The authors would like to thank the Rare Disease Models and Mechanism (RDMM) Network (funded by the Canadian Institutes of Health Research (CIHR) and Genome Canada). CK is supported by a UROP scholarship award (University of Ottawa). DLJ is supported by a Vanier Canada Graduate Scholarship. IAP is supported by a Canadian Institutes of Health Research (CIHR) postdoctoral fellowship award. The authors thank the Genetics Society of America for granting permission to publish this protocol and to use an adaptation of the Supplemental Figure 5 from Pena, et al.

Procedures involving animal subjects have been approved and are in accordance with animal care guidelines provided by the Canadian Council on Animal Care and the University of Ottawa animal care committees.
1. Preparation
<ol>
	<li>Prepare the dissection surface by taping a 9 cm Petri dish lid with autoclave tape across its interior surface. Position it under a stereo-microscope.</li>
	<li>Place 3 - 5 days post-fertilization (dpf) zebrafish larvae in a Petri dish and anesthetize in &#732;1.5 mM Tricaine in 1x E3 embryo media (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4, pH 7.2).</li>
	<li>Cut off the end of a P1000 pipette tip with a pair of scissors or a razor blade to a diameter of 2 mm to accommodate a 3 dpf larva with minimal stress.</li>
</ol>
2. Fin Clipping of Zebrafish Larvae
<ol>
	<li>Pick up an anesthetized larva using the modified P1000 micropipette and place the zebrafish larva onto the positioned autoclave tape on the Petri dish lid.</li>
	<li>Remove excess Tricaine solution surrounding the larva using a micropipette. The area should be as dry as possible to successfully section and pick up the fin but still wet to ensure survival.</li>
	<li>Using a micro scalpel, section the caudal fin under a stereo-microscope as indicated in Figure 1, as described in Wilkinson&#160;et al16. During the incision, apply a steady downward pressure within the pigment gap site of the caudal fin, distal to the limit of the blood circulation (Figure 1A). Apply gentle pressure to avoid damaging the notochord.</li>
	<li>Visualize the sectioned fin under the microscope and position the piece on top of the tip of the microscalpel blade (Figure 2A). Place the microscalpel containing the fin onto the surface of a small piece of filter paper. 
	NOTE: Due to the presence of melanocytes in the transected tissue, this step allows visualization of the fin as a small black spot (Figure 2B).</li>
	<li>Using scissors and tweezers (Figure 2C), transfer the filter paper containing the fin to a 96-well PCR plate, containing 25 &#956;L of 50 mM NaOH solution per well (Figure 2D).</li>
	<li>Prepare a flat-bottom 96-well plate, numbering the well according to the label of the PCR plate. Using the modified P1000 pipette filled with 200 &#181;L of fresh 1x E3 embryo media, carefully collect the zebrafish larva using gentle pressure. Dispense the larva in the 96-well tissue culture plate, in the same well number as the PCR plate (Figure 2E).</li>
	<li>Make sure that the filter paper is well submerged in the solution (Figure 2F). Clean the blade of the microscapel and tweezers between each cut to prevent cross-contamination by dipping it into 70% EtOH solution. Wipe the blade with clean paper tissue/wipe.</li>
	<li>Repeat steps 2.1 - 2.7 until all embryos have been clipped.</li>
	<li>Store the larvae in the 96-well plate at 28 &#176;C until the genotypes have been identified. 
	NOTE: The embryo media does not need to be changed, since the protocol can be easily completed in less than 2 days. If more time is needed, a media change is recommended.</li>
</ol>
3. Genomic DNA Extraction
<ol>
	<li>Seal the 96-well PCR plate and centrifuge the samples at 1,000 x g for 1 min to make sure all the filter papers are submerged within the NaOH solution.</li>
	<li>For tissue lysis, heat the samples in a thermocycler at 95 &#176;C for 5 min followed by cooling to 4 &#176;C for 10 min.</li>
	<li>Add 6 &#956;L of 500 mM Tris-HCl, pH 8.0 to each sample. Vortex.</li>
	<li>Briefly centrifuge the plate at 1,500 x g, for 5 min at room temperature.</li>
	<li>Use 1.5 &#956;L of the DNA supernatant per PCR reaction.</li>
	<li>Prepare a PCR to genotype the larvae, as shown in Table 1-2. 
	NOTE: This protocol was tested for two applications: multiplex PCR reactions revealing the genotype using agarose gels9, and heteroduplex melting assay (HMA) revealing the genotypes using polyacrylamide gel electrophoresis (PAGE)17.</li>
</ol>
4. Alternative Genomic Extraction Using a Chelating Resin
<ol>
	<li>Complete step 2.5 by adding filter paper with the clipped fin to a 96-well PCR plate, containing 30 &#956;L of 5% chelating resin (styrene-divinylbenzene copolymer containing paired iminodiacetate ions). 
	NOTE: This type of resin is commonly used for tissue lysis and preparation of PCR-ready DNA in a fast and efficient manner18.</li>
	<li>Follow steps 2.6 - 2.8 as indicated.</li>
	<li>Seal the 96-well PCR plate and briefly centrifuge the samples to make sure all filter paper is submerged within the chelating resin.</li>
	<li>For tissue lysis, heat the samples in a thermocycler at 95 &#176;C for 15 min followed by cooling to 4 &#176;C for 10 min. 
	NOTE: This step allows lysis and subsequent binding of the polar resin beads to cellular components while DNA and RNA remain in solution.</li>
	<li>Briefly centrifuge the samples to pellet the resin beads and obtain the DNA in the suspension.</li>
	<li>Follow steps 3.5 - 3.6 to complete genotyping procedure.</li>
</ol>
5. Application 1: Multiplex PCR Followed by Agarose Gel Analysis
<ol>
	<li>Following Table 1, set up a multiplex PCR reaction to enable the discrimination between homozygous mutants, heterozygotes, and WT larvae. Use a 96-well thermal cycler to perform the PCR reactions using the cycling conditions described in Table 3. 
	NOTE: The PCR reactions can also be performed in any other conventional PCR thermal cycler. This example describes the PCR strategy used by Pena, et al.9 to identify aldh7a1 WT and 5-bp insertion mutant alleles in the same multiplex reaction.</li>
	<li>Prepare a 1% agarose gel in sodium borate buffer, stained with 1x GelRed. 
	NOTE: 20x sodium borate buffer stock is made of 40 mL of 10 M NaOH, 1800 mL of ddH2O, pH adjusted to 8.5 with 76 g of boric acid and then completed to 2 L final volume using ddH2O.</li>
	<li>Run the gel in 1x sodium borate buffer for 15 min at 250 V. To facilitate the process, if possible, use a gel system that is compatible with multichannel pipets to enable higher throughput capabilities by reducing the time to load samples.</li>
	<li>Image the gel under ultraviolet (UV) light to allow discrimination of the different genotypes.</li>
</ol>
6. Application 2: Heteroduplex Melting Assay
<ol>
	<li>Prepare PAGE 12% gels using the 1.5 mm spacer plate and the 15-samples combs. Prepare two PAGE gels using 12.21 mL of ddH2O, 2.52 mL of 10x Tris/Borate/EDTA (TBE), 10 mL of 30% acrylamide, 250 &#956;L of 10% ammonium persulfate (APS) and 20 &#956;L of tetramethylethylenediamine (TEMED). Use 1x TBE buffer (0.089 M Tris-HCl, 0.089 M boric acid and 0.002 M ethylenediaminetetraacetic acid (EDTA)) as running buffer.</li>
	<li>Heat the PCR products at 94 &#176;C for 5 min.</li>
	<li>Cool the tubes on ice or in the thermocycler for 10 min at 4 &#176;C.</li>
	<li>Load 10 &#956;L per PCR sample and 8 &#956;L of molecular weight marker.</li>
	<li>Run the PAGE at 150 V in 1x TBE using a vertical electrophoresis system for 1 h or until the molecular weight marker almost reaches the foot line of the glass plate.</li>
	<li>Image the gel under UV light to allow discrimination of the different genotypes.</li>
</ol>

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The authors declare that they have no competing financial interests.

Gene editing tools have been employed to precisely introduce mutations and transgenes in zebrafish over the past years5,6,7,8. When performing disease modeling in zebrafish, early larval or juvenile phenotypes are often observed9,12,13,14. The protocol presented here ...

The zebrafish (Danio rerio) is a vertebrate organism widely used as a model for disease investigation as well as for preclinical testing of therapeutic hypotheses1,2,3. 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 embryos4. Through the increasing development of novel transgenic and gene editing technologies such as Zinc-Finger nucleases (ZFN)5, TAL-like Effector Nucleases (TALENs)6,7, and the Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/CRISPR-associated 9 (Cas9) system8, the zebrafish is poised to substantially improve the understanding of several pathologic conditions. These technologies have been used to create targeted knockouts in both somatic and germline cells in zebrafish, efficiently engendering genetically modified animals8. Recent examples in the literature include the successful development of genetic models of epilepsy and metabolic disorders in zebrafish3,9,10,11,12.
With the progress made in zebrafish genome editing, rapid and reliable genotyping has become an undeniable rate-limiting step. Identification of specific DNA mutations adjacent to the predicted genome-editing target site is an essential stage of the protocol. Traditionally, genotyping techniques by fin clipping are usually only performed in juvenile or adult zebrafish. However, the time spent raising zebrafish to adulthood (&#62;2 months) considerably delays advancements in research. In many cases, adulthood cannot be achieved due to the knockout of essential genes (e.g., in disease modeling) or gain-of-function mutations leading to detrimental phenotypic effects. In addition, several assays require pooling of larvae, such as in RNA or metabolite extraction, and thus involve prior identification of the genotypes, which can be challenging when only post hoc genotyping techniques are available. These aforementioned examples highlight the importance of larval genotyping techniques which are at the same time precise and enable full recovery and normal development of the larvae. Early stage zebrafish genotyping does not currently appear to be widely used; this is reflected by recent publications of disease models in which larval genotypes are identified via post-mortem genotyping rather than genotyping prior to the execution of the experiment12,13,14. In this paper, a larval fin clip strategy is presented aiming to improve previously established genotyping procedures.
In the past, strategies employing proteinase K digestion to genotype live zebrafish larvae have led to variable polymerase chain reaction (PCR) efficiency15. Although a more recently published microscopic tail biopsy technique provided more promising results16, we and other groups were not able to reproduce the high efficiency and DNA recovery reported by the authors. A major setback of the protocol described by Wilkinson&#160;et al.16 could be the transfer of the tail tissue to the PCR tube. Due to its microscopic size, it is hard to ensure that the fin was properly collected and dispensed by pipetting. To address this barrier, we have developed an improved protocol for genotyping large numbers of live zebrafish larvae with nearly 100% efficiency9. Using a microscalpel, the tip of the fin tail is removed and placed onto a piece of filter paper. The piece of filter paper containing the tissue can be readily visualized and correctly placed into a PCR tube. Genomic DNA extraction is then performed, followed by PCR amplification of the region of interest to genotype the specimen. The advantages of this method include a high PCR efficiency, a low false positive rate and a low mortality rate. Additionally, genotyping of large numbers of embryos is possible using this protocol. The protocol described herein allows fin-clipping of hundreds of larvae within 2 - 3 h using this approach and correct identification of genotyped fish and tail regeneration within two days.

<table><tbody><tr><td>Petri dish</td><td>Corning</td><td>430167</td><td></td></tr><tr><td>Microscalpel</td><td>World Precision Instruments</td><td>500249</td><td></td></tr><tr><td>Stereomicroscope</td><td>Nikon</td><td>SMZ1500</td><td></td></tr><tr><td>Tricaine/Ethyl 3-aminobenzoate methanesulfonate</td><td>Sigma-Aldrich</td><td>E10521</td><td></td></tr><tr><td>96-well Tissue-culture plate</td><td>Costar</td><td>3595</td><td></td></tr><tr><td>96-well PCR plate</td><td>Fisher</td><td>14230232</td><td></td></tr><tr><td>NaOH</td><td>Fisher</td><td>S25548A&amp;nbsp;</td><td></td></tr><tr><td>Tris base&amp;nbsp;</td><td>Sigma-Aldrich</td><td>T1503</td><td></td></tr><tr><td>NaCl</td><td>Fisher</td><td>BP358-10</td><td></td></tr><tr><td>KCl</td><td>Sigma-Aldrich</td><td>P3911</td><td></td></tr><tr><td>MgSO4</td><td>Sigma-Aldrich</td><td>230391</td><td></td></tr><tr><td>5% Chelex 100 sodium form</td><td>Sigma-Aldrich</td><td>C7901</td><td></td></tr><tr><td>GelRed 1x</td><td>Biotium</td><td>41003</td><td></td></tr><tr><td>Boric Acid</td><td>Sigma-Aldrich</td><td>B6768</td><td></td></tr><tr><td>Sub-Cell Model 192 system</td><td>Bio-Rad&amp;nbsp;</td><td>1704508</td><td></td></tr><tr><td>30 % Acrylamide/Bis Solution, 37.5: 1</td><td>Bio-Rad&amp;nbsp;</td><td>1610158</td><td></td></tr><tr><td>GoTaq Green Master Mix, 2X</td><td>Promega</td><td>M7123</td><td></td></tr><tr><td>paper wipes (Kimwipes&amp;reg; disposable wipers)</td><td>Sigma-Aldrich</td><td>Z188956</td><td></td></tr><tr><td>1kb-plus molecylar weight marker&amp;nbsp;</td><td>Thermo Scientific</td><td>SM1343</td><td></td></tr><tr><td>Nuclease free water&amp;nbsp;</td><td>Sigma-Aldrich</td><td>W4502</td><td></td></tr><tr><td>Sub-Cell Model 192 system (high-throughput agarose gel system)</td><td>Bio-Rad&amp;nbsp;</td><td>1704508</td><td></td></tr><tr><td>vertical gel electrophoresis system Mini-PROTEAN Tetra Cell&amp;nbsp;</td><td>Bio-Rad&amp;nbsp;</td><td>1658004</td><td></td></tr></tbody></table>

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.

The efficiency of the protocoll was demonstrated by genotyping the offspring of an heterozygous cross (aldh7a1+/ot100, here shown as aldh7a1+/-) (Figure 3A-C)9. The aldh7a1ot100 mutant allele has a 5-base pair (bp) insertion in the first coding exon of the zebrafish aldh7a1 gene that leads to frameshift and loss-of-function due to an ...

Watch this Scientific Journal Video about High-throughput DNA Extraction and Genotyping of 3dpf Zebrafish Larvae by Fin Clipping at JoVE.com

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

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 ...

high-throughput-dna-extraction-genotyping-3dpf-zebrafish-larvae-fin

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Genetics

13.9K Views.  Children's Hospital of Eastern Ontario Research Institute. 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.

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

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 ...

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA mutation rates. Changes in replication timing occur during development and in cancer, but the role replication timing plays in development and disease is not known. Zebrafish were recently established as an in vivo model system to study replication timing. Here is detailed the protocols for using the zebrafish to determine DNA replication timing. After sorting cells from embryos and adult zebrafish, high-resolution genome-wide DNA replication timing patterns can be constructed by determining changes in DNA copy number through analysis of next generation sequencing data. The zebrafish model system allows for evaluation of the replication timing changes that occur in vivo throughout development, and can also be used to assess changes in individual cell types, disease models, or mutant lines. These methods will enable studies investigating the mechanisms and determinants of replication timing establishment and maintenance during development, the role replication timing plays in mutations and tumorigenesis, and the effects of perturbing replication timing on development and disease.

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

Zebrafish were recently used as an in vivo model system to study DNA replication timing during development. Here is detailed the protocols for using zebrafish embryos to profile replication timing. This protocol can be easily adapted to study replication timing in mutants, individual cell types, disease models, and other species.

DNA replication timing is an important cellular characteristic, exhibiting significant relationships with chromatin structure, transcription, and DNA ...

Isolation and Characterization of Single Cells from Zebrafish Embryos

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been described at the morphological level, little is known about the molecular basis of cellular changes that occur as the organism develops. With recent advancements in microfluidics and multiplexing technologies, it is now possible to characterize gene expression in single cells. This allows for investigation of heterogeneity between individual cells of specific cell populations to identify and classify cell subtypes, characterize intermediate states that occur during cell differentiation, and explore differential cellular responses to stimuli. This study describes a protocol to isolate viable, single cells from zebrafish embryos for high throughput multiplexing assays. This method may be rapidly applied to any zebrafish embryonic cell type with fluorescent markers. An extension of this method may also be used in combination with high throughput sequencing technologies to fully characterize the transcriptome of single cells. As proof of principle, the relative abundance of cardiac differentiation markers was assessed in isolated, single cells derived from nkx2.5 positive cardiac progenitors. By evaluation of gene expression at the single cell level and at a single time point, the data support a model in which cardiac progenitors coexist with differentiating progeny. The method and work flow described here is broadly applicable to the zebrafish research community, requiring only a labeled transgenic fish line and access to microfluidics technologies.

This protocol describes a method for isolating single cells from zebrafish embryos, enriching for cells of interest, capturing zebrafish cells in microfluidic based single cell multiplex systems, and assessing gene expression from single cells.

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been ...

Developmental Biology

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 ...

Genotyping and Quantification of In Situ Hybridization Staining in Zebrafish

In situ hybridization (ISH) is an important technique that enables researchers to study mRNA distribution in situ and has been a critical technique in developmental biology for decades. Traditionally, most gene expression studies relied on visual evaluation of the ISH signal, a method that is prone to bias, particularly in cases where sample identities are known a priori. We have previously reported on a method to circumvent this bias and provide a more accurate quantification of ISH signals. Here, we present a simple guide to apply this method to quantify the expression levels of genes of interest in ISH-stained embryos and correlate that with their corresponding genotypes. The method is particularly useful to quantify spatially restricted gene expression signals in samples of mixed genotypes and it provides an unbiased and accurate alternative to the traditional visual scoring methods.

Gene editing technologies have enabled researchers to generate zebrafish mutants to investigate gene function with relative ease. Here, we provide a guide to perform parallel embryo genotyping and quantification of in situ hybridization signals in zebrafish. This unbiased approach provides greater accuracy in phenotypical analyses based on in situ hybridization.

In situ hybridization (ISH) is an important technique that enables researchers to study mRNA distribution in situ and has been a critical technique in ...

Examining Muscle Regeneration in Zebrafish Models of Muscle Disease

Skeletal muscle has a remarkable ability to regenerate following injury, which is driven by obligate tissue resident muscle stem cells. Following injury, the muscle stem cell is activated and undergoes cell proliferation to generate a pool of myoblasts, which subsequently differentiate to form new muscle fibers. In many muscle wasting conditions, including muscular dystrophy and ageing, this process is impaired resulting in the inability of muscle to regenerate. The process of muscle regeneration in zebrafish is highly conserved with mammalian systems providing an excellent system to study muscle stem cell function and regeneration, in muscle wasting conditions such as muscular dystrophy. Here, we present a method to examine muscle regeneration in zebrafish models of muscle disease. The first step involves the use of a genotyping platform that allows the determination of the genotype of the larvae prior to eliciting an injury. Having determined the genotype, the muscle is injured using a needle stab, following which polarizing light microscopy is used to determine the extent of muscle regeneration. We therefore provide a high throughput pipeline which allows the examination of muscle regeneration in zebrafish models of muscle disease.

Skeletal muscle regeneration is driven by tissue resident muscle stem cells, which are impaired in many muscle diseases such as muscular dystrophy, and this results in the inability of muscle to regenerate. Here, we describe a protocol that allows the examination of muscle regeneration in zebrafish models of muscle disease.

Skeletal muscle has a remarkable ability to regenerate following injury, which is driven by obligate tissue resident muscle stem cells. Following injury, ...

Processing Gut Cells from Zebrafish Larvae for Single-Cell RNA Sequencing

Gut Isolation from Zebrafish Larvae for Single-cell RNA Sequencing

The gastrointestinal (GI) tract performs a range of functions essential for life. Congenital defects affecting its development can lead to enteric neuromuscular disorders, highlighting the importance to understand the molecular mechanisms underlying GI development and dysfunction. In this study, we present a method for gut isolation from zebrafish larvae at 5 days post fertilization to obtain live,&#160;viable cells which can be used for single-cell RNA sequencing (scRNA-seq) analysis. This protocol is based on the manual dissection of the zebrafish intestine, followed by enzymatic dissociation with papain. Subsequently, cells are submitted to fluorescence-activated cell sorting, and viable cells are collected for scRNA-seq. With this method, we were able to successfully identify different intestinal cell types, including epithelial, stromal, blood, muscle, and immune cells, as well as enteric neurons and glia. Therefore, we consider it to be a valuable resource for studying the composition of the GI tract in health and disease, using the zebrafish.

Author Spotlight: Isolating and Analyzing Intestinal Cells of Zebrafish Larvae for Investigating Transcriptomic Aspects of Gastrointestinal Development 

Here, we describe a method for gut isolation from zebrafish larvae at 5 days post fertilization, for single-cell RNA sequencing analysis.

The gastrointestinal (GI) tract performs a range of functions essential for life. Congenital defects affecting its development can lead to enteric ...

Gene Trapping Using Gal4 in Zebrafish

Large clutch size and external development of optically transparent embryos make zebrafish an exceptional vertebrate model system for in vivo insertional mutagenesis using fluorescent reporters to tag expression of mutated genes. Several laboratories have constructed and tested enhancer- and gene-trap vectors in zebrafish, using fluorescent proteins, Gal4- and lexA- based transcriptional activators as reporters 1-7. These vectors had two potential drawbacks: suboptimal stringency (e.g. lack of ability to differentiate between enhancer- and gene-trap events) and low mutagenicity (e.g. integrations into genes rarely produced null alleles). Gene Breaking Transposon (GBTs) were developed to address these drawbacks 8-10. We have modified one of the first GBT vectors, GBT-R15, for use with Gal4-VP16 as the primary gene trap reporter and added UAS:eGFP as the secondary reporter for direct detection of gene trap events. Application of Gal4-VP16 as the primary gene trap reporter provides two main advantages. First, it increases sensitivity for genes expressed at low expression levels. Second, it enables researchers to use gene trap lines as Gal4 drivers to direct expression of other transgenes in very specific tissues. This is especially pertinent for genes with non-essential or redundant functions, where gene trap integration may not result in overt phenotypes. The disadvantage of using Gal4-VP16 as the primary gene trap reporter is that genes coding for proteins with N-terminal signal sequences are not amenable to trapping, as the resulting Gal4-VP16 fusion proteins are unlikely to be able to enter the nucleus and activate transcription. Importantly, the use of Gal4-VP16 does not pre-select for nuclear proteins: we recovered gene trap mutations in genes encoding proteins which function in the nucleus, the cytoplasm and the plasma membrane.

This protocol describes the method of gene trap insertional mutagenesis using Gal4-VP16 as the primary reporter and GFP/RFP as secondary reporters in zebrafish. Approximately one in ten high-expressing F0 fish yield gene trap progeny co-expressing GFP and RFP. The screening procedure can be readily scaled to adapt to the size of the laboratory performing the insertional mutagenesis screen.

Large clutch size and external development of optically transparent embryos make zebrafish an exceptional vertebrate model system for in vivo insertional ...

Biology

Rapid and Efficient Zebrafish Genotyping Using PCR with High-resolution Melt Analysis

Zebrafish is a powerful vertebrate model system for studying development, modeling disease, and performing drug screening. Recently a variety of genetic tools have been introduced, including multiple strategies for inducing mutations and generating transgenic lines. However, large-scale screening is limited by traditional genotyping methods, which are time-consuming and labor-intensive. Here we describe a technique to analyze zebrafish genotypes by PCR combined with high-resolution melting analysis (HRMA).&#160;This approach is rapid, sensitive, and inexpensive, with lower risk of contamination artifacts.&#160;Genotyping by PCR with HRMA can be used for embryos or adult fish, including in high-throughput screening protocols.

PCR combined with high-resolution melt analysis (HRMA) is demonstrated as a rapid and efficient method to genotype zebrafish.

Zebrafish is a powerful vertebrate model system for studying development, modeling disease, and performing drug screening. Recently a variety of genetic ...

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

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

Profiling DNA Replication Timing Using Zebrafish as an In Vivo Model System

Isolation and Characterization of Single Cells from Zebrafish Embryos

Screening Sperm for the Rapid Isolation of Germline Edits in Zebrafish

Fluorescence Labeling to Visualize Low-Expressed Proteins in Zebrafish

Genotyping and Quantification of In Situ Hybridization Staining in Zebrafish

Examining Muscle Regeneration in Zebrafish Models of Muscle Disease

Gut Isolation from Zebrafish Larvae for Single-cell RNA Sequencing

Gene Trapping Using Gal4 in Zebrafish

Rapid and Efficient Zebrafish Genotyping Using PCR with High-resolution Melt Analysis