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: Figure 2 provides a summary overview of the steps to engineer precise edits using the two-sgRNA CRISPR-Cas9 approach.</li>
	<li>Next, use a design software tool, such as CRISPOR24, which includes selection for Danio rerio as a species and of the Cas enzyme to be used. 
	NOTE: The gene of interest for this study was zkcnh6a (Ensembl Transcript ID: ENSDART00000090809.6; UniProt Protein ID: B3DJX4), and the target mutation was amino acid, R56Q.
	<ol>
		<li>For the two-sgRNA approach, choose one sgRNA location that precedes the target exon and a second sgRNA that is located within the immediate downstream intron.</li>
		<li>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. Do not consider guides with no mismatches in the seed sequence of potential off-targets.</li>
		<li>Identify the most likely potential off-target sites (based on CRISPOR scores, select the top three exon potential off-target sites) for PCR-based Sanger sequencing genotyping in step 6.1.3.</li>
		<li>Once the two-sgRNAs have been selected, obtain the reverse complement for each, such that there are four oligonucleotides to be used: two complementary oligos that precede the mutation and two complementary oligos that are downstream.</li>
		<li>On each of the four oligos, add compatible restriction sites for incorporation into a guide plasmid of choice; for integration into the DR274 plasmid, use a 5&#39; BsaI restriction site to create an overhang. Ensure that the Bsa1 recognition site is engineered at the 5&#39; end of the guide selected from CRISPOR and the Bsa1 recognition site is engineered at the 5&#39; end of the complementary strands to ensure correct orientation of the guides in the DR274 plasmid (see Figure 3).</li>
	</ol>
	</li>
	<li>To design the exogenous template used for HDR in zebrafish (Figure 1), choose two sequence fragments that will flank the mVenus YFP reporter gene housed in the pKHR5 plasmid. 
	NOTE: The intended modification/edit can be included in either the upstream or downstream fragment.
	<ol>
		<li>Using Ensembl, locate the target site within the gene sequence of interest, including approximately the 2 kb flanking sequence (homology arm), which will be used to make the template. 
		NOTE: Homology arms may be symmetrical or asymmetrical25,26 and approximately 1 kb each, upstream and downstream of the target site.</li>
		<li>Divide the template into two segments that will be inserted either side of the mVenus YFP reporter gene (see Figure 4). Ensure that the split site is in an intron so that the coding sequence is not interrupted. 
		NOTE: If the gene is well characterized, check for functional roles in the intron, such as splicing sites or regulatory regions. Regions close to the 5&#39; or 3&#39; ends are more often involved in mRNA splicing.</li>
		<li>Incorporate modifications into the template sequence that include i) silent mutations in the guide protospacer adjacent motif (PAM) or seed sequence (be aware of alternate PAM sites that the Cas enzyme might target) to prevent Cas enzyme recutting; ii) the modification of interest; iii) the creation of restriction endonuclease sites to facilitate cloning into the pKHR5 plasmid, which contains the mVenus YFP reporter gene (see Figure 3). 
		NOTE: In this study, the first template segment contained XhoI upstream of the R56Q mutation site and SalI downstream, while the second template segment had EcoRI upstream of the guide target sequence and BamHI downstream. If any of the selected restriction sites are present within the template sequence, mutations to silence these will be required, or alternate approaches such as Gibson Assembly can be used.</li>
	</ol>
	</li>
</ol>
2. Preparation of CRISPR components for embryo microinjection
<ol>
	<li>Prepare the Cas9 for microinjection 1 week prior to the microinjections. Use Cas9 protein or prepare Cas9 mRNA via in vitro transcription. 
	NOTE: In this study, Cas9 mRNA was used, since efficiencies tended to be higher.
	<ol>
		<li>Amplify bacterial cultures of commercially available XL1 Blue bacterial agar stab (containing Cas9 plasmid) using an appropriate antibiotic such as ampicillin. Use 675 &#181;L of liquid culture (with 325 &#181;L of glycerol) to create a backup glycerol stock for long-term storage at &#8722;80 &#176;C.</li>
		<li>Use the remainder of the liquid culture for a miniprep purification, according to the protocol provided with the miniprep kit. Resuspend the final purified DNA in 50 &#181;L of the provided elution buffer. Quantify the product via a spectrophotometer to examine the yield and purity.</li>
		<li>Linearize 2 &#181;g of the purified DNA via restriction digest using an appropriate restriction enzyme and using the appropriate buffer and incubation time as listed for the enzyme of interest.</li>
		<li>Purify the linearized plasmid using a PCR Purification Kit, resuspending it in 30 &#181;L of the provided elution buffer.</li>
		<li>After quantifying the product, use this as a template for in vitro transcription using the appropriate transcription kit for the promoter of interest. Follow the provided protocol and purify via lithium chloride precipitation27. Resuspend the purified RNA in 10 &#181;L of nuclease-free H2O and quantify it before storing at &#8722;20 &#176;C for use in the microinjection mix.</li>
	</ol>
	</li>
	<li>Prepare the two sgRNA guides.
	<ol>
		<li>Prepare the sgRNA plasmid with a scaffold by amplifying bacterial cultures from the commercially available XL1 Blue bacterial agar stab (see Table of Materials for details) in the same way as MLM3613 above (step 2.1.1), except use kanamycin instead of ampicillin.</li>
		<li>Anneal the two pairs of complementary single-stranded oligonucleotides (ssODNs) for the sgRNA guides designed above by first resuspending the ssODNs in 1x annealing buffer to a concentration of 100 &#181;M.</li>
		<li>In separate reactions for each of the two sgRNAs, anneal the pair of complementary ssODNs using a thermal cycler. Mix 2 &#181;g of each complementary ssODN pair with 50 &#181;L of annealing buffer and incubate at 95 &#176;C for 2 min, then cool to 25 &#176;C over 45 min.</li>
		<li>Digest a commercially available plasmid that contains a gRNA scaffold. Digest 2 &#181;g of the DR274 plasmid using 1 &#181;L of BsaI, 2 &#181;L of appropriate buffer, and ddH2O to 20 &#181;L at 37 &#176;C for 1 h. Confirm linearization (optional: purify using a PCR Purification Kit) using gel electrophoresis28.</li>
		<li>In two separate ligation reactions (one for each sgRNA), ligate the annealed ssODNs with the linearized DR274 plasmid. Use a molar insert:vector ratio of 3:1, calculating the appropriate mass through an online ligation calculator. Mix the required mass of the insert and vector with 1 &#181;L of T4 DNA ligase, 2 &#181;L of ligation buffer, and ddH2O to 12 &#181;L and incubate at room temperature for 12 h.</li>
		<li>Transform 2 &#181;L of the ligated product into appropriate competent cells (such as 10&#946; cells) using standard approaches, and then amplify and purify the product using a commercially available Miniprep Kit. Optional: create a glycerol stock of this product.</li>
		<li>Transcribe the two sgRNAs by linearizing 2 &#181;g of each guide using a 3&#39; downstream restriction site that is as close to the end of the space sequence as possible. For the DR274 plasmid, linearize with HindIII, and then purify the RNA template using a PCR Purification Kit, resuspending in 30 &#181;L of the elution buffer.</li>
		<li>Transcribe the two guides using an RNA transcription kit. Follow the manufacturer&#39;s protocol and purify via lithium chloride precipitation27. Resuspend the two purified sgRNA guides in 10 &#181;L of nuclease-free H2O, quantify, and store at &#8722;20 &#176;C for use in the microinjection mix. 
		NOTE: The RNA transcription kit cannot incorporate a 5&#39; cap or poly-A tail.</li>
	</ol>
	</li>
	<li>Prepare the double-stranded, exogenous HDR reporter template. 
	NOTE: The template is synthesized in two parts, one upstream and one downstream of the mVenus YFP reporter gene. These two segments are synthetic constructs ordered through a commercial provider and then ligated into the pKHR5 (which contains mVenus YFP) plasmid sequentially.
	<ol>
		<li>Prepare the pKHR5 plasmid by amplifying bacterial cultures from the commercially available DH5&#945; bacterial strain (see Table of Materials for details) in the same way as MLM3613 above (step 2.1.1). 
		NOTE: The pKHR5 plasmid contains the mVenus YFP reporter gene sequence.</li>
		<li>Resuspend the two template segments designed above to 100 &#181;M in TE buffer and then transform into 10&#946; cells.</li>
		<li>Digest the first template segment (the one upstream of the mVenus YFP reporter gene) and the pKHR5 plasmid using the restriction enzymes selected in step 1.3.3. 
		NOTE: Sequential digest of pKHR5 is necessary due to the close proximity of the selected restriction sites in the MCS.</li>
		<li>To prepare the pKHR5 plasmid for the upstream template segment, digest 4 &#181;g of pKHR5 (as per step 2.2.4) with SalI and then purify using a PCR purification kit. Resuspend in 30 &#181;L of ddH2O and use this as a template for the second digestion reaction with XhoI. Purify the product using the PCR Purification Kit.</li>
		<li>Prepare the first template segment by digesting 2 &#181;g of the template segment (step 2.3.2), 1 &#181;L of XhoI, 1 &#181;L of SalI, 2 &#181;L of appropriate buffer, and ddH2O to 20 &#181;L for 1 h at 37 &#176;C and gel-purify the product.</li>
		<li>Ligate the upstream template segment from step 2.3.5 into the prepared pKHR5 using the reaction conditions described in step 2.2.5. Transform the ligated product into competent 10&#946; cells, amplify, and purify using a miniprep (optional: create a glycerol stock of this product).</li>
		<li>Use the ligated product from step 2.3.6 (which contains the first template segment ligated into pKHR5 plasmid upstream of the mVenus YFP reporter gene) and digest to prepare for the second (downstream of the mVenus reporter) template segment. Digest 4 &#181;g of the ligated product (from step 2.3.6, as in step 2.2.4) with BamHI, and then purify using a PCR purification kit. Resuspend in 30 &#181;L of ddH2O and use this as the template for the second digest reaction with EcoRI. Purify the product using the PCR Purification Kit.</li>
		<li>Prepare the second template segment by digesting 2 &#181;g of the template segment (step 2.3.2), 1 &#181;L of BamHI, 1 &#181;L of EcoRI, 2 &#181;L of appropriate buffer, and ddH2O to 20 &#181;L for 1 h at 37 &#176;C and gel-purify the product.</li>
		<li>Ligate the downstream template segment into the prepared pKHR5 (from step 2.3.7) using the reaction conditions described in step 2.2.5. Transform the ligated product into competent cells, amplify, and purify using a miniprep kit. Create a glycerol stock of this final product, which contains both template segments ligated either side of the mVenus YFP reporter gene within pKHR5.</li>
	</ol>
	</li>
	<li>Prepare the microinjection mix using the Cas9 mRNA, two sgRNAs, and the exogenous HDR reporter template.
	<ol>
		<li>Mix 200 ng/&#181;L of Cas9 mRNA, 100 ng/&#181;L of each sgRNA, and 200 ng/&#181;L of exogenous HDR reporter template in 1x injection buffer to a final volume of 20 &#181;L.</li>
		<li>Store the microinjection mix at &#8722;20 &#176;C and discard the unused mix after three freeze-thaw cycles. 
		NOTE: Use 4 nL of this microinjection mix for microinjection into the yolk sac of each embryo.</li>
	</ol>
	</li>
</ol>
3. Breeding of zebrafish and embryo microinjection
NOTE: Protocols for zebrafish breeding and the microinjection of single-cell embryos have been described previously29,30,31.
<ol>
	<li>For breeding, use zebrafish of the AB strain and that are 6-12 months of age. Inject the embryos in the one-cell stage at approximately 40 min post fertilization (see Figure 5). 
	NOTE: The biological sex of the injected embryos was not known; sexual dimorphism is not evident until approximately 3 months of age32.</li>
</ol>
4. Reporter gene screening of CRISPR-Cas9-edited larval zebrafish
<ol>
	<li>Visualize YFP integration in zebrafish larvae following the microinjection of CRISPR-Cas9 components to screen for successful HDR edits.
	<ol>
		<li>In a 25 mm Petri dish, anesthetize 24 zebrafish larvae at 3 days post fertilization (dpf) in 0.3% tricaine methane sulfonate (MS-222, buffered to pH 7.0-7.4 with HEPES and sodium hydroxide) until they lose their self-righting reflex (typically 1-2 min). Once anesthetized, transfer each larva into an individual well of a 24-well plate.</li>
		<li>Using a microscope capable of detecting GFP/YFP, screen for reporter gene fluorescence in the eyes of each individual larva.</li>
		<li>Capture images of each larva and document the presence or absence of reporter gene expression.</li>
	</ol>
	</li>
</ol>
5. Phenotyping of CRISPR-Cas9-edited larval zebrafish
<ol>
	<li>Following reporter gene screening, perform cardiac phenotyping (heart rate, pericardial dimensions, ECG) on each larva. Phenotype an equal number of reporter gene-positive and -negative larvae.
	<ol>
		<li>Use a CCD camera (e.g., blackfly USB3) and video and image recording software (e.g., Micromanager for ImageJ) to measure heart rate and pericardial dimensions while the larvae are anesthetized.</li>
		<li>To measure the heart rate, using Micromanager, create a region of interest (ROI) so as to capture the heart and exclude other structures.
		<ol>
			<li>Import the video into ImageJ as an image sequence, and ensure the correct number of frames is entered under number of images.</li>
			<li>After the file is open, use the rectangle selection tool to draw an ROI within the heart but excluding other moving elements, and save the ROI in the ROI manager (analyze | tools | ROI manager).</li>
			<li>Click plugins | install, and select the heart rate algorithm to install the code in the default folder, then select the plugin at the bottom of the plugins tab. Record the beats per minute (bpm) from the pop-up window. 
			NOTE: Image detection algorithms were custom-written to detect heart rate by measuring individual pixel density changes associated with ventricular systolic contraction. The code can be found at&#160;https://github.com/dpoburko/zFish_HR.</li>
		</ol>
		</li>
		<li>Measure pericardial dimensions using a free tool such as ImageJ to free-draw ROIs around the pericardial sac and one of the eyes. Open the image in ImageJ and use the polygon selection tool to draw an ROI first around the pericardial sac, saving it in the ROI manager as in step 5.1.2, and repeat for the eye. Select these two ROIs in the ROI manager, then click measure. Record the area for each to later calculate the area of the pericardial sac normalized to the eye area in each larva.</li>
		<li>Following heart rate and pericardial measurements, record ECG from individual larvae. 
		NOTE: Protocols for recording zebrafish ECG have been described previously33,34,35,36.</li>
	</ol>
	</li>
</ol>
6. Genotyping of CRISPR-Cas9-edited larval zebrafish
<ol>
	<li>Following phenotypic analyses, conduct on- and potential off-target genotyping to confirm accurate and precise HDR gene editing.
	<ol>
		<li>Anesthetize each 3 dpf larva in 0.3% MS-222 and tail clip to isolate gDNA using the HOTShot method37. Incubate each excised tail clip in 15 &#181;L of 25 mM NaOH at 95 &#176;C for 20 min. Then, neutralize with 1.5 &#181;L of Tris-HCl and centrifuge at 13,800 x g for 30 s. Retain the supernatant, which contains extracted gDNA.</li>
		<li>Recover the larvae in E3 media and return them to the housing system if further development or study is intended.</li>
		<li>Using the extracted gDNA as a template, perform PCR-based Sanger sequencing of on-target and potential off-target sites. 
		NOTE: Optional: a nested PCR approach may be beneficial for some gene regions.</li>
		<li>Ensure that 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 identified in step 1.2.3. 
		NOTE: Guide-design software programs often suggest primers to use, but customization may be necessary to achieve optimal results.</li>
		<li>Compile on- and off-target genotyping, heart rate, pericardial dimensions, ECG phenotyping, and reporter gene data identifiable for each zebrafish.</li>
	</ol>
	</li>
</ol>

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D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Enhanced+CRISPR/Cas9-mediated+precise+genome+editing+by+improved+design+and+delivery+of+gRNA,+Cas9+nuclease,+and+donor+DNA.">Enhanced CRISPR/Cas9-mediated precise genome editing by improved design and delivery of gRNA, Cas9 nuclease, and donor DNA.</a> Journal of Biotechnology. 241, 136-146 (2017).</li><li>Richardson, C. D., Ray, G. J., DeWitt, M. A., Curie, G. L., Corn, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Enhancing+homology-directed+genome+editing+by+catalytically+active+and+inactive+CRISPR-Cas9+using+asymmetric+donor+DNA.">Enhancing homology-directed genome editing by catalytically active and inactive CRISPR-Cas9 using asymmetric donor DNA.</a> Nature Biotechnology. 34 (3), 339-344 (2016).</li><li>Sambrook, J., Fritsch, E. F., Maniatis, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Precipitation+of+Large+RNAs+with+Lithium+Chloride.">Precipitation of Large RNAs with Lithium Chloride.</a> Molecular Cloning: A Laboratory Manual, Book 3. E.15. , (1989).</li><li>Sambrook, J., Fritsch, E. F., Maniatis, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Agarose+Gel+Electrophoresis.">Agarose Gel Electrophoresis.</a> Molecular Cloning: A Laboratory Manual, Book 1. , 3-20 (1989).</li><li>Sorlien, E. L., Witucki, M. A., Ogas, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficient+production+and+identification+of+CRISPR/Cas9-generated+gene+knockouts+in+the+model+system+Danio+rerio.">Efficient production and identification of CRISPR/Cas9-generated gene knockouts in the model system Danio rerio.</a> Journal of Visualized Experiments. (138), e56969 (2018).</li><li>Avdesh, 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=Regular+care+and+maintenance+of+a+zebrafish+(Danio+rerio)+laboratory:+An+introduction.">Regular care and maintenance of a zebrafish (Danio rerio) laboratory: An introduction.</a> Journal of Visualized Experiments. (69), e4196 (2012).</li><li>Rosen, J. N., Sweeney, M. F., Mably, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microinjection+of+zebrafish+embryos+to+analyze+gene+function.">Microinjection of zebrafish embryos to analyze gene function.</a> Journal of Visualized Experiments. (25), e1115 (2009).</li><li>Kossack, M. E., Draper, B. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+regulation+of+sex+determination+and+maintenance+in+zebrafish+(Danio+rerio).">Genetic regulation of sex determination and maintenance in zebrafish (Danio rerio).</a> Current Topics in Developmental Biology. 134, 119-149 (2019).</li><li>Tanaka, Y., 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=Functional+analysis+of+KCNH2+gene+mutations+of+type+2+long+QT+syndrome+in+larval+zebrafish+using+microscopy+and+electrocardiography.">Functional analysis of KCNH2 gene mutations of type 2 long QT syndrome in larval zebrafish using microscopy and electrocardiography.</a> Heart and Vessels. 34 (1), 159-166 (2019).</li><li>Dhillon, S. S., 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=Optimisation+of+embryonic+and+larval+ECG+measurement+in+zebrafish+for+quantifying+the+effect+of+QT+prolonging+drugs.">Optimisation of embryonic and larval ECG measurement in zebrafish for quantifying the effect of QT prolonging drugs.</a> PLoS One. 8 (4), 60552 (2013).</li><li>Yu, 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=Evolving+cardiac+conduction+phenotypes+in+developing+zebrafish+larvae:+Implications+to+drug+sensitivity.">Evolving cardiac conduction phenotypes in developing zebrafish larvae: Implications to drug sensitivity.</a> Zebrafish. 7 (4), 325-331 (2010).</li><li>Hurst, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Development and optimization of tools for embryonic electrocardiograph recording for heart dysfunction in zebrafish. , (2018).</li><li>Meeker, N. D., Hutchinson, S. A., Ho, L., Trede, N. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Method+for+isolation+of+PCR-ready+genomic+DNA+from+zebrafish+tissues.">Method for isolation of PCR-ready genomic DNA from zebrafish tissues.</a> BioTechniques. 43 (5), 610-614 (2007).</li><li>Arnaout, R., 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=Zebrafish+model+for+human+long+QT+syndrome.">Zebrafish model for human long QT syndrome.</a> Proceedings of the National Academy of Sciences of the United States of America. 104 (27), 11316-11321 (2007).</li><li>Milan, D. J., Peterson, T. A., Ruskin, J. N., Peterson, R. T., MacRae, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drugs+that+induce+repolarization+abnormalities+cause+bradycardia+in+zebrafish.">Drugs that induce repolarization abnormalities cause bradycardia in zebrafish.</a> Circulation. 107 (10), 1355-1358 (2003).</li><li>Langheinrich, U., Vacun, G., Wagner, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+embryos+express+an+orthologue+of+HERG+and+are+sensitive+toward+a+range+of+QT-prolonging+drugs+inducing+severe+arrhythmia.">Zebrafish embryos express an orthologue of HERG and are sensitive toward a range of QT-prolonging drugs inducing severe arrhythmia.</a> Toxicology and Applied Pharmacology. 193 (3), 370-382 (2003).</li><li>MacRae, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+arrhythmia:+In+vivo+screening+in+the+zebrafish+to+overcome+complexity+in+drug+discovery.">Cardiac arrhythmia: In vivo screening in the zebrafish to overcome complexity in drug discovery.</a> Expert Opinion on Drug Discovery. 5 (7), 619-632 (2010).</li></ol>

The authors have no conflicts of interest to disclose.

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">
	<tbody>
		<tr>
			<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>
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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 ...

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

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

3.3K Views.  Simon Fraser University. 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.

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

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 ...

Semi-automated Optical Heartbeat Analysis of Small Hearts

We have developed a method for analyzing high speed optical recordings from Drosophila, zebrafish and embryonic mouse hearts (Fink, et. al., 2009). Our Semi-automatic Optical Heartbeat Analysis (SOHA) uses a novel movement detection algorithm that is able to detect cardiac movements associated with individual contractile and relaxation events. The program provides a host of physiologically relevant readouts including systolic and diastolic intervals, heart rate, as well as qualitative and quantitative measures of heartbeat arrhythmicity. The program also calculates heart diameter measurements during both diastole and systole from which fractional shortening and fractional area changes are calculated. Output is provided as a digital file compatible with most spreadsheet programs. Measurements are made for every heartbeat in a record increasing the statistical power of the output. We demonstrate each of the steps where user input is required and show the application of our methodology to the analysis of heart function in all three genetically tractable heart models.

We have developed a Semi-automated Optical Heartbeat Analysis method (SOHA) for analyzing high speed optical recordings from Drosophila, zebrafish and embryonic mouse hearts. We demonstrate the application of our methodology to the analysis of heart function in fruit fly and embryonic mouse hearts.

We have developed a method for analyzing high speed optical recordings from Drosophila, zebrafish and embryonic mouse hearts (Fink, et. al., 2009). Our ...

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.

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

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Chapters in this video

Optical Mapping of Langendorff-perfused Rat Hearts

Semi-automated Optical Heartbeat Analysis of Small Hearts

In Situ Hybridization for the Precise Localization of Transcripts in Plants

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

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

Gavaging Adult Zebrafish

Cell Labeling and Injection in Developing Embryonic Mouse Hearts

Recovery of Adult Zebrafish Hearts for High-throughput Applications

Nephrotoxin Microinjection in Zebrafish to Model Acute Kidney Injury

Local Field Fluorescence Microscopy: Imaging Cellular Signals in Intact Hearts