We thank Barbara Garbers, PhD, from Liwen Bianji (Edanz) for editing the English text of a draft of this manuscript. This work was supported by the Key-Area Research and Development Program of Guangdong Province (2019B030335001), the National Key R&#38;D Program of China (2019YFE0106700), the National Natural Science Foundation of China (32070819, 31970782), and the Research Fund Program of Guangdong Provincial Key Lab of Pathogenic Biology and Epidemiology for Aquatic Economic Animals (PBEA2020YB05).

This study was conducted in strict compliance with the guidelines of the Care and Use Committee of the South China Normal University.
1. Preparing synthetically modified gRNA and zSpRY-ABE8e mRNA
<ol>
	<li>Design the gRNA according to previous publications37,38.
	<ol>
		<li>Preferentially select NRN as the PAM sequence, as zSpRY-ABE8e shows a higher preference for NRN PAM than NYN PAM (where R is A or G, and Y is C or T)18. Ensure that the target adenine nucleotide is in the highly active editing window (third to ninth nucleotide along the protospacer).</li>
	</ol>
	</li>
	<li>Order EasyEdit sgRNA (EE gRNA) modified with 2&#8242;-O-methyl-3&#8242;-phosphorothioate (MS) at both ends (Figure 1). 
	NOTE: The EE gRNA should be dissolved in RNase-free water and stored at &#8722;80 &#176;C.</li>
	<li>Linearize the zSpRY-ABE8e plasmid18.
	<ol>
		<li>Linearize 6 &#181;g of the plasmid with the XbaI enzyme (Table of Materials) in a metal bath at 37 &#176;C for 3 h.</li>
		<li>Purify the linearized plasmid using the DNA purification kit (see Table of Materials) according to the manufacturer&#39;s instructions.</li>
	</ol>
	</li>
	<li>Transcribe the mRNA using the in vitro transcription kit (Table of Materials).
	<ol>
		<li>Transcribe the zSpRY-ABE8e mRNA with the linearized plasmid as template according to the manufacturer&#39;s instructions. 
		NOTE: All operations involving mRNA and gRNA require the use of RNase-free laboratory supplies.</li>
	</ol>
	</li>
	<li>Use the RNA purification kit (Table of Materials) to purify the mRNA according to the manufacturer&#39;s instructions. 
	NOTE: Before eluting, the ethanol must be dried to avoid cytotoxicity, and the mRNA should be stored at &#8722;80 &#176;C if not used immediately.</li>
</ol>
2. Preparing microinjection glass capillaries
<ol>
	<li>Set up the micropipette puller system, and preheat for at least 30 min.</li>
	<li>Run a ramp test to determine the heat value needed to melt the borosilicate glass capillaries (with 1 mm outer diameter and 0.58 mm inner diameter) according to the manufacturer&#39;s instructions.</li>
	<li>Select a program, and click on the number on the keyboard to enter the parameter values. Set the following parameters to pull the glass capillaries: heat = ramp test value, pull = 50, velocity = 100, time = 50, and pressure = 30.</li>
	<li>Install the capillary tube, ensuring it is in the groove. Press PULL START/STOP to run the program.</li>
	<li>Preserve the pulled capillaries by inserting them into foam that contains gaps, keeping the needle suspended.</li>
</ol>
3. Microinjection of the zSpRY-ABE8e mRNA and EE gRNA mixture into zebrafish embryos
<ol>
	<li>Set up breeding tanks the night before the injection. Insert a divider, place two females and one male AB strain zebrafish (3-18 months old) in each tank, and cover with the lid. Refer to previous publications for gender distinctions39. 
	NOTE: To obtain more embryos, select fish that have not been breeding for at least 1 week.</li>
	<li>The next morning before the injection, thaw the zSpRY-ABE8e mRNA and EE gRNA on ice. Mix the mRNA and gRNA to final concentrations of 400 ng/&#181;L and 200 ng/&#181;L, respectively.
	<ol>
		<li>Perform the entire procedure on ice using RNase-free tips and tubes, and handle with gloves.</li>
	</ol>
	</li>
	<li>Configure the pneumatic microinjector. Close the partial pressure valve of the air pump and open the main valve first, unscrew the gas cylinder, and adjust the pressure of the partial pressure valve to 0.2 MPa/29 psi.
	<ol>
		<li>Turn on the pneumatic microinjector, set the mode to TIMER, and adjust the initial parameter pressure value to 20.0 psi and the TIMER value to 0.040 s.</li>
	</ol>
	</li>
	<li>Use fine forceps to carefully break the needles of the pulled glass capillaries under a stereomicroscope, making sure the needles are cut at an angle to facilitate the piercing of the chorion and the egg. 
	NOTE: The break point of the needle needs to be in the right place and be strong enough to pierce the egg yet thin enough to minimize damage to the egg.</li>
	<li>Add the mixture of zSpRY-ABE8e mRNA and gRNA to the tip of a glass capillary using a microloader pipette tip, avoiding bubbles in the capillary. Attach the needle to the micromanipulator, and configure the pressure and TIMER value of the injector to ensure 2 nL of mixture is produced per injection using a microcap.</li>
	<li>Unplug the dividers of the breeding tanks, and allow the fish to breed for 10-15 min. Collect the eggs using a strainer, and examine the cell stage and quality of the eggs under a stereomicroscope. Select the eggs in the single-cell state for injection to improve the editing efficiency.</li>
	<li>Line the eggs on a 1.5% agarose injection plate, and inject 2 nL of the mixture into the cell, not the yolk, of the embryos under a microscope to improve the editing efficiency. Retain at least 15 eggs as the control group.</li>
	<li>Use 1x E3 buffer to collect the eggs in Petri dishes, and place them in an incubator at 28 &#176;C. Remove the dead eggs, and change the culture buffer every 12 h until 48 hours post fertilization (hpf).</li>
</ol>
4. Efficiency analysis of base editing with EditR
<ol>
	<li>At 48 hpf, collect three pools of six randomly selected injected embryos and six randomly selected control embryos into PCR tubes respectively. Use a pipette to aspirate the residual E3 buffer.</li>
	<li>Add 40 &#181;L of 50 mM NaOH into the PCR tubes. Put the tubes in a metal bath at 95 &#176;C for 20 min, and then vortex for 15 s.</li>
	<li>Add 4 &#181;L of 1 M Tris&#183;HCl (pH 8.0) into each tube to neutralize the NaOH. Briefly centrifuge at 2,000 x g for 10 s at room temperature to remove droplets from the tube wall. Collect the supernatant into new PCR tubes, and store them at &#8722;20 &#176;C.</li>
	<li>Design appropriate primers upstream and downstream of the gRNA target sites. Use 1 &#181;L of the above supernatant as template for the PCR reactions of 50 &#181;L volume. The primers used for the PCR in this study are listed in Supplementary Table 1.</li>
	<li>Perform a DNA agarose gel electrophoresis to ensure correct and specific bands, followed by Sanger sequencing as previously described40.</li>
	<li>Analyze the Sanger sequencing data with the EditR (1.0.10) program41.
	<ol>
		<li>Upload the sequencing file (ab1 format) into the &#34;Upload .ab1 File&#34; box.</li>
		<li>Enter the 20 bp gRNA sequence in 5&#8242;-3&#8242; orientation into the &#34;Enter gRNA sequence&#34; box.</li>
		<li>Open the sequencing file (ab1 format), and confirm the base positions where the 20 bp gRNA sequence starts and ends. Enter the corresponding base position in the &#34;5&#8242; start&#34; and &#34;3&#8242; end&#34; boxes.</li>
		<li>Click on Predicted Editing to obtain the base editing efficiency. Check the quality of the sequencing data near the gRNA target sequence to avoid disorderly peaks or indels. 
		NOTE: In general, the disorderly peaks can be effectively removed by adjusting the PCR annealing temperature.</li>
	</ol>
	</li>
</ol>

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The authors declare no conflicts of interest.

This protocol describes the construction of a zebrafish disease model using the base editor zSpRY-ABE8e. Compared with the traditional HDR pathway for base substitution, this protocol can achieve more efficient base editing and reduce the occurrence of indels. At the same time, this protocol involves implementing the recently proposed i-Silence gene-knockout strategy in zebrafish. Taken together, zSpRY-ABE8e will enhance the application of zebrafish models in disease research.
Off-target effec...

Single-nucleotide variants (SNVs) that cause missense or nonsense mutations are the most common source of mutations in the human genome1. To determine whether a particular SNV is pathogenic, and to shed light on its pathogenesis, precise animal models are required2. Zebrafish are good human disease models, exhibiting a high degree of physiological and genetic homology with humans, a short developmental cycle, and strong reproductive ability, which is advantageous for research into pathogenic characteristics and mechanisms, as well as drug screening3.
The clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9 system has been widely applied in the genome editing of various species, including zebrafish4. With gRNA guidance, the CRISPR/Cas9 system can generate DNA double-stranded breaks (DSBs) at the target site, which then allows single-base substitution through recombination of the target site with donor DNA templates via the homology-directed repair (HDR) pathway. However, the efficiency of this base replacement method is quite low as the cellular DNA repair process is mainly carried out by the non-homologous end-joining (NHEJ) pathway, which is usually accompanied by insertion and deletion (indel) mutations5. Fortunately, CRISPR/Cas9-based single-base editing technology significantly alleviates this problem by using base editors, which enable more efficient single-base editing without inducing DSBs. Two major classes of base editors, adenine base editors (ABEs) and cytosine base editors (CBEs), have been developed to implement base substitution editing for A&#183;T to G&#183;C and C&#183;G to T&#183;A, respectively6,7,8,9,10,11. These four types of base substitutions cover 30% of human pathogenic variants12. Both classes of base editors, including PmCDA1, BE system, CBE4max, ABE7.10, and ABE8e, have been reported to work in zebrafish, with BE4max and ABE8e especially reported to achieve high editing activity13,14,15,16,17,18,19.
Cas9 proteins from different species, including Staphylococcus aureus, Streptococcus pyogenes, and S. canis, have been implemented in zebrafish gene editing, with the Streptococcus pyogenes Cas9 (SpCas9) being used most widely20,21,22,23. However, SpCas9 can only recognize target sites with an NGG protospacer adjacent motif (PAM), which limits its editable range and can result in no suitable sequence being found near the pathogenic site of interest24. To expand the target range, a variety of SpCas9 variants have been engineered to recognize different PAMs through directed evolution and structure-guided design. However, few variants are effective in animals, especially in zebrafish, which limits the application of zebrafish in SNV-related disease research25,26,27,28. Recently, two variants of SpCas9, SpG and SpRY, with less stringent PAM restrictions (NGN for SpG and NNN for SpRY with a higher preference for NRN than NYN) have been reported to exhibit high editing activity in human cells and plants29,30,31,32. Subsequently, SpG and SpRY, as well as a number of their mediated base editors, such as SpRY-mediated CBEs and SpRY-mediated ABEs, have also been reported to work in zebrafish, which will enhance the application of zebrafish models in the mechanistic study and drug screening of SNV-related diseases18,33,34,35. Furthermore, i-Silence was proposed as an effective and accurate gene-knockout strategy through ABE-mediated start codon conversion from ATG to GTG or ACG36. The combination of the i-Silence strategy and the SpRY-mediated base editor zSpRY-ABE8e provides a new method for disease modeling18. This protocol demonstrates how to perform gene editing using zSpRY-ABE8e in zebrafish to construct a tsr2 (M1V) model using the i-Silence strategy. The editing efficiency and phenotypes that appear in zebrafish models were assessed and analyzed.

<table><tbody><tr><td>Agarose</td><td>Sigma-Aldrich</td><td>A9539</td><td>1.5% Agarose used to make an injection plate</td></tr><tr><td>Borosilicate Glass Capillaries&amp;nbsp;</td><td>Harvard Apparatus</td><td>BS4 30-0016</td><td></td></tr><tr><td>Cell culture dishes&amp;nbsp;</td><td>Falcon</td><td>351029</td><td></td></tr><tr><td>ClonExpress Ultra One Step Cloning Kit</td><td>Vazyme</td><td>C115</td><td>Kit for Infusion clone&amp;nbsp;</td></tr><tr><td>Codon optimization service</td><td>Sangon Biotech</td><td></td><td></td></tr><tr><td>Drummond Microcaps</td><td>Drummond Microcaps</td><td>P1299-1PAK</td><td>Length:32 mm, capacity:0.5 &amp;mu;L</td></tr><tr><td>EasyEdit gRNA service</td><td>GenScript</td><td></td><td></td></tr><tr><td>Fine Forceps</td><td>Fine Scientific Instrument</td><td>11254-20</td><td>Used to break meedle</td></tr><tr><td>Flaming/Brown Micropipette Puller&amp;nbsp;</td><td>Stutter Instrument</td><td>P-97</td><td>Used to pull the glass capillaries</td></tr><tr><td>HotStart Taq PCR StarMix</td><td>Genstar</td><td>A033-101</td><td>Used for PCR reaction</td></tr><tr><td>Intelligent artificial climate box</td><td>TENLIN</td><td>PRX-1000A</td><td>Used to culture zebrafish embryos</td></tr><tr><td>Methylcellulose</td><td>Sigma-Aldrich</td><td>M0512</td><td>Used to fix zebrafish when photographing</td></tr><tr><td>Microloader pipette tips&amp;nbsp;</td><td>Eppendorf</td><td>5242956003</td><td></td></tr><tr><td>mMACHINE kit&amp;nbsp;</td><td></td><td></td><td></td></tr><tr><td>Mut Express II Fast Mutagenesis Kit V2</td><td>Vazyme</td><td>C214-01</td><td>Kit for site-directed mutagenesis</td></tr><tr><td>Pneumatic Microinjector</td><td>ZGene Biotech</td><td>ZGPCP-1500 PLUS</td><td></td></tr><tr><td>pT3TS-zSpCas9</td><td>Addgene</td><td>46757</td><td></td></tr><tr><td>RNeasy FFPE kit&amp;nbsp;</td><td>Qiagen</td><td>73504</td><td>Kit for RNA purification</td></tr><tr><td>Sanger Sequencing service</td><td>Sangon Biotech</td><td></td><td></td></tr><tr><td>Sodium hydroxide, granular</td><td>Sangon</td><td>A100173-0500</td><td>NaOH used for genome extraction</td></tr><tr><td>Stereo Microscope</td><td>Olympus&amp;nbsp;</td><td>SZX10</td><td>Used for photograph of phenotype</td></tr><tr><td>SZ Series Zoom Stereo Microscope</td><td>CNOPTEC</td><td>SZ650</td><td></td></tr><tr><td>T3 mMESSAGE</td><td>Ambion</td><td>AM1348</td><td>Kit for in vitro transcription&amp;nbsp;</td></tr><tr><td>TIANprep Mini Plasmid Kit</td><td>TIANGEN</td><td>DP103-03</td><td>Kit for plasmid extraction kit</td></tr><tr><td>TIANquick Mini Purification Kit</td><td>TIANGEN</td><td>DP203-02</td><td>Kit for purification for linearized plasmid</td></tr><tr><td>Tricaine</td><td>Sigma-Aldrich</td><td>E10521</td><td>Used to anesthetize zebrafish</td></tr><tr><td>Tris (hydroxymethyl) aminomethane</td><td>Sangon</td><td>A600194-0500</td><td>Component of Tris&amp;middot;HCl used for genome extraction</td></tr><tr><td>&lt;em&gt;XbaI&lt;/em&gt;</td><td>New England Biolabs</td><td>R0145S</td><td>Restriction endonuclease used for plasmid linearization</td></tr></tbody></table>

efficient pam-less base editing for zebrafish modeling of human genetic disease with zspry-abe8e

CRISPR/Cas9 technology has increased the value of zebrafish for modeling human genetic diseases, studying disease pathogenesis, and drug screening, but protospacer adjacent motif (PAM) limitations are a major obstacle to creating accurate animal models of human genetic disorders caused by single-nucleotide variants (SNVs). Until now, some SpCas9 variants with broad PAM compatibility have shown efficiency in zebrafish. The application of the optimized SpRY-mediated adenine base editor (ABE), zSpRY-ABE8e, and synthetically modified gRNA in zebrafish has enabled efficient adenine-guanine base conversion without PAM restriction. Described here is a protocol for efficient adenine base editing without PAM limitation in zebrafish using zSpRY-ABE8e. By injecting a mixture of zSpRY-ABE8e mRNA and synthetically modified gRNA into zebrafish embryos, a zebrafish disease model was constructed with a precise mutation that simulated a pathogenic site of the TSR2 ribosome maturation factor (tsr2). This method provides a valuable tool for the establishment of accurate disease models for studying disease mechanisms and treatments.

The mutation of TSR2 has been reported to cause Diamond Blackfan anemia (DBA)42. Here, a DBA zebrafish model was constructed with a tsr2 (M1V) mutation using the i-Silence strategy. The adenine of the start codon of the zebrafish tsr2 was successfully converted to guanine using zSpRY-ABE8e (Figure 3).
The EditR analysis of the Sanger sequencing results showed that there was an A/G overlap at the adenine base of th...

Watch this Scientific Journal Video about Efficient PAM-Less Base Editing for Zebrafish Modeling of Human Genetic Disease with zSpRY-ABE8e at JoVE.com

Efficient PAM-Less Base Editing for Zebrafish Modeling of Human Genetic Disease with zSpRY-ABE8e

This protocol describes how to perform efficient adenine base editing without PAM limitation to construct a precise zebrafish disease model using zSpRY-ABE8e.

CRISPR/Cas9 technology has increased the value of zebrafish for modeling human genetic diseases, studying disease pathogenesis, and drug screening, but ...

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1.1K Views.  Ministry of Education. This protocol describes how to perform efficient adenine base editing without PAM limitation to construct a precise zebrafish disease model using zSpRY-ABE8e. 

Video: Efficient PAM-Less Base Editing for Zebrafish Modeling of Human Genetic Disease with zSpRY-ABE8e

Microbead Implantation in the Zebrafish Embryo

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic conservation, as well as similarities in cellular, molecular, and physiological processes, with other vertebrates including humans. During early ontogeny, zebrafish embryos are optically transparent, allowing researchers to visualize the dynamics of organogenesis using a simple stereomicroscope. Microbead implantation is a method that enables tissue manipulation through the alteration of factors in local environments. This allows researchers to assay the effects of any number of signaling molecules of interest, such as secreted peptides, at specific spatial and temporal points within the developing embryo. Here, we detail a protocol for how to manipulate and implant beads during early zebrafish development.

The zebrafish is an excellent model system for genetic and developmental studies. Bead implantation is a valuable tissue manipulation technique that can be used to interrogate developmental mechanisms by introducing alterations in local cellular environments. This protocol describes how to perform microbead implantation in the zebrafish embryo.

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic ...

Developmental Biology

Sample Preparation and Analysis of RNASeq-based Gene Expression Data from Zebrafish

The analysis of global gene expression changes is a valuable tool for identifying novel pathways underlying observed phenotypes. The zebrafish is an excellent model for rapid assessment of whole transcriptome from whole animal or individual cell populations due to the ease of isolation of RNA from large numbers of animals. Here a protocol for global gene expression analysis in zebrafish embryos using RNA sequencing (RNASeq) is presented. We describe preparation of RNA from whole embryos or from cell populations obtained using cell sorting in transgenic animals. We also describe an approach for analysis of RNASeq data to identify enriched pathways and Gene Ontology (GO) terms in global gene expression data sets. Finally, we provide a protocol for validation of gene expression changes using quantitative reverse transcriptase PCR (qRT-PCR). These protocols can be used for comparative analysis of control and experimental sets of zebrafish to identify novel gene expression changes, and provide molecular insight into phenotypes of interest.

This protocol presents an approach for whole transcriptome analysis from zebrafish embryos, larvae, or sorted cells. We include isolation of RNA, pathway analysis of RNASeq data, and qRT-PCR-based validation of gene expression changes.

The analysis of global gene expression changes is a valuable tool for identifying novel pathways underlying observed phenotypes. The zebrafish is an ...

Genetics

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

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.

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Targeted genome editing in the model system Danio rerio (zebrafish) has been greatly facilitated by the emergence of CRISPR-based approaches. Herein, we describe a streamlined, robust protocol for generation and identification of CRISPR-derived nonsense alleles that incorporates the heteroduplex mobility assay and identification of mutations using next-generation sequencing.

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development ...

Microinjection for Transgenesis and Genome Editing in Threespine Sticklebacks

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and behavioral phenotypes. The remarkably diverse phenotypes that have evolved as marine populations adapt to countless freshwater environments, combined with the ability to cross marine and freshwater forms, provide a rare vertebrate system in which genetics can be used to map genomic regions controlling evolved traits. Excellent genomic resources are now available, facilitating molecular genetic dissection of evolved changes. While mapping experiments generate lists of interesting candidate genes, functional genetic manipulations are required to test the roles of these genes. Gene regulation can be studied with transgenic reporter plasmids and BACs integrated into the genome using the Tol2 transposase system. Functions of specific candidate genes and cis-regulatory elements can be assessed by inducing targeted mutations with TALEN and CRISPR/Cas9 genome editing reagents. All methods require introducing nucleic acids into fertilized one-cell stickleback embryos, a task made challenging by the thick chorion of stickleback embryos and the relatively small and thin blastomere. Here, a detailed protocol for microinjection of nucleic acids into stickleback embryos is described for transgenic and genome editing applications to study gene expression and function, as well as techniques to assess the success of transgenesis and recover stable lines.

Transgenic manipulations and genome editing are critical for functionally testing the roles of genes and cis-regulatory elements. Here a detailed microinjection protocol for the generation of genomic modifications (including Tol2-mediated fluorescent reporter transgene constructs, TALENs, and CRISPRs) is presented for the emergent model fish, the threespine stickleback.

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and ...

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin condensation, microtubule-kinetochore attachment, chromosome segregation, and cytokinesis in a small time frame. Errors in the delicate process can result in human disease, including birth defects and cancer. Traditional approaches investigating human mitotic disease states often rely on cell culture systems, which lack the natural physiology and developmental/tissue-specific context advantageous when studying human disease. This protocol overcomes many obstacles by providing a way to visualize, with high resolution, chromosome dynamics in a vertebrate system, the zebrafish. This protocol will detail an approach that can be used to obtain dynamic images of dividing cells, which include: in vitro transcription, zebrafish breeding/collecting, embryo embedding, and time-lapse imaging. Optimization and modifications of this protocol are also explored. Using H2A.F/Z-EGFP (labels chromatin) and mCherry-CAAX (labels cell membrane) mRNA-injected embryos, mitosis in AB wild-type, auroraBhi1045, and esco2hi2865 mutant zebrafish is visualized. High resolution live imaging in zebrafish allows one to observe multiple mitoses to statistically quantify mitotic defects and timing of mitotic progression. In addition, observation of qualitative aspects that define improper mitotic processes (i.e., congression defects, missegregation of chromosomes, etc.) and improper chromosomal outcomes (i.e., aneuploidy, polyploidy, micronuclei, etc.) are observed. This assay can be applied to the observation of tissue differentiation/development and is amenable to the use of mutant zebrafish and pharmacological agents. Visualization of how defects in mitosis lead to cancer and developmental disorders will greatly enhance understanding of the pathogenesis of disease.

Mitosis is critical to every living organism and defects often lead to cancer and developmental disorders. Using this imaging protocol and zebrafish as a model system, researchers can visualize mitosis in a live vertebrate organism and the multitude of defects that arise when mitotic processes are defective.

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin ...

High-throughput DNA Extraction and Genotyping of 3dpf Zebrafish Larvae by Fin Clipping

Zebrafish (Danio rerio) possess orthologues for 84% of the genes known to be associated with human diseases. In addition, these animals have a short generation time, are easy to handle, display a high reproductive rate, low cost, and are easily amenable to genetic manipulations by microinjection of DNA in embryos. Recent advances in gene editing tools are enabling precise introduction of mutations and transgenes in zebrafish. Disease modeling in zebrafish often leads to larval phenotypes and early death which can be challenging to interpret if genotypes are unknown. This early identification of genotypes is also needed in experiments requiring sample pooling, such as in gene expression or mass spectrometry studies. However, extensive genotypic screening is limited by traditional methods, which in most labs are performed only on adult zebrafish or in postmortem larvae. We addressed this problem by adapting a method for the isolation of PCR-ready genomic DNA from live zebrafish larvae that can be achieved as early as 72 h post-fertilization (hpf). This time and cost-effective technique, improved from a previously published genotyping protocol, allows the identification of genotypes from microscopic fin biopsies. The fins quickly regenerate as the larvae develop. Researchers are then able to select and raise the desired genotypes to adulthood by utilizing this high-throughput PCR-based genotyping procedure.

Zebrafish have been used as reliable genetic model organisms in biomedical research, especially with the advent of gene-editing technologies. When larval phenotypes are expected, DNA extraction and genotype identification can be challenging. Here, we describe an efficient genotyping procedure for zebrafish larvae, by tail clipping, as early as 72-h post-fertilization.

Zebrafish (Danio rerio) possess orthologues for 84% of the genes known to be associated with human diseases. In addition, these animals have a short ...

Manipulation of Gene Function in Mexican Cavefish

Cave animals provide a compelling system for investigating the evolutionary mechanisms and genetic bases underlying changes in numerous complex traits, including eye degeneration, albinism, sleep loss, hyperphagia, and sensory processing. Species of cavefish from around the world display a convergent evolution of morphological and behavioral traits due to shared environmental pressures between different cave systems. Diverse cave species have been studied in the laboratory setting. The Mexican tetra, Astyanax mexicanus, with sighted and blind forms, has provided unique insights into biological and molecular processes underlying the evolution of complex traits and is well-poised as an emerging model system. While candidate genes regulating the evolution of diverse biological processes have been identified in A. mexicanus, the ability to validate a role for individual genes has been limited. The application of transgenesis and gene-editing technology has the potential to overcome this significant impediment and to investigate the mechanisms underlying the evolution of complex traits. Here, we describe a different methodology for manipulating gene expression in A. mexicanus. Approaches include the use of morpholinos, Tol2 transgenesis, and gene-editing systems, commonly used in zebrafish and other fish models, to manipulate gene function in A. mexicanus. These protocols include detailed descriptions of timed breeding procedures, the collection of fertilized eggs, injections, and the selection of genetically modified animals. These methodological approaches will allow for the investigation of the genetic and neural mechanisms underlying the evolution of diverse traits in A. mexicanus.&#160;

We describe approaches for the manipulation of genes in the evolutionary model system Astyanax mexicanus. Three different techniques are described: Tol2-mediated transgenesis, targeted manipulation of the genome using CRISPR/Cas9, and knockdown of expression using morpholinos. These tools should facilitate the direct investigation of genes underlying the variation between surface- and cave-dwelling forms.

Cave animals provide a compelling system for investigating the evolutionary mechanisms and genetic bases underlying changes in numerous complex traits, ...

Screening Sperm for the Rapid Isolation of Germline Edits in Zebrafish

The advent of targeted CRISPR-Cas nuclease technologies has revolutionized the ability to perform precise genome editing in both established and emerging model systems. CRISPR-Cas genome editing systems use a synthetic guide RNA (sgRNA) to target a CRISPR-associated (Cas) endonuclease to specific genomic DNA loci, where the Cas endonuclease generates a double-strand break. The repair of double-strand breaks by intrinsic error-prone mechanisms leads to insertions and/or deletions, disrupting the locus. Alternatively, the inclusion of double-stranded DNA donors or single-stranded DNA oligonucleotides in this process can elicit the inclusion of precise genome edits ranging from single nucleotide polymorphisms to small immunological tags or even large fluorescent protein constructs. However, a major bottleneck in this procedure can be finding and isolating the desired edit in the germline. This protocol outlines a robust method for screening and isolating germline mutations at specific loci in Danio rerio (zebrafish); however, these principles may be adaptable in any model where in vivo sperm collection is possible.

CRISPR-Cas technologies have revolutionized the field of genome editing. However, finding and isolating the desired germline edit remains a major bottleneck. Therefore, this protocol describes a robust method for quickly screening F0 CRISPR-injected zebrafish sperm for germline edits using standard PCR, restriction digest, and gel electrophoresis techniques.

The advent of targeted CRISPR-Cas nuclease technologies has revolutionized the ability to perform precise genome editing in both established and emerging ...

Fluorescent Protein Tagging in Zebrafish Larvae Using CRISPR/Cas9 Knock-In

Fluorescence Labeling to Visualize Low-Expressed Proteins in Zebrafish

CRISPR/Cas9-mediated knock-in (KI) technology allows for easier fluorescent-protein tagging in zebrafish (Danio rerio), a preferred model organism for in vivo imaging due to its transparency during the early developmental stage. Here, we provide a detailed protocol for performing high-efficiency fluorescence gene KI, rapid screening for KI founders, and low-abundance protein tracing in zebrafish larvae, which will lay a critical foundation for subsequent physio-pathological studies in zebrafish. The current protocol includes complete steps for the sgRNA design for the gene of interest, sgRNA in vitro transcription, Cas9 mRNA in vitro transcription, in vivo sgRNA screen for the one with the highest efficiency, donor plasmid design and construction, microinjection in zebrafish larvae, KI founder screen and zebrafish live imaging. Critical steps, troubleshooting tips, quality control methods, and advantages and applications of this protocol are included and discussed. This protocol assures quick and accurate results at a low cost and has been validated by multiple trials.

Here, we present a protocol to label proteins with a fluorescence protein tag in zebrafish larvae, a newly developed and modified in vivo system especially useful for visualizing low-abundance proteins in zebrafish.

CRISPR/Cas9-mediated knock-in (KI) technology allows for easier fluorescent-protein tagging in zebrafish (Danio rerio), a preferred model organism for in ...

In Vivo Modeling of the Morbid Human Genome using Danio rerio

Here, we present methods for the development of assays to query potentially clinically significant nonsynonymous changes using in vivo complementation in zebrafish. Zebrafish (Danio rerio) are a useful animal system due to their experimental tractability; embryos are transparent to enable facile viewing, undergo rapid development ex vivo, and can be genetically manipulated.1 These aspects have allowed for significant advances in the analysis of embryogenesis, molecular processes, and morphogenetic signaling. Taken together, the advantages of this vertebrate model make zebrafish highly amenable to modeling the developmental defects in pediatric disease, and in some cases, adult-onset disorders. Because the zebrafish genome is highly conserved with that of humans (~70% orthologous), it is possible to recapitulate human disease states in zebrafish. This is accomplished either through the injection of mutant human mRNA to induce dominant negative or gain of function alleles, or utilization of morpholino (MO) antisense oligonucleotides to suppress genes to mimic loss of function variants. Through complementation of MO-induced phenotypes with capped human mRNA, our approach enables the interpretation of the deleterious effect of mutations on human protein sequence based on the ability of mutant mRNA to rescue a measurable, physiologically relevant phenotype. Modeling of the human disease alleles occurs through microinjection of zebrafish embryos with MO and/or human mRNA at the 1-4 cell stage, and phenotyping up to seven days post fertilization (dpf). This general strategy can be extended to a wide range of disease phenotypes, as demonstrated in the following protocol. We present our established models for morphogenetic signaling, craniofacial, cardiac, vascular integrity, renal function, and skeletal muscle disorder phenotypes, as well as others.

In Vivo Modeling of the Morbid Human Genome using Danio rerio

Here, we present a systematic approach for developing physiologically relevant, sensitive and specific in vivo assays for interpreting variation in human pathology. Transient genetic manipulation via microinjection of WT and mutant human mRNA and morpholino (MO) antisense oligonucleotides harness the tractability of the developing zebrafish embryo to rapidly assay pathogenic mutations, especially, but not exclusively, in the context of human developmental disorders.

Here,  we present methods for the development of assays to query potentially clinically  significant nonsynonymous changes using in  vivo complementation ...

Efficient PAM-Less Base Editing for Zebrafish Modeling of Human Genetic Disease with zSpRY-ABE8e

Summary

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Chapters in this video

Microbead Implantation in the Zebrafish Embryo

Sample Preparation and Analysis of RNASeq-based Gene Expression Data from Zebrafish

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Microinjection for Transgenesis and Genome Editing in Threespine Sticklebacks

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

High-throughput DNA Extraction and Genotyping of 3dpf Zebrafish Larvae by Fin Clipping

Manipulation of Gene Function in Mexican Cavefish

Screening Sperm for the Rapid Isolation of Germline Edits in Zebrafish

Fluorescence Labeling to Visualize Low-Expressed Proteins in Zebrafish

In Vivo Modeling of the Morbid Human Genome using Danio rerio