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      Characterizing Sources of Variability in Zebrafish Embryo Screening Protocols

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          Abstract

          There is a need for fast, efficient, and cost-effective hazard identification and characterization of chemical hazards. This need is generating increased interest in the use of zebrafish embryos as both a screening tool and an alternative to mammalian test methods. A Collaborative Workshop on Aquatic Models and 21st Century Toxicology identified the lack of appropriate and consistent testing protocols as a challenge to the broader application of the zebrafish embryo model. The National Toxicology Program established the Systematic Evaluation of the Application of Zebrafish in Toxicology (SEAZIT) initiative to address the lack of consistent testing guidelines and identify sources of variability for zebrafish-based assays. This report summarizes initial SEAZIT information-gathering efforts. Investigators in academic, government, and industry laboratories that routinely use zebrafish embryos for chemical toxicity testing were asked about their husbandry practices and standard protocols. Information was collected about protocol components including zebrafish strains, feed, system water, disease surveillance, embryo exposure conditions, and endpoints. Literature was reviewed to assess issues raised by the investigators. Interviews revealed substantial variability across design parameters, data collected, and analysis procedures. The presence of the chorion and renewal of exposure media (static versus static-renewal) were identified as design parameters that could potentially influence study outcomes and should be investigated further with studies to determine chemical uptake from treatment solution into embryos. The information gathered in this effort provides a basis for future SEAZIT activities to promote more consistent practices among researchers using zebrafish embryos for toxicity evaluation.

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          Most cited references149

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          Stages of embryonic development of the zebrafish.

          We describe a series of stages for development of the embryo of the zebrafish, Danio (Brachydanio) rerio. We define seven broad periods of embryogenesis--the zygote, cleavage, blastula, gastrula, segmentation, pharyngula, and hatching periods. These divisions highlight the changing spectrum of major developmental processes that occur during the first 3 days after fertilization, and we review some of what is known about morphogenesis and other significant events that occur during each of the periods. Stages subdivide the periods. Stages are named, not numbered as in most other series, providing for flexibility and continued evolution of the staging series as we learn more about development in this species. The stages, and their names, are based on morphological features, generally readily identified by examination of the live embryo with the dissecting stereomicroscope. The descriptions also fully utilize the optical transparancy of the live embryo, which provides for visibility of even very deep structures when the embryo is examined with the compound microscope and Nomarski interference contrast illumination. Photomicrographs and composite camera lucida line drawings characterize the stages pictorially. Other figures chart the development of distinctive characters used as staging aid signposts.
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            The identification of genes with unique and essential functions in the development of the zebrafish, Danio rerio.

            In a large-scale screen, we isolated mutants displaying a specific visible phenotype in embryos or early larvae of the zebrafish, Danio rerio. Males were mutagenized with ethylnitrosourea (ENU) and F2 families of single pair matings between sibling F1 fish, heterozygous for a mutagenized genome, were raised. Egg lays were obtained from several crosses between F2 siblings, resulting in scoring of 3857 mutagenized genomes. F3 progeny were scored at the second, third and sixth day of development, using a stereomicroscope. In a subsequent screen, fixed embryos were analyzed for correct retinotectal projection. A total of 4264 mutants were identified. Two thirds of the mutants displaying rather general abnormalities were eventually discarded. We kept and characterized 1163 mutants. In complementation crosses performed between mutants with similar phenotypes, 894 mutants have been assigned to 372 genes. The average allele frequency is 2.4. We identified genes involved in early development, notochord, brain, spinal cord, somites, muscles, heart, circulation, blood, skin, fin, eye, otic vesicle, jaw and branchial arches, pigment pattern, pigment formation, gut, liver, motility and touch response. Our collection contains alleles of almost all previously described zebrafish mutants. From the allele frequencies and other considerations we estimate that the 372 genes defined by the mutants probably represent more than half of all genes that could have been discovered using the criteria of our screen. Here we give an overview of the spectrum of mutant phenotypes obtained, and discuss the limits and the potentials of a genetic saturation screen in the zebrafish.
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              Production of clones of homozygous diploid zebra fish (Brachydanio rerio).

              Homozygous diploid zebra fish have been produced on a large scale by the application of simple physical treatments. Clones of homozygous fish have been produced from individual homozygotes. These clones and associated genetic methods will facilitate genetic analyses of this vertebrate.
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                Author and article information

                Journal
                100953980
                21906
                ALTEX
                ALTEX
                ALTEX
                1868-596X
                1868-8551
                6 August 2023
                2019
                10 November 2018
                14 August 2023
                : 36
                : 1
                : 103-120
                Affiliations
                [1 ]Integrated Laboratory Systems, Research Triangle Park, North Carolina, USA
                [2 ]Division of the National Toxicology Program, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, North Carolina, USA
                [3 ]National Toxicology Program Interagency Center for the Evaluation of Alternative Toxicological Methods, Division of the National Toxicology Program, National Institute of Environmental Health Sciences, National Institutes of Health, Research Triangle Park, North Carolina, USA
                [4 ]Battelle, Life Sciences Research, Columbus, Ohio, USA
                [5 ]Integrated Systems Toxicology Division, National Health and Environmental Effects Research Laboratory, Office of Research and Development, U.S. Environmental Protection Agency, Research Triangle Park, North Carolina, USA
                [6 ]United States Army Engineer Research and Development Center, Vicksburg, Mississippi, USA
                [7 ]Department of Biological Sciences and Center for Human Health and the Environment, North Carolina State University, Raleigh, North Carolina, USA
                [8 ]Pfizer Pharmaceuticals, New London/Norwich, Connecticut, USA
                [9 ]Department of Environmental & Molecular Toxicology, Oregon State University, Corvallis, Oregon, USA
                [10 ]Department of Environmental Sciences, University of California, Riverside, California, USA
                Author notes
                Correspondence: Jon Hamm, PhD, ILS, P.O. Box 13501, Research Triangle Park, NC 27709 USA, ( hammjt@ 123456niehs.nih.gov )
                Article
                NIHMS1920022
                10.14573/altex.1804162
                10424490
                30415271
                ffb91911-94c9-4a0e-8b3d-7e93f6418ed1

                This is an Open Access article distributed under the terms of the Creative Commons Attribution 4.0 International license ( http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution and reproduction in any medium, provided the original work is appropriately cited.

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