Name: crispr-cas9-mediated precise knock-in edits in zebrafish hearts
Uploaded: 2022-09-13T13:50:00
Description: Watch this Scientific Journal Video about CRISPR-Cas9-Mediated Precise Knock-In Edits in Zebrafish Hearts at JoVE.com

This research was supported by a Canadian Institutes of Health Research Project grant (T.W.C.) and Natural Sciences and Engineering Research Council of Canada Discovery grants (T.W.C.).

Studies using zebrafish were conducted in agreement with the policies and procedures of the Simon Fraser University Animal Care Committee and the Canadian Council of Animal Care and were completed under protocol # 1264K-18.
1. Design of CRISPR components for precise edits
<ol>
	<li>To design the two-sgRNA guides that will be used to excise the sequence containing the KI target site, first identify the zebrafish ortholog for the gene of interest. 
	NOTE: <strong class="x...

<ol>
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B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Homology+directed+knockin+of+point+mutations+in+the+zebrafish+tardbp+and+fus+genes+in+ALS+using+the+CRISPR/Cas9+system.">Homology directed knockin of point mutations in the zebrafish tardbp and fus genes in ALS using the CRISPR/Cas9 system.</a> PLoS One. 11 (3), 0150188 (2016).</li><li>Bai, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR/Cas9-mediated+precise+genome+modification+by+a+long+ssDNA+template+in+zebrafish.">CRISPR/Cas9-mediated precise genome modification by a long ssDNA template in zebrafish.</a> BMC Genomics. 21 (1), 67 (2020).</li><li>de Vrieze, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+generation+of+knock-in+zebrafish+models+for+inherited+disorders+using+CRISPR-Cas9+ribonucleoprotein+complexes.">Efficient generation of knock-in zebrafish models for inherited disorders using CRISPR-Cas9 ribonucleoprotein complexes.</a> International Journal of Molecular Sciences. 22 (17), 9429 (2021).</li><li>Eschstruth, A., Schneider-Maunoury, S., Giudicelli, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Creation+of+zebrafish+knock-in+reporter+lines+in+the+nefma+gene+by+Cas9-mediated+homologous+recombination.">Creation of zebrafish knock-in reporter lines in the nefma gene by Cas9-mediated homologous recombination.</a> Genesis. 58 (1), 23340 (2020).</li><li>Irion, U., Krauss, J., N&#252;sslein-Volhard, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Precise+and+efficient+genome+editing+in+zebrafish+using+the+CRISPR/Cas9+system.">Precise and efficient genome editing in zebrafish using the CRISPR/Cas9 system.</a> Development. 141 (24), 4827-4830 (2014).</li><li>Kimura, Y., Hisano, Y., Kawahara, A., Higashijima, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+generation+of+knock-in+transgenic+zebrafish+carrying+reporter/driver+genes+by+CRISPR/Cas9-mediated+genome+engineering.">Efficient generation of knock-in transgenic zebrafish carrying reporter/driver genes by CRISPR/Cas9-mediated genome engineering.</a> Scientific Reports. 4 (1), 6545 (2014).</li><li>Levic, D. S., Yamaguchi, N., Wang, S., Knaut, H., Bagnat, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Knock-in+tagging+in+zebrafish+facilitated+by+insertion+into+non-coding+regions.">Knock-in tagging in zebrafish facilitated by insertion into non-coding regions.</a> Development. 148 (19), (2021).</li><li>Prykhozhij, S. V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optimized+knock-in+of+point+mutations+in+zebrafish+using+CRISPR/Cas9.">Optimized knock-in of point mutations in zebrafish using CRISPR/Cas9.</a> Nucleic Acids Research. 46 (17), 102 (2018).</li><li>Wierson, W. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+targeted+integration+directed+by+short+homology+in+zebrafish+and+mammalian+cells.">Efficient targeted integration directed by short homology in zebrafish and mammalian cells.</a> eLife. 9, 53968 (2020).</li><li>Boel, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR/Cas9-mediated+homology-directed+repair+by+ssODNs+in+zebrafish+induces+complex+mutational+patterns+resulting+from+genomic+integration+of+repair-template+fragments.">CRISPR/Cas9-mediated homology-directed repair by ssODNs in zebrafish induces complex mutational patterns resulting from genomic integration of repair-template fragments.</a> Disease Models &amp; Mechanisms. 11 (10), (2018).</li><li>Tessadori, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effective+CRISPR/Cas9-based+nucleotide+editing+in+zebrafish+to+model+human+genetic+cardiovascular+disorders.">Effective CRISPR/Cas9-based nucleotide editing in zebrafish to model human genetic cardiovascular disorders.</a> Disease Models &amp; Mechanisms. 11 (10), (2018).</li><li>Zhang, Y., Zhang, Z., Ge, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+efficient+platform+for+generating+somatic+point+mutations+with+germline+transmission+in+the+zebrafish+by+CRISPR/Cas9-mediated+gene+editing.">An efficient platform for generating somatic point mutations with germline transmission in the zebrafish by CRISPR/Cas9-mediated gene editing.</a> The Journal of Biological Chemistry. 293 (17), 6611-6622 (2018).</li><li>Vornanen, M., Hassinen, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+heart+as+a+model+for+human+cardiac+electrophysiology.">Zebrafish heart as a model for human cardiac electrophysiology.</a> Channels. 10 (2), 101-110 (2016).</li><li>Nemtsas, P., Wettwer, E., Christ, T., Weidinger, G., Ravens, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adult+zebrafish+heart+as+a+model+for+human+heart?+An+electrophysiological+study.">Adult zebrafish heart as a model for human heart? 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The engineering of precise gene edits using CRISPR-Cas9 is challenged by the low efficiencies of HDR mechanisms and their efficient detection. Here, a CRISPR-Cas9-based two-sgRNA exon replacement approach is described that produces precise edits in zebrafish with straightforward visual detection of positive edits. The efficacy of this approach is demonstrated by generating precise edits in the zkcnh6a gene. This paper shows how cardiac function in gene-edited zebrafish larvae may be assessed using non-invasive p...

CRISPR-based gene editing strategies in animal models enable the study of genetically heritable disease, development, and toxicology at the whole-organism level1,2,3. Zebrafish provide a powerful model that is closer in numerous physiological aspects to humans than murine or human-derived cell models4. An extensive array of genetic tools and strategies have been used in zebrafish for both forward5 and reverse genetic screening6. Comprehensive genetic mapping and annotation in zebrafish have facilitated gene-editing approaches as a primary technique to engineer targeted gene knockouts (KOs) and precise knock-ins (KIs)7.
Despite this, generating precise KI edits in zebrafish is limited by low efficiencies and the difficulty of accurate detection. Although transcription factor-like effector nucleases (TALENs) have been successfully used and optimized for KIs8, CRISPR provides an improved gene-editing strategy with simpler sgRNA targeting. Numerous studies have used CRISPR to generate precise KIs in zebrafish9,10,11,12,13,14,15,16,17,18,19,20, although these edits generated through CRISPR-mediated homology-directed repair (HDR) tend to be inefficient with low intrinsic success rates that require genotyping as a primary screen9,10,14,21. This demonstrates the need for an efficient KI CRISPR system in zebrafish, as well as a reliable high-throughput system for detecting precise edits.
The goal of this study was to describe a platform for generating a precise cardiac gene KI in zebrafish hearts with simple and high-throughput detection of successful edits. A CRISPR-Cas9-based two-sgRNA exon replacement approach is described, which is based on a TALEN approach8. This approach involves excision of the target sequence using two-sgRNA guides and replacement with an exogenous template sequence that contains the KI of interest as well as a genetically encoded intronic reporter gene (Figure 1). The integration of a genetically encoded fluorescent reporter within the target gene intronic sequence enables the efficient detection of positive edits. A phenotyping platform is then described for assessing cardiac electrical function in zebrafish larvae for non-invasive characterization of the gene variants associated with inherited LQTS, a cardiac electrical disorder that predisposes individuals to sudden cardiac death.
These approaches will enhance the access to and use of zebrafish KI gene edits to model inherited diseases and address biological and physiological questions, such as mapping gene expression patterns, and developmental regulation. Since zebrafish hearts better parallel human cardiac electrophysiological characteristics than murine models, they may be particularly attractive as a genetically tractable system for cardiac disease modeling7,22,23.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
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			<td>Program</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CRISPOR</td>
			<td>TEFOR Infrastructure</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ENSEMBL</td>
			<td>European Bioinformatics Institute</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ImageJ</td>
			<td>National Institutes of Health (NIH)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Micro-Manager</td>
			<td>Open Source (Github)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>NEBiocalculator</td>
			<td>New England Biolabs (NEB)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>EQUIPMENT</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>24-well Plate</td>
			<td>VWR</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>25 mm Petri Dish</td>
			<td>VWR</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Blackfly USB3 Camera</td>
			<td>Teledyne FLIR</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>C1000 Thermal Cycler</td>
			<td>Bio-Rad</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge 5415C</td>
			<td>Eppendorf</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>EZNA Gel Extraction Kit</td>
			<td>Omega Biotek</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MAXIscript T7 Transcription Kit</td>
			<td>Invitrogen</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MaxQ 5000 Incubator</td>
			<td>Barnstead Lab Line</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Miniprep Kit</td>
			<td>Qiagen</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>mMessage mMachine T7 Ultra Transcription Kit</td>
			<td>Invitrogen</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ND1000 Spectrophotometer</td>
			<td>Nanodrop</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PCR Purification Kit</td>
			<td>Qiagen</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PLI 100A Picoinjector</td>
			<td>Harvard Apparatus</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PowerPac Basic Power Supply</td>
			<td>Bio-Rad</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Stemi 305 Steroscope</td>
			<td>Zeiss</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Wide Mini Sub Cell GT Electrophoresis System</td>
			<td>Bio-Rad</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ZebTec Zebrafish Housing System</td>
			<td>Tecniplast</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>SERVICES</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Gene Synthesis</td>
			<td>Genewiz</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sanger Sequencing</td>
			<td>Genewiz</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>REAGENTS</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>10&#946; Competent Cells</td>
			<td>NEB</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>10X PCR Buffer</td>
			<td>Qiagen</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>100 mM Nucleotide Mixture</td>
			<td>ABM</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ampicillin</td>
			<td>Sigma</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>BamHI Endonuclease w/ buffer</td>
			<td>NEB</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>BsaI Endonuclease w/ buffer</td>
			<td>NEB</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>DR274 Plasmid (XL1 Blue bacterial agar stab)</td>
			<td>Addgene</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>EcoRI Endonuclease w/ buffer</td>
			<td>NEB</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glycerol</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>HindIII Endonuclease w/ buffer</td>
			<td>NEB</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Kanamycin</td>
			<td>Sigma</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Methylene Blue</td>
			<td>Sigma</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MLM3613 Plasmid (XL1 Blue bacterial agar stab)</td>
			<td>Addgene</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MS-222 (Tricaine)</td>
			<td>Sigma</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>pKHR5 Plasmid (DH5&#945; bacterial agar stab)</td>
			<td>Addgene</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PmeI Endonuclease w/ buffer</td>
			<td>NEB</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>SalI Endonuclease w/ buffer</td>
			<td>NEB</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Hydroxide</td>
			<td>Sigma</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>T4 Ligase w/ buffer</td>
			<td>Sigma</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Taq Polymerase</td>
			<td>Qiagen</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>TE Buffer</td>
			<td>Sigma</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Tris Hydrochloride</td>
			<td>Sigma</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>XhoI Endonuclease w/ buffer</td>
			<td>NEB</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>RECIPES</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Solution</td>
			<td>Component</td>
			<td>Supplier</td>
			<td></td>
		</tr>
		<tr>
			<td>Annealing Buffer (pH 7.5-8.0)</td>
			<td>10 mM Tris</td>
			<td>Sigma</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>50 mM NaCl</td>
			<td>Sigma</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>1 mM EDTA</td>
			<td>Sigma</td>
			<td></td>
		</tr>
		<tr>
			<td>E3 Media (pH 7.2)</td>
			<td>5 mM NaCl</td>
			<td>Sigma</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>0.17 mM KCl</td>
			<td>Sigma</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>0.33 mM CaCl2</td>
			<td>Sigma</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>0.33 mM MgSO4</td>
			<td>Sigma</td>
			<td></td>
		</tr>
		<tr>
			<td>Injection Buffer (pH 7.5)</td>
			<td>20 mM HEPES</td>
			<td>Sigma</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>150 mM KCl</td>
			<td>Sigma</td>
			<td></td>
		</tr>
	</tbody>
</table>

crispr-cas9-mediated precise knock-in edits in zebrafish hearts

Clustered regularly interspaced short palindromic repeats (CRISPR) in animal models enable precise genetic manipulation for the study of physiological phenomena. Zebrafish have been used as an effective genetic model to study numerous questions related to heritable disease, development, and toxicology at the whole-organ and -organism level. Due to the well-annotated and mapped zebrafish genome, numerous tools for gene editing have been developed. However, the efficacy of generating and ease of detecting precise knock-in edits using CRISPR is a limiting factor. Described here is a CRISPR-Cas9-based knock-in approach with the simple detection of precise edits in a gene responsible for cardiac repolarization and associated with the electrical disorder, Long QT Syndrome (LQTS). This two-single-guide RNA (sgRNA) approach excises and replaces the target sequence and links a genetically encoded reporter gene. The utility of this approach is demonstrated by describing non-invasive phenotypic measurements of cardiac electrical function in wild-type and gene-edited zebrafish larvae. This approach enables the efficient study of disease-associated variants in a whole organism. Furthermore, this strategy offers possibilities for the insertion of exogenous sequences of choice, such as reporter genes, orthologs, or gene editors.

The successful use of this two-sgRNA exon replacement CRISPR approach is highlighted by the introduction and simple detection of a precise edit to engineer the LQTS-associated variant, R56Q, in the zkcnh6a gene in zebrafish. Figure 6 shows a representative 3 dpf larvae injected at the one-cell embryo stage with CRISPR components as described above. Figure 6A shows the presence of the YFP mVenus reporter gene expression in the eye lens as a positive repo...

Watch this Scientific Journal Video about CRISPR-Cas9-Mediated Precise Knock-In Edits in Zebrafish Hearts at JoVE.com

CRISPR-Cas9-Mediated Precise Knock-In Edits in Zebrafish Hearts

This protocol describes an approach to facilitate precise knock-in edits in zebrafish embryos using CRISPR-Cas9 technology. A phenotyping pipeline is presented to demonstrate the applicability of these techniques to model a Long QT Syndrome-associated gene variant.

Clustered regularly interspaced short palindromic repeats (CRISPR) in animal models enable precise genetic manipulation for the study of physiological ...

Our protocol provides a pipeline for both precise integration and simple visual detection of CRISPR edits, as well as an example of post-edit phenotyping. The main advantage of this approach is ease of CRISPR edit detection, allowing for efficient and simplified identification of successful edits. This approach allows for simple generation and characterization of disease-associated variants in zebrafish.Here, we are examining long QT syndrome. Follow-up studies on pharmaceutical therapies are then feasible to explore. Begin by designing two sgRNA guides to excise the knock-in target site sequence by identifying the zebrafish ortholog for the gene of interest.Use Ensembl to locate the target site within the gene sequence of interest, including the two kilobase flanking sequence used to make the template. Use a design software tool such as CRISPOR and select Danio rerio species and Cas enzyme. For the two sgRNA approach, choose one sgRNA before the target exon and a second sgRNA within the immediate downstream intron.Ensure that the selected sgRNAs have high specificity and low predicted off-target binding. Use CRISPOR rankings to identify guides with minimal off-target binding. Now identify the most likely potential off-target sites for PCR-based Sanger sequencing genotyping.After selecting the two sgRNAs, obtain the reverse complement for each, two complementary oligos that precede the mutation and two complementary oligos downstream of the mutation. Add compatible restriction sites on each oligo to incorporate the guide in the plasmid of choice. For the integration of the selected guide sequences into the DR274 plasmid, create an overhang using a five prime BSA1 restriction site and engineer the BSA1 recognition sites at the five prime ends of the guides.Next, design the exogenous template for HDR in zebrafish by choosing two sequence fragments that will flank M venous YFP reporter gene housed in the PKHR5 plasmid. Divide the template into two segments to insert it on either side of the M venous YFP reporter gene. Ensure that the split site is in an intron to avoid cutting the coding sequence.Incorporate all the required modifications mentioned in the manuscript to facilitate cloning into the PKHR5 plasmid. Inject the embryos in the one cell stage at approximately 40 minutes post-fertilization. After three days of fertilization, anesthetize zebrafish larva by transferring it to a 25 millimeter Petri dish containing 0.3%MS222 until they lose their self-righting reflex.Once anesthetized, transfer the larva into the well of a 24-well plate. Using a microscope capable of detecting GFP or YFP, screen for reporter gene fluorescence in the eyes of the larva. After capturing the images of the larva, document the presence or absence of the reporter gene expression.To confirm accurate and precise HDR gene editing, perform genotyping by first anesthetizing the larva three days post-fertilization as demonstrated earlier and isolating the genomic DNA from the tail clip using the hotshot method. Add the excised tail clip to 15 microliters of 25 millimolar sodium hydroxide and incubate it 95 degrees Celsius for 20 minutes, then neutralize with 1.5 microliters of tris hydrochloric acid. Centrifuge at 13, 800 times G for 30 seconds, and retain the supernatant containing the extracted genomic DNA.Recover the larva in E3 media and return it to the housing system if further study is intended. Using the extracted genomic DNA as a template, perform PCR-based Sanger sequencing of on-target and potential off-target sites. Ensure the on-target primer design captures the mutation site and the closest sgRNA binding site.Design a separate sequencing primer to detect the transition from the inserted homology arm and the target gene to confirm integration into the gene of interest. Design primers to sequence the top three potential off-target sites. Compile on and off-target genotyping, heart rate, pericardial dimensions, ECG phenotyping, and reporter gene data identifiable for each zebrafish.The YFP M venous reporter gene expression was observed in the islands, acting as a positive reporter of successful template integration. Reporter gene positive fish were found to have the precise edit G2A which introduces the R56Q variant into ZKCNH6A. The measurement of pericardial dimensions as a ratio of the eye area is shown here.A trend of bradycardia with increasing pericardial edema associated with the disorders of cardiac repolarization in zebrafishes was observed in the R56Q gene edited larvae. ECG recording from a zebrafish larva heart three days post-fertilization is shown here. The most important aspect of this approach is guide design, as is often the case in CRISPR procedures.Effective guides are essential for successful CRISPR edits. Alternative CRISPR methods can be followed allowing for more precision or other functionalities. Additionally, large genes can be inserted such as a prime editing complex providing an in vivo inducible editing system.

crispr-cas9-mediated-precise-knock-in-edits-in-zebrafish-hearts

Research

JoVE Journal

Biology

Optical Mapping of Langendorff-perfused Rat Hearts

Optical mapping of the cardiac surface with voltage-sensitive fluorescent dyes has become an important tool to investigate electrical excitation in experimental models that range in scale from cell cultures to whole-organs[1, 2]. Using state-of-the-art optical imaging systems, generation and propagation of action potentials during normal cardiac rhythm or throughout initiation and maintenance of arrhythmias can be visualized almost instantly[1]. The latest commercially-available systems can provide information at exceedingly high spatiotemporal resolutions and were based on custom-built equipment initially developed to overcome the obstacles imposed by more conventional electrophysiological methods[1]. Advancements in high-resolution and high-speed complementary metal-oxide-semiconductor (CMOS) cameras and intensely-bright, light-emitting diodes (LEDs) as well as voltage-sensitive dyes, optics, and filters have begun to make electrical signal acquisition practical for cardiovascular cell biologists who are more accustomed to working with microscopes. Although the newest generation of CMOS cameras can acquire 10,000 frames per second on a 16,384 pixel array, depending on the type of sample preparation, long-established fluorescence acquisition technologies such as photodiode arrays, laser scanning systems, and cooled charged-coupled device (CCD) cameras still have some distinct advantages with respect to dynamic range, signal-to-noise ratio, and quantum efficiency[1, 3]. In the present study, Lewis rat hearts were perfused ex vivo with a crystalloid perfusate (Krebs-Henseleit solution) at 37&deg;C on a modified Langendorff apparatus. After a 20 minute stabilization period, the hearts were intermittently perfused with 11 mMol/L 2,3-butanedione monoxime to eliminate contraction-associated motion during image acquisition. For optical mapping, we loaded hearts with the fast-response potentiometric probe di-8-ANEPPS[4] (5 &mu;Mol/L) and briefly illuminated the preparation with 475&plusmn;15 nm excitation light. During a typical 2 second period of illumination, &gt;605 nm light emitted from the cardiac preparation was imaged with a high-speed CMOS camera connected to a horizontal macroscope. For this demonstration, hearts were paced at 300 beats per minute with a coaxial electrode connected to an isolated electrical stimulation unit. Simultaneous bipolar electrographic recordings were acquired and analyzed along with the voltage signals using readily-available software. In this manner, action potentials on the surface of Langendorff-perfused rat hearts can be visualized and registered with electrographic signals.

This article describes a high temporal and spatial resolution technique to optically image action potential movement on the surface of Langendorff-perfused rat hearts using a potentiometric dye (di-8-ANEPPS).

Optical mapping of the cardiac surface with voltage-sensitive fluorescent dyes has become an important tool to investigate electrical excitation in ...

In Situ Hybridization for the Precise Localization of Transcripts in Plants

With the advances in genomics research of the past decade, plant biology has seen numerous studies presenting large-scale quantitative analyses of gene expression. Microarray and next generation sequencing approaches are being used to investigate developmental, physiological and stress response processes, dissect epigenetic and small RNA pathways, and build large gene regulatory networks1-3. While these techniques facilitate the simultaneous analysis of large gene sets, they typically provide a very limited spatiotemporal resolution of gene expression changes. This limitation can be partially overcome by using either profiling method in conjunction with lasermicrodissection or fluorescence-activated cell sorting4-7. However, to fully understand the biological role of a gene, knowledge of its spatiotemporal pattern of expression at a cellular resolution is essential. Particularly, when studying development or the effects of environmental stimuli and mutants can the detailed analysis of a gene's expression pattern become essential. For instance, subtle quantitative differences in the expression levels of key regulatory genes can lead to dramatic phenotypes when associated with the loss or gain of expression in specific cell types. 
Several methods are routinely used for the detailed examination of gene expression patterns. One is through analysis of transgenic reporter lines. Such analysis can, however, become time-consuming when analyzing multiple genes or working in plants recalcitrant to transformation. Moreover, an independent validation to ensure that the transgene expression pattern mimics that of the endogenous gene is typically required. Immunohistochemical protein localization or mRNA in situ hybridization present relatively fast alternatives for the direct visualization of gene expression within cells and tissues. The latter has the distinct advantage that it can be readily used on any gene of interest. In situ hybridization allows detection of target mRNAs in cells by hybridization with a labeled anti-sense RNA probe obtained by in vitro transcription of the gene of interest. 
Here we outline a protocol for the in situ localization of gene expression in plants that is highly sensitivity and specific. It is optimized for use with paraformaldehyde fixed, paraffin-embedded sections, which give excellent preservation of histology, and DIG-labeled probes that are visualized by immuno-detection and alkaline-phosphatase colorimetric reaction. This protocol has been successfully applied to a number of tissues from a wide range of plant species, and can be used to analyze expression of mRNAs as well as small RNAs8-14.

In Situ Hybridization for the Precise Localization of Transcripts in Plants

The in situ hybridization protocol described here allows a direct localization of mRNA and small RNA expression at the cellular level with high sensitivity and specificity. The procedure is optimized for paraffin-embedded plant tissue sections, is applicable to a wide range of plants and tissues, and can be completed within ten days.

With the advances in genomics research of the past decade, plant biology has seen numerous studies presenting large-scale quantitative analyses of gene ...

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal neurons. In contrast, the adult zebrafish retina possesses the robust ability to spontaneously regenerate any neuronal class that is lost in a variety of different retinal damage models, including retinal puncture, chemical ablation, concentrated high temperature, and intense light treatment 1-8. Our lab extensively characterized regeneration of photoreceptors following constant intense light treatment and inner retinal neurons after intravitreal ouabain injection 2, 5, 9. In all cases, resident M&uuml;ller glia re-enter the cell cycle to produce neuronal progenitors, which continue to proliferate and migrate to the proper retinal layer, where they differentiate into the deficient neurons. 
	We characterized five different stages during regeneration of the light-damaged retina that were highlighted by specific cellular responses. We identified several differentially expressed genes at each stage of retinal regeneration by mRNA microarray analysis 10. Many of these genes are also critical for ocular development. To test the role of each candidate gene/protein during retinal regeneration, we needed to develop a method to conditionally limit the expression of a candidate protein only at times during regeneration of the adult retina. 
	Morpholino oligos are widely used to study loss of function of specific proteins during the development of zebrafish, Xenopus, chick, mouse, and tumors in human xenografts 11-14. These modified oligos basepair with complementary RNA sequence to either block the splicing or translation of the target RNA. Morpholinos are stable in the cell and can eliminate or "knockdown" protein expression for three to five days 12. 
	Here, we describe a method to efficiently knockdown target protein expression in the adult zebrafish retina. This method employs lissamine-tagged antisense morpholinos that are injected into the vitreous of the adult zebrafish eye. Using electrode forceps, the morpholino is then electroporated into all the cell types of the dorsal and central retina. Lissamine provides the charge on the morpholino for electroporation and can be visualized to assess the presence of the morpholino in the retinal cells. 
	Conditional knockdown in the retina can be used to examine the role of specific proteins at different times during regeneration. Additionally, this approach can be used to study the role of specific proteins in the undamaged retina, in such processes as visual transduction and visual processing in second order neurons.

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

A method to conditionally knockdown a target protein&#x2019;s expression in the adult zebrafish retina is described, which involves intravitreally injecting antisense morpholinos and electroporating them into the retina. The resulting protein is knocked down for several days, which allows testing the protein&#x2019;s role in the regenerating or intact retina.

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal ...

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration of adult tissues, such as the heart1, retina2, spinal cord3, optic nerve4, sensory hair cells5, and fins6.
The zebrafish fin is a relatively simple appendage that is easily manipulated to study multiple stages in epimorphic regeneration. Classically, fin regeneration was characterized by three distinct stages: wound healing, blastema formation, and fin outgrowth. After amputating part of the fin, the surrounding epithelium proliferates and migrates over the wound. At 33 &deg;C, this process occurs within six hours post-amputation (hpa, Figure 1B)6,7. Next, underlying cells from different lineages (ex. bone, blood, glia, fibroblast) re-enter the cell cycle to form a proliferative blastema, while the overlying epidermis continues to proliferate (Figure 1D)8. Outgrowth occurs as cells proximal to the blastema re-differentiate into their respective lineages to form new tissue (Figure 1E)8. Depending on the level of the amputation, full regeneration is completed in a week to a month.
The expression of a large number of gene families, including wnt, hox, fgf, msx, retinoic acid, shh, notch, bmp, and activin-betaA genes, is up-regulated during specific stages of fin regeneration9-16. However, the roles of these genes and their encoded proteins during regeneration have been difficult to assess, unless a specific inhibitor for the protein exists13, a temperature-sensitive mutant exists or a transgenic animal (either overexpressing the wild-type protein or a dominant-negative protein) was generated7,12. We developed a reverse genetic technique to quickly and easily test the function of any gene during fin regeneration.
Morpholino oligonucleotides are widely used to study loss of specific proteins during zebrafish, Xenopus, chick, and mouse development17-19. Morpholinos basepair with a complementary RNA sequence to either block pre-mRNA splicing or mRNA translation. We describe a method to efficiently introduce fluorescein-tagged antisense morpholinos into regenerating zebrafish fins to knockdown expression of the target protein. The morpholino is micro-injected into each blastema of the regenerating zebrafish tail fin and electroporated into the surrounding cells. Fluorescein provides the charge to electroporate the morpholino and to visualize the morpholino in the fin tissue.
This protocol permits conditional protein knockdown to examine the role of specific proteins during regenerative fin outgrowth. In the Discussion, we describe how this approach can be adapted to study the role of specific proteins during wound healing or blastema formation, as well as a potential marker of cell migration during blastema formation.

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

We describe a method to conditionally knockdown the expression of a target protein during adult zebrafish fin regeneration. This technique involves micro-injecting and electroporating antisense oligonucleotide morpholinos into fin tissue, which allows testing the protein&#x2019;s role in various stages of fin regeneration, including wound healing, blastema formation, and regenerative outgrowth.

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration ...

Gavaging Adult Zebrafish

The zebrafish has become an important in vivo model in biomedical research. Effective methods must be developed and utilized to deliver compounds or agents in solutions for scientific research. Current methods for administering compounds orally to adult zebrafish are inaccurate due to variability in voluntary consumption by the fish. A gavage procedure was developed to deliver precise quantities of infectious agents to zebrafish for study in biomedical research. Adult zebrafish over 6 months of age were anesthetized with 150 mg/L of buffered MS-222 and gavaged with 5 &mu;l of solution using flexible catheter implantation tubing attached to a cut 22-G needle tip. The flexible tubing was lowered into the oral cavity of the zebrafish until the tip of the tubing extended past the gills (approximately 1 cm). The solution was then injected slowly into the intestinal tract. This method was effective 88% of the time, with fish recovering uneventfully. This procedure is also efficient as one person can gavage 20-30 fish in one hour. This method can be used to precisely administer agents for infectious diseases studies, or studies of other compounds in adult zebrafish.

The increasing use of zebrafish as an animal model requires the development of effective methods for the delivery of known quantities of compounds and solutions. The gavage procedure described below allows for the oral delivery of precise volumes of solution reliably, safely and efficiently to adult zebrafish.

The zebrafish has become an important in vivo model in biomedical research. Effective methods must be developed and utilized to deliver compounds or ...

Cell Labeling and Injection in Developing Embryonic Mouse Hearts

Testing the fate of embryonic or pluripotent stem cell-derivatives in in vitro protocols has led to controversial outcomes that do not necessarily reflect their in vivo potential. Preferably, these cells should be placed in a proper embryonic environment in order to acquire their definite phenotype. Furthermore, cell lineage tracing studies in the mouse after labeling cells with dyes or retroviral vectors has remained mostly limited to early stage mouse embryos with still poorly developed organs. To overcome these limitations, we designed standard and ultrasound-mediated microinjection protocols to inject various agents in targeted regions of the heart in mouse embryos at E9.5 and later stages of development. &#160;Embryonic explant or embryos are then cultured or left to further develop in utero. These agents include fluorescent dyes, virus, shRNAs, or stem cell-derived progenitor cells. Our approaches allow for preservation of the function of the organ while monitoring migration and fate of labeled and/or injected cells. These technologies can be extended to other organs and will be very helpful to address key biological questions in biology of development.

We describe a series of methods to inject dyes, DNA vectors, virus, and cells in order to monitor both cell fate and phenotype of endogenous and grafted cells derived from embryonic or pluripotent cells within mouse embryos at embryonic day (E)9.5 and later stages of development.

Testing the fate of embryonic or pluripotent stem cell-derivatives in in vitro protocols has led to controversial outcomes that do not necessarily reflect ...

Recovery of Adult Zebrafish Hearts for High-throughput Applications

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and purification of RNA, DNA, and proteins. All of these applications demand the rapid recovery of significant numbers of zebrafish hearts to avoid gene regulatory, metabolic, and other changes that begin after death. Adult zebrafish hearts are also required for studying heart structure for a variety of mutants and for studying heart regeneration. However, the traditional zebrafish heart dissection is slow and difficult and requires specialized tools, making large-scale dissection of adult zebrafish hearts tedious. Traditional methods also harbor the risk of damaging the heart during the dissection. Here, we describe a method for dissection of adult zebrafish hearts that is fast, reproducible, and preserves heart architecture. Furthermore, this method does not require specialized tools, is painless for the zebrafish, can be performed on fresh or fixed specimens, and can be performed on zebrafish as young as one month old. The approach described expands the use of adult zebrafish for cardiovascular research.

Use of zebrafish for cardiovascular research is expanding towards research on adult hearts. For these applications, quick and simple isolation of cardiac tissues is key to avoid post-mortem changes and to obtain an adequate number of samples. Here, we describe a fast and reproducible method for dissecting adult zebrafish hearts.

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and ...

Nephrotoxin Microinjection in Zebrafish to Model Acute Kidney Injury

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid decline in renal function and development of the clinical syndrome known as acute kidney injury (AKI). Pharmacological agents used to treat medical circumstances ranging from bacterial infection to cancer, when administered individually or in combination with other drugs, can initiate AKI. Zebrafish are a useful animal model to study the chemical effects on renal function in vivo, as they form an embryonic kidney comprised of nephron functional units that are conserved with higher vertebrates, including humans. Further, zebrafish can be utilized to perform genetic and chemical screens, which provide opportunities to elucidate the cellular and molecular facets of AKI and develop therapeutic strategies such as the identification of nephroprotective molecules. Here, we demonstrate how microinjection into the zebrafish embryo can be utilized as a paradigm for nephrotoxin studies.

Renal injuries incurred from nephrotoxins, which include drugs ranging from antibiotics to chemotherapeutics, can result in complex disorders whose pathogenesis remains incompletely understood. This protocol demonstrates how zebrafish can be used for disease modeling of these conditions, which can be applied to the identification of renoprotective measures.

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid ...

Local Field Fluorescence Microscopy: Imaging Cellular Signals in Intact Hearts

In the heart, molecular signaling studies are usually performed in isolated myocytes. However, many pathological situations such as ischemia and arrhythmias can only be fully understood at the whole organ level. Here, we present the spectroscopic technique of local field fluorescence microscopy (LFFM) that allows the measurement of cellular signals in the intact heart. The technique is based on a combination of a Langendorff perfused heart and optical fibers to record fluorescent signals. LFFM has various applications in the field of cardiovascular physiology to study the heart under normal and pathological conditions. Multiple cardiac variables can be monitored using different fluorescent indicators. These include cytosolic [Ca2+], intra-sarcoplasmic reticulum [Ca2+] and membrane potentials. The exogenous fluorescent probes are excited and the emitted fluorescence detected with three different arrangements of LFFM epifluorescence techniques presented in this paper. The central differences among these techniques are the type of light source used for excitation and on the way the excitation light is modulated. The pulsed LFFM (PLFFM) uses laser light pulses while continuous wave LFFM (CLFFM) uses continuous laser light for excitation. Finally, light-emitting diodes (LEDs) were used as a third light source. This non-coherent arrangement is called pulsed LED fluorescence microscopy (PLEDFM).

In the heart, molecular events coordinate the electrical and contractile function of the organ. A set of local field fluorescence microscopy techniques presented here enables the recording of cellular variables in intact hearts. Identifying mechanisms defining the cardiac function is critical in understanding how the heart works under pathological situations.

In the heart, molecular signaling studies are usually performed in isolated myocytes. However, many pathological situations such as ischemia and ...

Collecting Hemolymph from Small Arthropods

A Precise and Quantifiable Method for Collecting Hemolymph from Small Arthropods

Arthropods are known to transmit a variety of viruses of medical and agricultural importance through their hemolymph, which is essential for virus transmission. Hemolymph collection is the basic technology for studying virus-vector interactions. Here, we describe a novel and simple method for the quantitative collection of hemolymph from small arthropods using Laodelphax striatellus (the small brown planthopper, SBPH) as a research model, as this arthropod is the main vector of rice stripe virus (RSV). In this protocol, the process begins by gently pinching off one leg of the frozen arthropod with fine-tipped tweezers and pressing the hemolymph out of the wound. Then, a simple micropipette consisting of a capillary and a pipette bulb is used to collect the transudative hemolymph from the wound according to the principle of capillary forces. Finally, the collected hemolymph can be dissolved into a specific buffer for further study. This new method for collecting hemolymph from small arthropods is a useful and efficient tool for further research on arboviruses and vector-virus interactions.

Author Spotlight: A Precise and Quantifiable Method for Collecting Hemolymph from Small Arthropods  

We describe a method to collect quantifiable hemolymph efficiently from small arthropods for subsequent analysis.