Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development of a customizable platform to rapidly generate gene modifications in a wide variety of organisms, including zebrafish. CRISPR-based genome editing uses a single guide RNA (sgRNA) to target a CRISPR-associated (Cas) endonuclease to a genomic DNA (gDNA) target of interest, where the Cas endonuclease generates a double-strand break (DSB). Repair of DSBs by error-prone mechanisms lead to insertions and/or deletions (indels). This can cause frameshift mutations that often introduce a premature stop codon within the coding sequence, thus creating a protein-null allele. CRISPR-based genome engineering requires only a few molecular components and is easily introduced into zebrafish embryos by microinjection. This protocol describes the methods used to generate CRISPR reagents for zebrafish microinjection and to identify fish exhibiting germline transmission of CRISPR-modified genes. These methods include in vitro transcription of sgRNAs, microinjection of CRISPR reagents, identification of indels induced at the target site using a PCR-based method called a heteroduplex mobility assay (HMA), and characterization of the indels using both a low throughput and a powerful next-generation sequencing (NGS)-based approach that can analyze multiple PCR products collected from heterozygous fish. This protocol is streamlined to minimize both the number of fish required and the types of equipment needed to perform the analyses. Furthermore, this protocol is designed to be amenable for use by laboratory personal of all levels of experience including undergraduates, enabling this powerful tool to be economically employed by any research group interested in performing CRISPR-based genomic modification in zebrafish.
Gene expression and neurogenesis in vertebrates involves a wide variety of cellular machinery to coordinate the transcriptional changes that direct cellular differentiation. Critical aspects of this epigenetic machinery are chromatin remodeling factors. One such factor is CHD5, a vertebrate‐specific member of a family of ATP‐dependent chromatin remodeling proteins. CHD5 is expressed primarily in neural tissue where it is thought to contribute to neurogenesis and has been strongly linked to tumor suppression. Currently, little is known regarding the molecular mechanisms by which CHD5 contributes to neurogenesis or tumor suppression. We hypothesize that loss of CHD5 contributes to development of neuroblastoma because neural precursor cells lacking CHD5 fail to alter some aspect of their neural precursor transcriptome/epigenome. To study the biochemical properties and functional importance of CHD5 remodelers, we have developed a zebrafish model to study Chd5, the zebrafish homolog of human CHD5. Using the CRISPR/Cas9 system, we have engineered chd5 knockout zebrafish embryos. These zebrafish did not display any overt developmental phenotypes, however next‐generation sequencing of brain tissue from zebrafish embryos reveals that loss of chd5 is associated with broad changes in gene expression. These genes are both up and down regulated, suggesting Chd5 is necessary for both activation and repression of genes during brain development in zebrafish. Analysis of these genes and of their possible contribution to Chd5‐dependent phenotypes is underway. Support or Funding Information Purdue Center for Cancer Research; Purdue College of Agriculture
CHD5 is a vertebrate specific member of a family of ATP‐dependent chromatin remodeling proteins. Chromodomain/Helicase/DNA‐binding (CHD) proteins play a variety of roles in vertebrate development, and loss of CHD5 causes failure of proper expression in a cohort of genes involved in neural development. CHD5 plays a critical role tumor suppression in humans, and loss of CHD5 contributes to the formation or progression of numerous cancers, in particular neuroblastoma. To understand how CHD5 plays a role in tumor suppression, it is necessary to understand how CHD5 contributes to normal development and how its normal functions are misregulated during tumor progression. Our lab is establishing zebrafish as model system in which to study the role of chd5 dependent processes.We have engineered chd5 knockout fish using CRISPR‐Cas9 technology. The resulting fish do not exhibit any overt developmental phenotypes, and do not produce tumors during adulthood (N=165, aged >16 months). We hypothesize that the closely related remodelers CHD3/4 function redundantly with CHD5, and propose to use a novel knock‐in approach to generate zebrafish carrying a dominant negative mutation in the ATPase domain that has previously been shown to result in strong mutant phenotypes when generated in closely related remodelers.To engineer a dominant negative allele of chd5 we have engineered a homologous recombination (HR) strategy that uses piggyBac transposase technology, which has previously been used in Drosophila, mice, and human cells to engineer scarless genomic modifications. This design uses two‐steps to first, introduce a homologous donor carrying the mutation of interest and a fluorescent marker for easy identification. Second, the HR positive fish will be bred with piggyBac expressing fish lines to seamlessly excise the exogenous sequences, thus leaving only the desired genome modification.Once we have engineered this method into the fish, we will be able to use piggyBac to control when and where we express the dominant negative allele of chd5 so that we can score the resulting fish for developmental and tumor phenotypes. The screening of the dominant negative, knock‐in lines is in progress. The resulting dominant negative lines will give great insight into the role of chd5 in neural development and tumor suppression.Support or Funding InformationFunding made available by: Purdue University Center for Cancer Research, Purdue University Department of Biochemistry, Purdue University College of AgricultureThis abstract is from the Experimental Biology 2019 Meeting. There is no full text article associated with this abstract published in The FASEB Journal.
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