune system of bacteria, the development trend of Cas9 will inevitably fuel the vital applications from basic research to biotechnology and bio-medicine.
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Affiliation(s)
- Ming Shao
- Faculty of Life Science and Technology, Kunming University of Science and Technology, Kunming Yunnan 650500, China;Key Laboratory of Animal Models and Human Disease Mechanisms of the Chinese Academy of Sciences and Yunnan Province, Kunming Institute of Zoology, Chinese Academy of Sciences, Kunming Yunnan 650223, China
| | - Tian-Rui Xu
- Faculty of Life Science and Technology, Kunming University of Science and Technology, Kunming Yunnan 650500, China.
| | - Ce-Shi Chen
- Key Laboratory of Animal Models and Human Disease Mechanisms of the Chinese Academy of Sciences and Yunnan Province, Kunming Institute of Zoology, Chinese Academy of Sciences, Kunming Yunnan 650223, China.
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326
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Additive reductions in zebrafish PRPS1 activity result in a spectrum of deficiencies modeling several human PRPS1-associated diseases. Sci Rep 2016; 6:29946. [PMID: 27425195 PMCID: PMC4947902 DOI: 10.1038/srep29946] [Citation(s) in RCA: 25] [Impact Index Per Article: 3.1] [Reference Citation Analysis] [Abstract] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/22/2016] [Accepted: 06/27/2016] [Indexed: 01/08/2023] Open
Abstract
Phosphoribosyl pyrophosphate synthetase-1 (PRPS1) is a key enzyme in nucleotide biosynthesis, and mutations in PRPS1 are found in several human diseases including nonsyndromic sensorineural deafness, Charcot-Marie-Tooth disease-5, and Arts Syndrome. We utilized zebrafish as a model to confirm that mutations in PRPS1 result in phenotypic deficiencies in zebrafish similar to those in the associated human diseases. We found two paralogs in zebrafish, prps1a and prps1b and characterized each paralogous mutant individually as well as the double mutant fish. Zebrafish prps1a mutants and prps1a;prps1b double mutants showed similar morphological phenotypes with increasingly severe phenotypes as the number of mutant alleles increased. Phenotypes included smaller eyes and reduced hair cell numbers, consistent with the optic atrophy and hearing impairment observed in human patients. The double mutant also showed abnormal development of primary motor neurons, hair cell innervation, and reduced leukocytes, consistent with the neuropathy and recurrent infection of the human patients possessing the most severe reductions of PRPS1 activity. Further analyses indicated the phenotypes were associated with a prolonged cell cycle likely resulting from reduced nucleotide synthesis and energy production in the mutant embryos. We further demonstrated the phenotypes were caused by delays in the tissues most highly expressing the prps1 genes.
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327
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Hao H, Wang X, Jia H, Yu M, Zhang X, Tang H, Zhang L. Large fragment deletion using a CRISPR/Cas9 system in Saccharomyces cerevisiae. Anal Biochem 2016; 509:118-123. [PMID: 27402178 DOI: 10.1016/j.ab.2016.07.008] [Citation(s) in RCA: 22] [Impact Index Per Article: 2.8] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/10/2016] [Revised: 07/04/2016] [Accepted: 07/06/2016] [Indexed: 11/18/2022]
Abstract
Large chromosomal modifications have been performed in natural and laboratory evolution studies and hold tremendous potential for use in foundational research, medicine, and biotechnology applications. Recently, the type II bacterial Clustered Regularly Interspaced Short Palindromic Repeat and CRISPR-associated (CRISPR/Cas9) system has emerged as a powerful tool for genome editing in various organisms. In this study, we applied the CRISPR/Cas9 system to preform large fragment deletions in Saccharomyces cerevisiae and compared the performance activity to that of a traditional method that uses the Latour system. Here we report in S. Cerevisiae the CRIPR/Cas9 system has been used to delete fragments exceeding 30 kb. The use of the CRISPR/Cas9 system for generating chromosomal segment excision showed some potential advantages over the Latour system. All the results indicated that CRISPR/Cas9 system was a rapid, efficient, low-cost, and versatile method for genome editing and that it can be applied in further studies in the fields of biology, agriculture, and medicine.
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Affiliation(s)
- Huanhuan Hao
- Key Laboratory of Microbial Diversity Research and Application of Hebei Province, Key Discipline of Biological Engineering of Hebei Province, College of Life Sciences, Hebei University, Baoding, 071002, China
| | - Xiaofei Wang
- Key Laboratory of Microbial Diversity Research and Application of Hebei Province, Key Discipline of Biological Engineering of Hebei Province, College of Life Sciences, Hebei University, Baoding, 071002, China
| | - Haiyan Jia
- Key Laboratory of Microbial Diversity Research and Application of Hebei Province, Key Discipline of Biological Engineering of Hebei Province, College of Life Sciences, Hebei University, Baoding, 071002, China
| | - Miao Yu
- Key Laboratory of Microbial Diversity Research and Application of Hebei Province, Key Discipline of Biological Engineering of Hebei Province, College of Life Sciences, Hebei University, Baoding, 071002, China
| | - Xiaoyu Zhang
- Key Laboratory of Microbial Diversity Research and Application of Hebei Province, Key Discipline of Biological Engineering of Hebei Province, College of Life Sciences, Hebei University, Baoding, 071002, China
| | - Hui Tang
- Key Laboratory of Microbial Diversity Research and Application of Hebei Province, Key Discipline of Biological Engineering of Hebei Province, College of Life Sciences, Hebei University, Baoding, 071002, China
| | - Liping Zhang
- Key Laboratory of Microbial Diversity Research and Application of Hebei Province, Key Discipline of Biological Engineering of Hebei Province, College of Life Sciences, Hebei University, Baoding, 071002, China.
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328
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Xu W, Jin M, Hu R, Wang H, Zhang F, Yuan S, Cao Y. The Joubert Syndrome Protein Inpp5e Controls Ciliogenesis by Regulating Phosphoinositides at the Apical Membrane. J Am Soc Nephrol 2016; 28:118-129. [PMID: 27401686 DOI: 10.1681/asn.2015080906] [Citation(s) in RCA: 34] [Impact Index Per Article: 4.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/18/2015] [Accepted: 05/30/2016] [Indexed: 12/23/2022] Open
Abstract
Phosphoinositides, a family of phosphorylated derivatives of phosphatidylinositol (PtdIns), are tightly regulated both temporally and spatially by PtdIns phosphatases and kinases. Mutations in inositol polyphosphate 5-phosphatase E (INPP5E) cause Joubert syndrome, a human disorder associated with numerous ciliopathic defects, including renal cyst formation, linking phosphoinositides to ciliopathies. However, the molecular mechanism by which INPP5E-mediated PtdIns signaling regulates ciliogenesis and cystogenesis is unclear. Here, we utilized an in vivo vertebrate model of renal cystogenesis to show that Inpp5e enzymatic activity at the apical membrane directs apical docking of basal bodies in renal epithelia. Knockdown or knockout of inpp5e led to ciliogenesis defects and cystic kidneys in zebrafish. Furthermore, knockdown of inpp5e in embryos led to defects in cell polarity, cortical organization of F-actin, and apical segregation of PtdIns(4,5)P2 and PtdIns(3,4,5)P3 Knockdown of the ezrin gene, which encodes an ezrin/radixin/moesin (ERM) protein that crosslinks PtdIns(4,5)P2 and F-actin, phenocopied inpp5e knockdowns. Notably, overexpression of the ezrin gene rescued inpp5e morphants. Finally, treatment with the PI 3-kinase inhibitor LY294002, which decreases PtdIns(3,4,5)P3 levels, rescued the cellular, phenotypic, and renal functional defects in inpp5e-knockdown embryos. Together, our data indicate that Inpp5e functions as a key regulator of cell polarity in the renal epithelia, by inhibiting PtdIns(3,4,5)P3 and subsequently stabilizing PtdIns(4,5)P2 and recruiting Ezrin, F-actin, and basal bodies to the apical membrane, and suggest a possible novel approach for treating human ciliopathies.
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Affiliation(s)
- Wenyan Xu
- Department of Molecular and Cell Biology, Tongji University School of Life Sciences and Technology, Shanghai, China
| | - Miaomiao Jin
- Department of Molecular and Cell Biology, Tongji University School of Life Sciences and Technology, Shanghai, China
| | - Ruikun Hu
- Department of Molecular and Cell Biology, Tongji University School of Life Sciences and Technology, Shanghai, China
| | - Hong Wang
- Department of Molecular and Cell Biology, Tongji University School of Life Sciences and Technology, Shanghai, China
| | - Fan Zhang
- Department of Molecular and Cell Biology, Tongji University School of Life Sciences and Technology, Shanghai, China
| | - Shiaulou Yuan
- Department of Pediatrics, Yale University School of Medicine, New Haven, Connecticut; and
| | - Ying Cao
- Department of Molecular and Cell Biology, Tongji University School of Life Sciences and Technology, Shanghai, China; .,Tongji University and Shanghai Changzheng Hospital Joint Research Center for Translational Medicine, Changzheng Hospital, Shanghai, China
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329
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Haeussler M, Schönig K, Eckert H, Eschstruth A, Mianné J, Renaud JB, Schneider-Maunoury S, Shkumatava A, Teboul L, Kent J, Joly JS, Concordet JP. Evaluation of off-target and on-target scoring algorithms and integration into the guide RNA selection tool CRISPOR. Genome Biol 2016; 17:148. [PMID: 27380939 PMCID: PMC4934014 DOI: 10.1186/s13059-016-1012-2] [Citation(s) in RCA: 1086] [Impact Index Per Article: 135.8] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/15/2016] [Accepted: 06/17/2016] [Indexed: 11/10/2022] Open
Abstract
BACKGROUND The success of the CRISPR/Cas9 genome editing technique depends on the choice of the guide RNA sequence, which is facilitated by various websites. Despite the importance and popularity of these algorithms, it is unclear to which extent their predictions are in agreement with actual measurements. RESULTS We conduct the first independent evaluation of CRISPR/Cas9 predictions. To this end, we collect data from eight SpCas9 off-target studies and compare them with the sites predicted by popular algorithms. We identify problems in one implementation but found that sequence-based off-target predictions are very reliable, identifying most off-targets with mutation rates superior to 0.1 %, while the number of false positives can be largely reduced with a cutoff on the off-target score. We also evaluate on-target efficiency prediction algorithms against available datasets. The correlation between the predictions and the guide activity varied considerably, especially for zebrafish. Together with novel data from our labs, we find that the optimal on-target efficiency prediction model strongly depends on whether the guide RNA is expressed from a U6 promoter or transcribed in vitro. We further demonstrate that the best predictions can significantly reduce the time spent on guide screening. CONCLUSIONS To make these guidelines easily accessible to anyone planning a CRISPR genome editing experiment, we built a new website ( http://crispor.org ) that predicts off-targets and helps select and clone efficient guide sequences for more than 120 genomes using different Cas9 proteins and the eight efficiency scoring systems evaluated here.
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Affiliation(s)
- Maximilian Haeussler
- Santa Cruz Genomics Institute, MS CBSE, University of California, 1156 High Street, Santa Cruz, CA, 95064, USA.
| | - Kai Schönig
- Central Institute of Mental Health, Medical Faculty Mannheim, Heidelberg University, Square J5, Mannheim, 68159, Germany
| | - Hélène Eckert
- Institut Curie, CNRS UMR3215, INSERM U934, Paris, Cedex, 05 75248, France
| | - Alexis Eschstruth
- CNRS UMR 7622, INSERM U1156, Sorbonne Université Paris 06, Paris, France
| | | | | | | | - Alena Shkumatava
- Institut Curie, CNRS UMR3215, INSERM U934, Paris, Cedex, 05 75248, France
| | | | - Jim Kent
- Santa Cruz Genomics Institute, MS CBSE, University of California, 1156 High Street, Santa Cruz, CA, 95064, USA
| | | | - Jean-Paul Concordet
- INSERM U1154, CNRS UMR 7196, Muséum National d'Histoire Naturelle, Paris, France.
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330
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Pelegri F, Mullins MC. Genetic screens for mutations affecting adult traits and parental-effect genes. Methods Cell Biol 2016; 135:39-87. [PMID: 27443920 DOI: 10.1016/bs.mcb.2016.05.006] [Citation(s) in RCA: 6] [Impact Index Per Article: 0.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 02/08/2023]
Abstract
Forward genetics remains an important approach for the unbiased identification of factors involved in biological pathways. Forward genetic analysis in the zebrafish has until now largely been restricted to the developmental period from zygotic genome activation through the end of embryogenesis. However, the use of the zebrafish as a model system for the analysis of late larval, juvenile and adult traits, including fertility and maternal and paternal effects, continues to gain momentum. Here, we describe two approaches, based on an F3-extended family and gynogenetic methods, that allow genetic screening for, and recovery of mutations affecting post-embryonic stages, including adult traits, fertility, and parental effects. For each approach, we also describe strategies to maintain, map, and molecularly clone the identified mutations.
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Affiliation(s)
- F Pelegri
- University of Wisconsin-Madison, Madison, WI, United States
| | - M C Mullins
- University of Pennsylvania Perelman School of Medicine, Philadelphia, PA, United States
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331
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Kowalko JE, Ma L, Jeffery WR. Genome Editing in Astyanax mexicanus Using Transcription Activator-like Effector Nucleases (TALENs). J Vis Exp 2016. [PMID: 27404092 DOI: 10.3791/54113] [Citation(s) in RCA: 11] [Impact Index Per Article: 1.4] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/12/2022] Open
Abstract
Identifying alleles of genes underlying evolutionary change is essential to understanding how and why evolution occurs. Towards this end, much recent work has focused on identifying candidate genes for the evolution of traits in a variety of species. However, until recently it has been challenging to functionally validate interesting candidate genes. Recently developed tools for genetic engineering make it possible to manipulate specific genes in a wide range of organisms. Application of this technology in evolutionarily relevant organisms will allow for unprecedented insight into the role of candidate genes in evolution. Astyanax mexicanus (A. mexicanus) is a species of fish with both surface-dwelling and cave-dwelling forms. Multiple independent lines of cave-dwelling forms have evolved from ancestral surface fish, which are interfertile with one another and with surface fish, allowing elucidation of the genetic basis of cave traits. A. mexicanus has been used for a number of evolutionary studies, including linkage analysis to identify candidate genes responsible for a number of traits. Thus, A. mexicanus is an ideal system for the application of genome editing to test the role of candidate genes. Here we report a method for using transcription activator-like effector nucleases (TALENs) to mutate genes in surface A. mexicanus. Genome editing using TALENs in A. mexicanus has been utilized to generate mutations in pigmentation genes. This technique can also be utilized to evaluate the role of candidate genes for a number of other traits that have evolved in cave forms of A. mexicanus.
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Affiliation(s)
| | - Li Ma
- Department of Biological Sciences, University of Cincinnati
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332
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Huang B, Gomez-Rodriguez J, Preite S, Garrett LJ, Harper UL, Schwartzberg PL. CRISPR-Mediated Triple Knockout of SLAMF1, SLAMF5 and SLAMF6 Supports Positive Signaling Roles in NKT Cell Development. PLoS One 2016; 11:e0156072. [PMID: 27258160 PMCID: PMC4892526 DOI: 10.1371/journal.pone.0156072] [Citation(s) in RCA: 15] [Impact Index Per Article: 1.9] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/24/2016] [Accepted: 04/08/2016] [Indexed: 01/04/2023] Open
Abstract
The SLAM family receptors contribute to diverse aspects of lymphocyte biology and signal via the small adaptor molecule SAP. Mutations affecting SAP lead to X-linked lymphoproliferative syndrome Type 1, a severe immunodysregulation characterized by fulminant mononucleosis, dysgammaglobulinemia, and lymphoproliferation/lymphomas. Patients and mice having mutations affecting SAP also lack germinal centers due to a defect in T:B cell interactions and are devoid of invariant NKT (iNKT) cells. However, which and how SLAM family members contribute to these phenotypes remains uncertain. Three SLAM family members: SLAMF1, SLAMF5 and SLAMF6, are highly expressed on T follicular helper cells and germinal center B cells. SLAMF1 and SLAMF6 are also implicated in iNKT development. Although individual receptor knockout mice have limited iNKT and germinal center phenotypes compared to SAP knockout mice, the generation of multi-receptor knockout mice has been challenging, due to the genomic linkage of the genes encoding SLAM family members. Here, we used Cas9/CRISPR-based mutagenesis to generate mutations simultaneously in Slamf1, Slamf5 and Slamf6. Genetic disruption of all three receptors in triple-knockout mice (TKO) did not grossly affect conventional T or B cell development and led to mild defects in germinal center formation post-immunization. However, the TKO worsened defects in iNKT cells development seen in SLAMF6 single gene-targeted mice, supporting data on positive signaling and potential redundancy between these receptors.
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Affiliation(s)
- Bonnie Huang
- Genetic Disease Research Branch, National Human Genome Research Institute, Bethesda, Maryland, United States of America
| | - Julio Gomez-Rodriguez
- Genetic Disease Research Branch, National Human Genome Research Institute, Bethesda, Maryland, United States of America
| | - Silvia Preite
- Genetic Disease Research Branch, National Human Genome Research Institute, Bethesda, Maryland, United States of America
| | - Lisa J. Garrett
- Embryonic Stem Cell and Transgenic Mouse Core, National Human Genome Research Institute, Bethesda, Maryland, United States of America
| | - Ursula L. Harper
- Genomics Core, National Human Genome Research Institute, National Human Genome Research Institute, Bethesda, Maryland, United States of America
| | - Pamela L. Schwartzberg
- Genetic Disease Research Branch, National Human Genome Research Institute, Bethesda, Maryland, United States of America
- * E-mail:
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333
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Burger A, Lindsay H, Felker A, Hess C, Anders C, Chiavacci E, Zaugg J, Weber LM, Catena R, Jinek M, Robinson MD, Mosimann C. Maximizing mutagenesis with solubilized CRISPR-Cas9 ribonucleoprotein complexes. Development 2016; 143:2025-37. [PMID: 27130213 DOI: 10.1242/dev.134809] [Citation(s) in RCA: 173] [Impact Index Per Article: 21.6] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/06/2016] [Accepted: 04/12/2016] [Indexed: 12/14/2022]
Abstract
CRISPR-Cas9 enables efficient sequence-specific mutagenesis for creating somatic or germline mutants of model organisms. Key constraints in vivo remain the expression and delivery of active Cas9-sgRNA ribonucleoprotein complexes (RNPs) with minimal toxicity, variable mutagenesis efficiencies depending on targeting sequence, and high mutation mosaicism. Here, we apply in vitro assembled, fluorescent Cas9-sgRNA RNPs in solubilizing salt solution to achieve maximal mutagenesis efficiency in zebrafish embryos. MiSeq-based sequence analysis of targeted loci in individual embryos using CrispRVariants, a customized software tool for mutagenesis quantification and visualization, reveals efficient bi-allelic mutagenesis that reaches saturation at several tested gene loci. Such virtually complete mutagenesis exposes loss-of-function phenotypes for candidate genes in somatic mutant embryos for subsequent generation of stable germline mutants. We further show that targeting of non-coding elements in gene regulatory regions using saturating mutagenesis uncovers functional control elements in transgenic reporters and endogenous genes in injected embryos. Our results establish that optimally solubilized, in vitro assembled fluorescent Cas9-sgRNA RNPs provide a reproducible reagent for direct and scalable loss-of-function studies and applications beyond zebrafish experiments that require maximal DNA cutting efficiency in vivo.
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Affiliation(s)
- Alexa Burger
- Institute of Molecular Life Sciences, University of Zürich, Zürich 8057, Switzerland
| | - Helen Lindsay
- Institute of Molecular Life Sciences, University of Zürich, Zürich 8057, Switzerland SIB Swiss Institute of Bioinformatics, University of Zürich, Zürich 8057, Switzerland
| | - Anastasia Felker
- Institute of Molecular Life Sciences, University of Zürich, Zürich 8057, Switzerland
| | - Christopher Hess
- Institute of Molecular Life Sciences, University of Zürich, Zürich 8057, Switzerland
| | - Carolin Anders
- Institute of Biochemistry, University of Zürich, Zürich 8057, Switzerland
| | - Elena Chiavacci
- Institute of Molecular Life Sciences, University of Zürich, Zürich 8057, Switzerland
| | - Jonas Zaugg
- Institute of Molecular Life Sciences, University of Zürich, Zürich 8057, Switzerland
| | - Lukas M Weber
- Institute of Molecular Life Sciences, University of Zürich, Zürich 8057, Switzerland SIB Swiss Institute of Bioinformatics, University of Zürich, Zürich 8057, Switzerland
| | - Raul Catena
- Institute of Molecular Life Sciences, University of Zürich, Zürich 8057, Switzerland
| | - Martin Jinek
- Institute of Biochemistry, University of Zürich, Zürich 8057, Switzerland
| | - Mark D Robinson
- Institute of Molecular Life Sciences, University of Zürich, Zürich 8057, Switzerland SIB Swiss Institute of Bioinformatics, University of Zürich, Zürich 8057, Switzerland
| | - Christian Mosimann
- Institute of Molecular Life Sciences, University of Zürich, Zürich 8057, Switzerland
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334
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Yin L, Maddison LA, Chen W. Multiplex conditional mutagenesis in zebrafish using the CRISPR/Cas system. Methods Cell Biol 2016; 135:3-17. [PMID: 27443918 DOI: 10.1016/bs.mcb.2016.04.018] [Citation(s) in RCA: 12] [Impact Index Per Article: 1.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/13/2022]
Abstract
The clustered regularly interspaced short palindromic repeat (CRISPR)/CRISPR-associated protein (Cas) system is a powerful tool for genome editing in numerous organisms. However, the system is typically used for gene editing throughout the entire organism. Tissue and temporal specific mutagenesis is often desirable to determine gene function in a specific stage or tissue and to bypass undesired consequences of global mutations. We have developed the CRISPR/Cas system for conditional mutagenesis in transgenic zebrafish using tissue-specific and/or inducible expression of Cas9 and U6-driven expression of sgRNA. To allow mutagenesis of multiple targets, we have isolated four distinct U6 promoters and designed Golden Gate vectors to easily assemble transgenes with multiple sgRNAs. We provide experimental details on the reagents and applications for multiplex conditional mutagenesis in zebrafish.
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Affiliation(s)
- L Yin
- Vanderbilt University School of Medicine, Nashville, TN, United States
| | - L A Maddison
- Vanderbilt University School of Medicine, Nashville, TN, United States
| | - W Chen
- Vanderbilt University School of Medicine, Nashville, TN, United States
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335
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Abstract
Zebrafish (Danio rerio) is a unique model organism at the functional intersection between a high fecundity and conserved vertebrate physiology while being amenable to a multitude of genome editing techniques. The genome engineering field has experienced an unprecedented rate of growth in the recent years since the introduction of designer endonucleases, such as zinc finger nucleases, transcription activator-like effector nucleases, and clustered regularly interspaced short palindromic repeats-Cas9 systems. With the ever-evolving toolset available to the scientific community, the important question one should ask is not simply how to make a mutant line, but rather how best to do so. For this purpose, understanding the toolset is just one end of the equation; understanding how DNA is repaired once double-strand breaks are induced by designer endonucleases, as well as understanding proper fish handling and line maintenance techniques, are also essential to rapidly edit the zebrafish genome. This chapter is outlined to provide a bird's-eye view on each of these three components. The goal of this chapter is to facilitate the adoption of the zebrafish as a model to study human genetic disease and to rapidly analyze the function of the vertebrate genome.
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Affiliation(s)
- H Ata
- Mayo Clinic, Rochester, MN, United States
| | - K J Clark
- Mayo Clinic, Rochester, MN, United States
| | - S C Ekker
- Mayo Clinic, Rochester, MN, United States
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336
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Shigeta M, Sakane Y, Iida M, Suzuki M, Kashiwagi K, Kashiwagi A, Fujii S, Yamamoto T, Suzuki KIT. Rapid and efficient analysis of gene function using CRISPR-Cas9 inXenopus tropicalisfounders. Genes Cells 2016; 21:755-71. [DOI: 10.1111/gtc.12379] [Citation(s) in RCA: 24] [Impact Index Per Article: 3.0] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/28/2016] [Accepted: 04/22/2016] [Indexed: 12/26/2022]
Affiliation(s)
- Mitsuki Shigeta
- Department of Mathematical and Life Sciences; Hiroshima University; 1-3-1 Kagamiyama Higashi-Hiroshima Hiroshima 739-8526 Japan
| | - Yuto Sakane
- Department of Mathematical and Life Sciences; Hiroshima University; 1-3-1 Kagamiyama Higashi-Hiroshima Hiroshima 739-8526 Japan
| | - Midori Iida
- Department of Bioscience and Bioinformatics; Kyushu Institute of Technology; 680-4 Kawazu Iizuka Fukuoka 820-8502 Japan
| | - Miyuki Suzuki
- Department of Mathematical and Life Sciences; Hiroshima University; 1-3-1 Kagamiyama Higashi-Hiroshima Hiroshima 739-8526 Japan
| | - Keiko Kashiwagi
- Institute for Amphibian Biology; Graduate School of Science; Hiroshima University; 1-3-1 Kagamiyama Higashi-Hiroshima Hiroshima 739-8526 Japan
| | - Akihiko Kashiwagi
- Institute for Amphibian Biology; Graduate School of Science; Hiroshima University; 1-3-1 Kagamiyama Higashi-Hiroshima Hiroshima 739-8526 Japan
| | - Satoshi Fujii
- Department of Bioscience and Bioinformatics; Kyushu Institute of Technology; 680-4 Kawazu Iizuka Fukuoka 820-8502 Japan
| | - Takashi Yamamoto
- Department of Mathematical and Life Sciences; Hiroshima University; 1-3-1 Kagamiyama Higashi-Hiroshima Hiroshima 739-8526 Japan
| | - Ken-ichi T. Suzuki
- Department of Mathematical and Life Sciences; Hiroshima University; 1-3-1 Kagamiyama Higashi-Hiroshima Hiroshima 739-8526 Japan
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337
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Haeussler M, Concordet JP. Genome Editing with CRISPR-Cas9: Can It Get Any Better? J Genet Genomics 2016; 43:239-50. [PMID: 27210042 PMCID: PMC5708852 DOI: 10.1016/j.jgg.2016.04.008] [Citation(s) in RCA: 39] [Impact Index Per Article: 4.9] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/12/2016] [Revised: 04/13/2016] [Accepted: 04/23/2016] [Indexed: 12/26/2022]
Abstract
The CRISPR-Cas revolution is taking place in virtually all fields of life sciences. Harnessing DNA cleavage with the CRISPR-Cas9 system of Streptococcus pyogenes has proven to be extraordinarily simple and efficient, relying only on the design of a synthetic single guide RNA (sgRNA) and its co-expression with Cas9. Here, we review the progress in the design of sgRNA from the original dual RNA guide for S. pyogenes and Staphylococcus aureus Cas9 (SpCas9 and SaCas9). New assays for genome-wide identification of off-targets have provided important insights into the issue of cleavage specificity in vivo. At the same time, the on-target activity of thousands of guides has been determined. These data have led to numerous online tools that facilitate the selection of guide RNAs in target sequences. It appears that for most basic research applications, cleavage activity can be maximized and off-targets minimized by carefully choosing guide RNAs based on computational predictions. Moreover, recent studies of Cas proteins have further improved the flexibility and precision of the CRISPR-Cas toolkit for genome editing. Inspired by the crystal structure of the complex of sgRNA-SpCas9 bound to target DNA, several variants of SpCas9 have recently been engineered, either with novel protospacer adjacent motifs (PAMs) or with drastically reduced off-targets. Novel Cas9 and Cas9-like proteins called Cpf1 have also been characterized from other bacteria and will benefit from the insights obtained from SpCas9. Genome editing with CRISPR-Cas9 may also progress with better understanding and control of cellular DNA repair pathways activated after Cas9-induced DNA cleavage.
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Affiliation(s)
- Maximilian Haeussler
- Santa Cruz Genomics Institute, MS CBSE, 1156 High Street, University of California, Santa Cruz, CA 95064, USA
| | - Jean-Paul Concordet
- Laboratoire Structure et Instabilité des Génomes, Inserm U1154, CNRS UMR7196, Muséum national d'Histoire naturelle, 43 rue Cuvier, 75005 Paris, France.
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338
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Bolukbasi MF, Gupta A, Wolfe SA. Creating and evaluating accurate CRISPR-Cas9 scalpels for genomic surgery. Nat Methods 2016; 13:41-50. [PMID: 26716561 DOI: 10.1038/nmeth.3684] [Citation(s) in RCA: 80] [Impact Index Per Article: 10.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/02/2015] [Accepted: 10/05/2015] [Indexed: 12/12/2022]
Abstract
The simplicity of site-specific genome targeting by type II clustered, regularly interspaced, short palindromic repeat (CRISPR)-Cas9 nucleases, along with their robust activity profile, has changed the landscape of genome editing. These favorable properties have made the CRISPR-Cas9 system the technology of choice for sequence-specific modifications in vertebrate systems. For many applications, whether the focus is on basic science investigations or therapeutic efficacy, activity and precision are important considerations when one is choosing a nuclease platform, target site and delivery method. Here we review recent methods for increasing the activity and accuracy of Cas9 and assessing the extent of off-target cleavage events.
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Affiliation(s)
- Mehmet Fatih Bolukbasi
- Department of Molecular, Cell and Cancer Biology, University of Massachusetts Medical School, Worcester, Massachusetts, USA.,Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, Massachusetts, USA
| | - Ankit Gupta
- Department of Molecular, Cell and Cancer Biology, University of Massachusetts Medical School, Worcester, Massachusetts, USA
| | - Scot A Wolfe
- Department of Molecular, Cell and Cancer Biology, University of Massachusetts Medical School, Worcester, Massachusetts, USA.,Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, Massachusetts, USA
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339
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Brown DR, Samsa LA, Qian L, Liu J. Advances in the Study of Heart Development and Disease Using Zebrafish. J Cardiovasc Dev Dis 2016; 3. [PMID: 27335817 PMCID: PMC4913704 DOI: 10.3390/jcdd3020013] [Citation(s) in RCA: 66] [Impact Index Per Article: 8.3] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 02/06/2023] Open
Abstract
Animal models of cardiovascular disease are key players in the translational medicine pipeline used to define the conserved genetic and molecular basis of disease. Congenital heart diseases (CHDs) are the most common type of human birth defect and feature structural abnormalities that arise during cardiac development and maturation. The zebrafish, Danio rerio, is a valuable vertebrate model organism, offering advantages over traditional mammalian models. These advantages include the rapid, stereotyped and external development of transparent embryos produced in large numbers from inexpensively housed adults, vast capacity for genetic manipulation, and amenability to high-throughput screening. With the help of modern genetics and a sequenced genome, zebrafish have led to insights in cardiovascular diseases ranging from CHDs to arrhythmia and cardiomyopathy. Here, we discuss the utility of zebrafish as a model system and summarize zebrafish cardiac morphogenesis with emphasis on parallels to human heart diseases. Additionally, we discuss the specific tools and experimental platforms utilized in the zebrafish model including forward screens, functional characterization of candidate genes, and high throughput applications.
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Affiliation(s)
- Daniel R. Brown
- Department of Pathology and Laboratory Medicine, University of North Carolina at Chapel Hill, Chapel Hill, NC 27599, USA; (D.R.B.); (L.Q.)
- McAllister Heart Institute, University of North Carolina at Chapel Hill, Chapel Hill, NC 27599, USA
| | - Leigh Ann Samsa
- Department of Cell Biology and Physiology; University of North Carolina at Chapel Hill, Chapel Hill, NC 27599, USA;
- McAllister Heart Institute, University of North Carolina at Chapel Hill, Chapel Hill, NC 27599, USA
| | - Li Qian
- Department of Pathology and Laboratory Medicine, University of North Carolina at Chapel Hill, Chapel Hill, NC 27599, USA; (D.R.B.); (L.Q.)
- McAllister Heart Institute, University of North Carolina at Chapel Hill, Chapel Hill, NC 27599, USA
| | - Jiandong Liu
- Department of Pathology and Laboratory Medicine, University of North Carolina at Chapel Hill, Chapel Hill, NC 27599, USA; (D.R.B.); (L.Q.)
- McAllister Heart Institute, University of North Carolina at Chapel Hill, Chapel Hill, NC 27599, USA
- Correspondence: ; Tel.: +1-919-962-0326; Fax: +1-919- 843-2063
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340
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Brocal I, White RJ, Dooley CM, Carruthers SN, Clark R, Hall A, Busch-Nentwich EM, Stemple DL, Kettleborough RNW. Efficient identification of CRISPR/Cas9-induced insertions/deletions by direct germline screening in zebrafish. BMC Genomics 2016; 17:259. [PMID: 27009152 PMCID: PMC4806435 DOI: 10.1186/s12864-016-2563-z] [Citation(s) in RCA: 15] [Impact Index Per Article: 1.9] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/06/2016] [Accepted: 03/02/2016] [Indexed: 12/26/2022] Open
Abstract
BACKGROUND The CRISPR/Cas9 system is a prokaryotic immune system that infers resistance to foreign genetic material and is a sort of 'adaptive immunity'. It has been adapted to enable high throughput genome editing and has revolutionised the generation of targeted mutations. RESULTS We have developed a scalable analysis pipeline to identify CRISPR/Cas9 induced mutations in hundreds of samples using next generation sequencing (NGS) of amplicons. We have used this system to investigate the best way to screen mosaic Zebrafish founder individuals for germline transmission of induced mutations. Screening sperm samples from potential founders provides much better information on germline transmission rates and crucially the sequence of the particular insertions/deletions (indels) that will be transmitted. This enables us to combine screening with archiving to create a library of cryopreserved samples carrying known mutations. It also allows us to design efficient genotyping assays, making identifying F1 carriers straightforward. CONCLUSIONS The methods described will streamline the production of large numbers of knockout alleles in selected genes for phenotypic analysis, complementing existing efforts using random mutagenesis.
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Affiliation(s)
- Isabel Brocal
- />Wellcome Trust Sanger Institute, Wellcome Trust Genome Campus, Hinxton, Cambridge, CB10 1SA UK
| | - Richard J. White
- />Wellcome Trust Sanger Institute, Wellcome Trust Genome Campus, Hinxton, Cambridge, CB10 1SA UK
| | - Christopher M. Dooley
- />Wellcome Trust Sanger Institute, Wellcome Trust Genome Campus, Hinxton, Cambridge, CB10 1SA UK
| | - Samantha N. Carruthers
- />Wellcome Trust Sanger Institute, Wellcome Trust Genome Campus, Hinxton, Cambridge, CB10 1SA UK
| | - Richard Clark
- />Wellcome Trust Sanger Institute, Wellcome Trust Genome Campus, Hinxton, Cambridge, CB10 1SA UK
| | - Amanda Hall
- />Wellcome Trust Sanger Institute, Wellcome Trust Genome Campus, Hinxton, Cambridge, CB10 1SA UK
| | | | - Derek L. Stemple
- />Wellcome Trust Sanger Institute, Wellcome Trust Genome Campus, Hinxton, Cambridge, CB10 1SA UK
| | - Ross N. W. Kettleborough
- />Wellcome Trust Sanger Institute, Wellcome Trust Genome Campus, Hinxton, Cambridge, CB10 1SA UK
- />Twist Bioscience, 455 Mission Bay Boulevard South, San Francisco, CA 94158 United States
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341
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Abstract
The zebrafish model is the only available high-throughput vertebrate assessment system, and it is uniquely suited for studies of in vivo cell biology. A sequenced and annotated genome has revealed a large degree of evolutionary conservation in comparison to the human genome. Due to our shared evolutionary history, the anatomical and physiological features of fish are highly homologous to humans, which facilitates studies relevant to human health. In addition, zebrafish provide a very unique vertebrate data stream that allows researchers to anchor hypotheses at the biochemical, genetic, and cellular levels to observations at the structural, functional, and behavioral level in a high-throughput format. In this review, we will draw heavily from toxicological studies to highlight advances in zebrafish high-throughput systems. Breakthroughs in transgenic/reporter lines and methods for genetic manipulation, such as the CRISPR-Cas9 system, will be comprised of reports across diverse disciplines.
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Affiliation(s)
- Gloria R Garcia
- Oregon State University, Department of Environmental and Molecular Toxicology, Environmental Health Sciences Center, Corvallis, OR 97331, USA
| | - Pamela D Noyes
- Oregon State University, Department of Environmental and Molecular Toxicology, Environmental Health Sciences Center, Corvallis, OR 97331, USA
| | - Robert L Tanguay
- Oregon State University, Department of Environmental and Molecular Toxicology, Environmental Health Sciences Center, Corvallis, OR 97331, USA.
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342
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Juntti SA, Hilliard AT, Kent KR, Kumar A, Nguyen A, Jimenez MA, Loveland JL, Mourrain P, Fernald RD. A Neural Basis for Control of Cichlid Female Reproductive Behavior by Prostaglandin F2α. Curr Biol 2016; 26:943-9. [PMID: 26996507 DOI: 10.1016/j.cub.2016.01.067] [Citation(s) in RCA: 59] [Impact Index Per Article: 7.4] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/23/2015] [Revised: 01/22/2016] [Accepted: 01/27/2016] [Indexed: 01/08/2023]
Abstract
In most species, females time reproduction to coincide with fertility. Thus, identifying factors that signal fertility to the brain can provide access to neural circuits that control sexual behaviors. In vertebrates, levels of key signaling molecules rise at the time of fertility to prime the brain for reproductive behavior [1-11], but how and where they regulate neural circuits is not known [12, 13]. Specifically, 17α,20β-dihydroxyprogesterone (DHP) and prostaglandin F2α (PGF2α) levels rise in teleost fish around the time of ovulation [10, 14, 15]. In an African cichlid fish, Astatotilapia burtoni, fertile females select a mate and perform a stereotyped spawning routine, offering quantifiable behavioral outputs of neural circuits. We show that, within minutes, PGF2α injection activates a naturalistic pattern of sexual behavior in female A. burtoni. We also identify cells in the brain that transduce the prostaglandin signal to mate and show that the gonadal steroid DHP modulates mRNA levels of the putative receptor for PGF2α (Ptgfr). We use CRISPR/Cas9 to generate the first targeted gene mutation in A. burtoni and show that Ptgfr is necessary for the initiation of sexual behavior, uncoupling sexual behavior from reproductive status. Our findings are consistent with a model in which PGF2α communicates fertility status via Ptgfr to circuits in the brain that drive female sexual behavior. Our targeted genome modification in a cichlid fish shows that dissection of gene function can reveal basic control mechanisms for behaviors in this large family of species with diverse and fascinating social systems [16, 17].
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Affiliation(s)
- Scott A Juntti
- Department of Biology, Stanford University, Stanford, CA 94305, USA.
| | | | - Kai R Kent
- Department of Biology, Stanford University, Stanford, CA 94305, USA
| | - Anusha Kumar
- Department of Biology, Stanford University, Stanford, CA 94305, USA
| | - Andrew Nguyen
- Department of Biology, Stanford University, Stanford, CA 94305, USA
| | | | | | - Philippe Mourrain
- Department of Psychiatry and Behavioral Sciences, Center for Sleep Sciences, Stanford University, Stanford, CA 94305, USA
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343
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Ma AC, McNulty MS, Poshusta TL, Campbell JM, Martínez-Gálvez G, Argue DP, Lee HB, Urban MD, Bullard CE, Blackburn PR, Man TK, Clark KJ, Ekker SC. FusX: A Rapid One-Step Transcription Activator-Like Effector Assembly System for Genome Science. Hum Gene Ther 2016; 27:451-63. [PMID: 26854857 PMCID: PMC4931509 DOI: 10.1089/hum.2015.172] [Citation(s) in RCA: 41] [Impact Index Per Article: 5.1] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/28/2023] Open
Abstract
Transcription activator-like effectors (TALEs) are extremely effective, single-molecule DNA-targeting molecular cursors used for locus-specific genome science applications, including high-precision molecular medicine and other genome engineering applications. TALEs are used in genome engineering for locus-specific DNA editing and imaging, as artificial transcriptional activators and repressors, and for targeted epigenetic modification. TALEs as nucleases (TALENs) are effective editing tools and offer high binding specificity and fewer sequence constraints toward the targeted genome than other custom nuclease systems. One bottleneck of broader TALE use is reagent accessibility. For example, one commonly deployed method uses a multitube, 5-day assembly protocol. Here we describe FusX, a streamlined Golden Gate TALE assembly system that (1) is backward compatible with popular TALE backbones, (2) is functionalized as a single-tube 3-day TALE assembly process, (3) requires only commonly used basic molecular biology reagents, and (4) is cost-effective. More than 100 TALEN pairs have been successfully assembled using FusX, and 27 pairs were quantitatively tested in zebrafish, with each showing high somatic and germline activity. Furthermore, this assembly system is flexible and is compatible with standard molecular biology laboratory tools, but can be scaled with automated laboratory support. To demonstrate, we use a highly accessible and commercially available liquid-handling robot to rapidly and accurately assemble TALEs using the FusX TALE toolkit. Together, the FusX system accelerates TALE-based genomic science applications from basic science screening work for functional genomics testing and molecular medicine applications.
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Affiliation(s)
- Alvin C Ma
- 1 Department of Biochemistry and Molecular Biology, Mayo Clinic , Rochester, Minnesota.,2 Department of Medicine, Li Ka Shing Faculty of Medicine, University of Hong Kong , Hong Kong
| | - Melissa S McNulty
- 1 Department of Biochemistry and Molecular Biology, Mayo Clinic , Rochester, Minnesota
| | - Tanya L Poshusta
- 1 Department of Biochemistry and Molecular Biology, Mayo Clinic , Rochester, Minnesota
| | - Jarryd M Campbell
- 1 Department of Biochemistry and Molecular Biology, Mayo Clinic , Rochester, Minnesota
| | | | - David P Argue
- 1 Department of Biochemistry and Molecular Biology, Mayo Clinic , Rochester, Minnesota
| | - Han B Lee
- 1 Department of Biochemistry and Molecular Biology, Mayo Clinic , Rochester, Minnesota
| | - Mark D Urban
- 1 Department of Biochemistry and Molecular Biology, Mayo Clinic , Rochester, Minnesota
| | - Cassandra E Bullard
- 1 Department of Biochemistry and Molecular Biology, Mayo Clinic , Rochester, Minnesota
| | - Patrick R Blackburn
- 1 Department of Biochemistry and Molecular Biology, Mayo Clinic , Rochester, Minnesota
| | - Toni K Man
- 2 Department of Medicine, Li Ka Shing Faculty of Medicine, University of Hong Kong , Hong Kong
| | - Karl J Clark
- 1 Department of Biochemistry and Molecular Biology, Mayo Clinic , Rochester, Minnesota
| | - Stephen C Ekker
- 1 Department of Biochemistry and Molecular Biology, Mayo Clinic , Rochester, Minnesota
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344
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Experimental approaches to studying the nature and impact of splicing variation in zebrafish. Methods Cell Biol 2016; 135:259-88. [PMID: 27443930 DOI: 10.1016/bs.mcb.2016.02.006] [Citation(s) in RCA: 2] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/30/2022]
Abstract
From a fixed number of genes carried in all cells, organisms create considerable diversity in cellular phenotype through differential regulation of gene expression. One prevalent source of transcriptome diversity is alternative pre-mRNA splicing, which is manifested in many different forms. Zebrafish models of splicing dysfunction due to mutated spliceosome components provide opportunity to link biochemical analyses of spliceosome structure and function with whole organism phenotypic outcomes. Drawing from experience with two zebrafish mutants: cephalophŏnus (a prpf8 mutant, isolated for defects in granulopoiesis) and caliban (a rnpc3 mutant, isolated for defects in digestive organ development), we describe the use of glycerol gradient sedimentation and native gel electrophoresis to resolve components of aberrant splicing complexes. We also describe how RNAseq can be employed to examine relatively rare alternative splicing events including intron retention. Such experimental approaches in zebrafish can promote understanding of how splicing variation and dysfunction contribute to phenotypic diversity and disease pathogenesis.
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345
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Chávez MN, Aedo G, Fierro FA, Allende ML, Egaña JT. Zebrafish as an Emerging Model Organism to Study Angiogenesis in Development and Regeneration. Front Physiol 2016; 7:56. [PMID: 27014075 PMCID: PMC4781882 DOI: 10.3389/fphys.2016.00056] [Citation(s) in RCA: 78] [Impact Index Per Article: 9.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/25/2015] [Accepted: 02/05/2016] [Indexed: 01/04/2023] Open
Abstract
Angiogenesis is the process through which new blood vessels are formed from preexisting ones and plays a critical role in several conditions including embryonic development, tissue repair and disease. Moreover, enhanced therapeutic angiogenesis is a major goal in the field of regenerative medicine and efficient vascularization of artificial tissues and organs is one of the main hindrances in the implementation of tissue engineering approaches, while, on the other hand, inhibition of angiogenesis is a key therapeutic target to inhibit for instance tumor growth. During the last decades, the understanding of cellular and molecular mechanisms involved in this process has been matter of intense research. In this regard, several in vitro and in vivo models have been established to visualize and study migration of endothelial progenitor cells, formation of endothelial tubules and the generation of new vascular networks, while assessing the conditions and treatments that either promote or inhibit such processes. In this review, we address and compare the most commonly used experimental models to study angiogenesis in vitro and in vivo. In particular, we focus on the implementation of the zebrafish (Danio rerio) as a model to study angiogenesis and discuss the advantages and not yet explored possibilities of its use as model organism.
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Affiliation(s)
- Myra N Chávez
- Department of Plastic Surgery and Hand Surgery, University Hospital rechts der Isar, Technische Universität MünchenMunich, Germany; Department of Biology, FONDAP Center for Genome Regulation, Faculty of Science, Universidad de ChileSantiago, Chile; Department of Biochemistry and Molecular Biology, FONDAP Advanced Center for Chronic Diseases (ACCDiS) and Center for Molecular Studies of the Cell (CEMC), Faculty of Chemical and Pharmaceutical Sciences, Faculty of Medicine, University of ChileSantiago, Chile
| | - Geraldine Aedo
- Department of Biology, FONDAP Center for Genome Regulation, Faculty of Science, Universidad de Chile Santiago, Chile
| | - Fernando A Fierro
- Department of Cell Biology and Human Anatomy, University of California Davis, Sacramento, CA, USA
| | - Miguel L Allende
- Department of Biology, FONDAP Center for Genome Regulation, Faculty of Science, Universidad de Chile Santiago, Chile
| | - José T Egaña
- Institute for Medical and Biological Engineering, Schools of Engineering, Biological Sciences and Medicine, Pontifícia Universidad Católica de Chile Santiago, Chile
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346
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Abstract
In the postgenomic era, the ability to quickly, efficiently, and inexpensively assign function to the zebrafish proteome is critical. Clustered regularly interspaced short palindromic repeats (CRISPRs) have revolutionized the ability to perform reverse genetics because of its simplicity and broad applicability. The CRISPR system is composed of an engineered, gene-specific single guide RNA (sgRNA) and a Cas9 enzyme that causes double-stranded breaks in DNA at the targeted site. This simple, two-part system, when injected into one-cell stage zebrafish embryos, efficiently mutates target loci at a frequency such that injected embryos phenocopy known mutant phenotypes. This property allows for CRISPR-based F0 screening in zebrafish, which provides a means to screen through a large number of candidate genes for their role in a phenotype of interest. While there are important considerations for any successful genetic screen, CRISPR screening has significant benefits over conventional methods and can be accomplished in any lab with modest molecular biology experience.
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Affiliation(s)
- A N Shah
- Fred Hutchinson Cancer Research Center, Seattle, WA, United States
| | - C B Moens
- Fred Hutchinson Cancer Research Center, Seattle, WA, United States
| | - A C Miller
- University of Oregon, Eugene, OR, United States
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347
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Pereira TCB, Campos MM, Bogo MR. Copper toxicology, oxidative stress and inflammation using zebrafish as experimental model. J Appl Toxicol 2016; 36:876-85. [PMID: 26888422 DOI: 10.1002/jat.3303] [Citation(s) in RCA: 140] [Impact Index Per Article: 17.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/16/2015] [Revised: 12/17/2015] [Accepted: 01/12/2016] [Indexed: 12/26/2022]
Abstract
Copper is an essential micronutrient and a key catalytic cofactor in a wide range of enzymes. As a trace element, copper levels are tightly regulated and both its deficit and excess are deleterious to the organism. Under inflammatory conditions, serum copper levels are increased and trigger oxidative stress responses that activate inflammatory responses. Interestingly, copper dyshomeostasis, oxidative stress and inflammation are commonly present in several chronic diseases. Copper exposure can be easily modeled in zebrafish; a consolidated model in toxicology with increasing interest in immunity-related research. As a result of developmental, economical and genetic advantages, this freshwater teleost is uniquely suitable for chemical and genetic large-scale screenings, representing a powerful experimental tool for a whole-organism approach, mechanistic studies, disease modeling and beyond. Copper toxicological and more recently pro-inflammatory effects have been investigated in both larval and adult zebrafish with breakthrough findings. Here, we provide an overview of copper metabolism in health and disease and its effects on oxidative stress and inflammation responses in zebrafish models. Copper-induced inflammation is highlighted owing to its potential to easily mimic pro-oxidative and pro-inflammatory features that combined with zebrafish genetic tractability could help further in the understanding of copper metabolism, inflammatory responses and related diseases. Copyright © 2016 John Wiley & Sons, Ltd.
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Affiliation(s)
- Talita Carneiro Brandão Pereira
- Programa de Pós-Graduação em Medicina e Ciências da Saúde, PUCRS, Porto Alegre, Brasil.,Laboratório de Biologia Genômica e Molecular, PUCRS, Porto Alegre, Brasil
| | - Maria Martha Campos
- Programa de Pós-Graduação em Medicina e Ciências da Saúde, PUCRS, Porto Alegre, Brasil.,Instituto de Toxicologia e Farmacologia, PUCRS, Porto Alegre, Brasil.,Programa de Pós-Graduação em Odontologia, PUCRS, Porto Alegre, Brasil
| | - Maurício Reis Bogo
- Programa de Pós-Graduação em Medicina e Ciências da Saúde, PUCRS, Porto Alegre, Brasil.,Laboratório de Biologia Genômica e Molecular, PUCRS, Porto Alegre, Brasil.,Instituto de Toxicologia e Farmacologia, PUCRS, Porto Alegre, Brasil.,Programa de Pós-Graduação em Biologia Celular e Molecular, PUCRS, Porto Alegre, Brasil
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348
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Raitskin O, Patron NJ. Multi-gene engineering in plants with RNA-guided Cas9 nuclease. Curr Opin Biotechnol 2016; 37:69-75. [PMID: 26707469 DOI: 10.1016/j.copbio.2015.11.008] [Citation(s) in RCA: 25] [Impact Index Per Article: 3.1] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/07/2015] [Revised: 10/26/2015] [Accepted: 11/02/2015] [Indexed: 11/17/2022]
Abstract
The use of RNA-guided Cas9 endonuclease for the concurrent engineering of multiple genes has been demonstrated in a number of plant species. Although Cas9 is a large monomeric protein, the single guide RNA (sgRNA) that directs it to a specific DNA target sequence is small and easy to reprogram. It is therefore relatively simple to produce numerous sgRNAs to target multiple endogenous sequences. Several approaches to express multiple sgRNAs and Cas9 in plants for the purpose of simultaneous editing or transcriptional regulation of many genes have recently been reported.
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Affiliation(s)
- Oleg Raitskin
- The Sainsbury Laboratory, Norwich Research Park, Norwich NR4 7UH, UK
| | - Nicola J Patron
- The Sainsbury Laboratory, Norwich Research Park, Norwich NR4 7UH, UK.
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349
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Van Otterloo E, Williams T, Artinger KB. The old and new face of craniofacial research: How animal models inform human craniofacial genetic and clinical data. Dev Biol 2016; 415:171-187. [PMID: 26808208 DOI: 10.1016/j.ydbio.2016.01.017] [Citation(s) in RCA: 44] [Impact Index Per Article: 5.5] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/09/2015] [Revised: 01/16/2016] [Accepted: 01/21/2016] [Indexed: 12/31/2022]
Abstract
The craniofacial skeletal structures that comprise the human head develop from multiple tissues that converge to form the bones and cartilage of the face. Because of their complex development and morphogenesis, many human birth defects arise due to disruptions in these cellular populations. Thus, determining how these structures normally develop is vital if we are to gain a deeper understanding of craniofacial birth defects and devise treatment and prevention options. In this review, we will focus on how animal model systems have been used historically and in an ongoing context to enhance our understanding of human craniofacial development. We do this by first highlighting "animal to man" approaches; that is, how animal models are being utilized to understand fundamental mechanisms of craniofacial development. We discuss emerging technologies, including high throughput sequencing and genome editing, and new animal repository resources, and how their application can revolutionize the future of animal models in craniofacial research. Secondly, we highlight "man to animal" approaches, including the current use of animal models to test the function of candidate human disease variants. Specifically, we outline a common workflow deployed after discovery of a potentially disease causing variant based on a select set of recent examples in which human mutations are investigated in vivo using animal models. Collectively, these topics will provide a pipeline for the use of animal models in understanding human craniofacial development and disease for clinical geneticist and basic researchers alike.
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Affiliation(s)
- Eric Van Otterloo
- Department of Craniofacial Biology, School of Dental Medicine, University of Colorado Anschutz Medical Campus, Aurora, CO 80045, USA.
| | - Trevor Williams
- Department of Craniofacial Biology, School of Dental Medicine, University of Colorado Anschutz Medical Campus, Aurora, CO 80045, USA
| | - Kristin Bruk Artinger
- Department of Craniofacial Biology, School of Dental Medicine, University of Colorado Anschutz Medical Campus, Aurora, CO 80045, USA.
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350
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Varshney GK, Zhang S, Pei W, Adomako-Ankomah A, Fohtung J, Schaffer K, Carrington B, Maskeri A, Slevin C, Wolfsberg T, Ledin J, Sood R, Burgess SM. CRISPRz: a database of zebrafish validated sgRNAs. Nucleic Acids Res 2016; 44:D822-6. [PMID: 26438539 PMCID: PMC4702947 DOI: 10.1093/nar/gkv998] [Citation(s) in RCA: 36] [Impact Index Per Article: 4.5] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/02/2015] [Accepted: 09/22/2015] [Indexed: 12/26/2022] Open
Abstract
CRISPRz (http://research.nhgri.nih.gov/CRISPRz/) is a database of CRISPR/Cas9 target sequences that have been experimentally validated in zebrafish. Programmable RNA-guided CRISPR/Cas9 has recently emerged as a simple and efficient genome editing method in various cell types and organisms, including zebrafish. Because the technique is so easy and efficient in zebrafish, the most valuable asset is no longer a mutated fish (which has distribution challenges), but rather a CRISPR/Cas9 target sequence to the gene confirmed to have high mutagenic efficiency. With a highly active CRISPR target, a mutant fish can be quickly replicated in any genetic background anywhere in the world. However, sgRNA's vary widely in their activity and models for predicting target activity are imperfect. Thus, it is very useful to collect in one place validated CRISPR target sequences with their relative mutagenic activities. A researcher could then select a target of interest in the database with an expected activity. Here, we report the development of CRISPRz, a database of validated zebrafish CRISPR target sites collected from published sources, as well as from our own in-house large-scale mutagenesis project. CRISPRz can be searched using multiple inputs such as ZFIN IDs, accession number, UniGene ID, or gene symbols from zebrafish, human and mouse.
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Affiliation(s)
- Gaurav K Varshney
- Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD 20892, USA
| | - Suiyuan Zhang
- Computational and Statistical Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD 20892, USA
| | - Wuhong Pei
- Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD 20892, USA
| | - Ashrifia Adomako-Ankomah
- Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD 20892, USA
| | - Jacob Fohtung
- Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD 20892, USA
| | - Katherine Schaffer
- Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD 20892, USA
| | - Blake Carrington
- Zebrafish Core, Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD 20892, USA
| | - Anoo Maskeri
- Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD 20892, USA
| | - Claire Slevin
- Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD 20892, USA
| | - Tyra Wolfsberg
- Computational and Statistical Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD 20892, USA
| | - Johan Ledin
- Department of Organismal Biology, Science for Life Laboratory, Uppsala University, SE-752 36 Uppsala, Sweden
| | - Raman Sood
- Zebrafish Core, Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD 20892, USA
| | - Shawn M Burgess
- Translational and Functional Genomics Branch, National Human Genome Research Institute, National Institutes of Health, Bethesda, MD 20892, USA
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