We would like to thank Chuck Kimmel for guidance, John Dowd for help in developing this breeding strategy, Macie Walker for her work in perfecting the skeletal stain, and Charline Walker and Bonnie Ullmann for helpful zebrafish advice. This work was supported by K99/R00 DE024190 to JTN.

All experiments described in this protocol were completed in accordance and compliance with the University of Colorado and the University of Oregon Institutional Animal Care and Use Committees (IACUC).
1. Preparing the Unselected Starting Stock
<ol>
	<li>Identify heterozygous carriers of the mutant allele of interest by fin clip11 and genotyping a stock of full-sibling animals by a method of choice, such as KASP12,13,14. This protocol was performed with the zebrafish mef2cab1086 strain.</li>
	<li>KASP genotyping 
	NOTE: Allele specific primers for KASP are designed by the company that provides the reagents (see Table of Materials). To obtain the appropriate 6-carboxyl-X-rhodamine (ROX) dye concentration for the specific real-time PCR machine visit the company website.
	<ol>
		<li>Place the zebrafish tail tissue in 50 &#181;L of lysis buffer (10 mM tris (pH 8.3), 50 mM KCl, 1.5 mM MgCl2, 0.3% non-ionic surfactant, 0.3% Octylphenyl-polyethylene glycol). Add 10 &#181;L of 10 mg/mL Proteinase-K per well of a 96-well PCR plate and digest in a thermal cycler at 55 &#176;C for 2 - 5 h followed by a 20 min 94 &#176;C inactivation step. Refrigerate the lysis product after digestion.&#160; 
		NOTE: Lysis products can be successfully used for several months after preparation. When tissue was lysed using 50 mM NaOH, the KASP reactions were unsuccessful. These findings suggest that the NaOH method of genomic DNA preparation is not compatible with KASP.</li>
		<li>Quantify the genomic DNA using a spectrophotometer. Dilute the genomic DNA template using molecular biology grade water so that each reaction contains approximately 20-30 ng. 
		NOTE: Quantify 4 or 5 samples, pool them, then prepare dilutions for all samples based on the average. When using this protocol for adult tail tissue, a 1:100 dilution is often sufficient. The precise quantification of DNA concentration is not required.</li>
		<li>Add 0.14 &#181;L of the KASP primer mix and 5 &#181;L of the KASP master mix per sample on ice into a 1.5 mL microcentrifuge tube and mix well before briefly centrifuging to collect contents at the bottom of the tube. 
		NOTE: KASP master mix is heat and light sensitive, keep stock on ice and avoid direct light.</li>
		<li>Pipet 5 &#181;L of primer/master mix solution into the wells of a 96-well optical PCR plate suitable for the real-time PCR machine being used.</li>
		<li>Pipet 5 &#181;L of diluted (approximately 5 - 7 ng/&#181;L) DNA template samples into each well and aspirate with a pipette to mix.</li>
		<li>Add 5 &#181;L of nuclease-free water to 3 wells to serve as no-template controls (NTC) and add 5 &#181;L of diluted confirmed homozygous wild type, heterozygous and homozygous mutant DNA template for positive controls.</li>
		<li>Place an optically clear film seal over the plate ensuring that the film is fully sealed on each well.</li>
		<li>Briefly centrifuge the plate at 600 x g to collect the contents at the bottom and to remove bubbles from the mix. 
		NOTE: Keep the plate on ice and shield from light until ready to proceed.</li>
		<li>Use the cycling program shown in Table 1 to perform the main KASP reaction.</li>
		<li>View the resulting scatter plot produced by the analysis computer. A successful reaction will show 4 distinct groupings of points on the plot (Figure 3): three sample groupings corresponding to genotype, and one grouping of the NTCs near the origin. The positive controls should segregate with the appropriate sample grouping.</li>
		<li>If the computer program does not recognize the plot groupings as corresponding to specific genotypes, additional cycling sets may be required (Table 2). Repeat the additional program until the computer recognizes tight groupings by genotype.</li>
		<li>Export the genotyping results to a spreadsheet file for easier use and analysis.</li>
	</ol>
	</li>
	<li>House the identified heterozygous animals together on the main water system.</li>
</ol>
2. The First Round of Progeny Scoring
<ol>
	<li>Pairwise cross sibling heterozygotes
	<ol>
		<li>After the animals recover from the fin clip, set up pairwise intercrosses. Use dividers to ensure embryos are synchronous16.</li>
		<li>Collect the embryos and keep the parental pairs isolated in breeding cages.</li>
		<li>Monitor isolated adults so that aggressive fish can be separated or provided with cover (shredded fish nets work nicely). Fish should be fed and their water changed following IACUC guidelines while in static water.</li>
	</ol>
	</li>
	<li>Embryo rearing
	<ol>
		<li>Stage and raise the zebrafish embryos to larvae under standard conditions17.</li>
		<li>Collect phenotypically wild-type animals using a glass pipette on day 5 or 6 when the swim bladder is inflated in 75% of the clutch. Place phenotypic wild types in the nursery recording of which parents yielded them. 
		Note: With mutant alleles that are recessive lethal, which is true of many skeletal mutants, homozygous mutants fail to form swim bladders18,19. Therefore, phenotypic wild types can be easily discerned from their mutant siblings based on inflated swim bladders.</li>
	</ol>
	</li>
	<li>Alcian Blue/Alizarin Red cartilage and bone staining
	<ol>
		<li>Anesthetize larvae that do not inflate their swim bladders using 0.17 g/L Tricaine-S in embryo media. Collect animals into 1.5 mL microcentrifuge tubes with a wide-bore glass Pasteur pipet. Add no more than 100 larvae per tube.</li>
		<li>Remove embryo water from each tube and fix animals in 1 mL of 2% paraformaldehyde in 1x PBS per tube. 
		CAUTION: Aqueous PFA is combustible and will cause serious skin and eye irritation. Wear appropriate personal protective equipment such as gloves and eye protection, and wash hands thoroughly after use.</li>
		<li>Rock for 1 h; longer fixation impairs bone staining. Check the tubes regularly during this and all subsequent rocking steps to ensure no larvae have become stuck to the side or clumped in the bottom of the tube.</li>
		<li>Remove the fixative from tubes using a glass pipette, add 1 mL of 50% ethanol, and rock for 10 min.</li>
		<li>Prepare fresh staining solution by adding 20 &#181;L of 0.5% Alizarin Red per 1 mL of Alcian premix (0.04% Alcian Blue/10 mM MgCl2/80% ethanol). Remove 50% ethanol, add 1 mL of staining solution to each tube, and rock overnight. Larvae can remain in the stain for up to 5 days.</li>
		<li>Remove the stain solution from the tubes and add 1 mL of 80% ethanol/10 mM MgCl2 solution to each tube. Rock the tubes for at least 1 h; rinsing overnight can produce a clearer Alcian Blue stain.</li>
		<li>Remove the 80% ethanol/10 mM MgCl2 solution from the tubes and add 1 mL of 50% ethanol; rock for 5 min.</li>
		<li>Remove the 50% ethanol solution from the tubes and add 1 mL of 25% ethanol and rock for 5 min.</li>
		<li>Remove the 50% ethanol solution from the tubes and add 1 mL freshly prepared 3% H2O2/0.5% KOH bleaching solution. Leave the caps open and let the tubes stand in in a micro-tube rack for approximately 10 min or until a faint brown coloration can be seen in the upper part of the solution. 
		NOTE: A layer of bubbles form on top of the solution. Careful timing is required at this step, as over bleaching will result in a poor double stain.</li>
		<li>Remove the bleaching solution and add 1 mL of 25% glycerol/0.1% KOH; rock for 10 min to 1 h.</li>
		<li>Remove 25% glycerol/0.1% KOH solution from the tubes, add 1 mL of 50% glycerol/0.1% KOH solution to each tube, rock for 10 min to overnight.</li>
		<li>Remove 50% glycerol/0.1% KOH solution from the tubes and again add 50% glycerol/0.1% KOH solution to each tube. Rocking overnight will help remove air bubbles that can accumulate inside larvae. Store stained skeletal preparations at 4 &#176;C when not being used.</li>
	</ol>
	</li>
	<li>Phenotype scoring
	<ol>
		<li>Score stained mutant offspring for the penetrance of the selected phenotype. In Nichols et al.1 mutants were scored for any occurrence of ectopic bone. 
		Note: Because the zebrafish parents are in static water and the Alizarin Red stain can fade with time, it is important to score skeletons within a few days of completing the skeletal preparation.</li>
		<li>Penetrance is the proportion of a genotype that has a phenotype. Calculate the percent penetrance by the following formula: 
		% penetrance = (mutants with phenotype)/(total number of mutants) &#215; 100</li>
	</ol>
	</li>
</ol>
3. Family Selection
<ol>
	<li>Choose two high penetrance and two low penetrance families for the next generation. In addition to penetrance, it is important to consider fecundity and vigor when choosing which families to propagate; see Discussion for more details on this.</li>
	<li>Give each parental pair a sub stock identifier: family .01, .02, etc.</li>
	<li>House parental pairs on the main system with several mixed sex &#39;companion&#39; fish. Companion fish can be any zebrafish line with permanent, obvious distinguishing features, such as altered pigmentation or fin structure. 
	NOTE: While many researchers successfully keep only breeding pairs in a tank, favorable results are seen by housing pairs with several companion fish. This optional step allows breeding pairs of animals to be kept in larger schools and easily retrieved so they can be repeatedly crossed as needed. Parental pairs can be crossed repeatedly to generate large full-sibling families; wait at least one week before repeat crossing the same parental pair.</li>
	<li>Cull the larval families from step 2.2.2. that were not selected for the next generation.</li>
	<li>Label the tanks containing the wild-type sibling larvae from selected families with the penetrance from their mutant siblings and raise to adulthood as normal.</li>
	<li>The next generation will have at least four full-sibling families, two for the upward line and two for the downward line.</li>
	<li>When larvae reach sexual maturity, repeat the protocol starting at step one with the new generation.</li>
</ol>

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G., 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=An+essential+role+of+variant+histone+H3.3+for+ectomesenchyme+potential+of+the+cranial+neural+crest.">An essential role of variant histone H3.3 for ectomesenchyme potential of the cranial neural crest.</a> PLoS Genet. 8 (9), e1002938 (2012).</li><li>DeLaurier, 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=Role+of+mef2ca+in+developmental+buffering+of+the+zebrafish+larval+hyoid+dermal+skeleton.">Role of mef2ca in developmental buffering of the zebrafish larval hyoid dermal skeleton.</a> Dev Biol. 385 (2), 189-199 (2014).</li><li>McGurk, P. D., Ben Lovely, C., Eberhart, J. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analyzing+Craniofacial+Morphogenesis+in+Zebrafish+Using+4D+Confocal+Microscopy.">Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy.</a> J Vis Exp. (83), e51190 (2014).</li><li>Kemp, H. A., Carmany-Rampey, A., Moens, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generating+chimeric+zebrafish+embryos+by+transplantation.">Generating chimeric zebrafish embryos by transplantation.</a> J Vis Exp. (29), e1394 (2009).</li><li>Lush, J. 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The authors have nothing to disclose.

Selective Breeding Unveils Subtleties of Gene Function
Shifting mutant phenotypes to be either more or less severe by selective breeding is a straightforward way to gain new insights into gene function. When compared with standard methods of unselected breeding, the protocol presented here can yield a much more complete understanding of mutant phenotypes. Specifically, by generating strains that are severe, the full breadth of mutant phenotypes may be revealed, including some ...

The zebrafish is a powerful model system for understanding skeletal development. With mutant zebrafish strains, biologists can decipher gene function during skeletogenesis. However, zebrafish skeletal mutant phenotypes can present with variable penetrance1,2,3,4 which can hinder developmental and genetic analyses. The purpose of this method is threefold. First, generating zebrafish mutant lines which consistently produce severe phenotypes enables downstream developmental studies like time-lapse recording5 and transplantation6. These sorts of studies can be crippled by attempting to study phenotypes that only manifest inconsistently. Second, inbreeding zebrafish strains can decrease genetic background variation, thus promoting experimental consistency and reproducibility. For example, performing all in situ hybridization analyses on one selectively inbred strain can reduce confounding variability and strengthen conclusions. Third, generating severe and mild strains will reveal the entire phenotypic series that can result from a particular mutation.
At first glance, selective breeding of lethal mutants seems impossible. How can one breed for penetrance when the animals that are scored for selection are dead? Fortunately, methods for selective breeding by family selection, specifically progeny testing, have demonstrated effectiveness in livestock breeding programs for many years7,8. These programs are mainly used for selective breeding for traits that are only present in one sex, like milk production in cows or egg production in hens. The males of these species cannot be scored directly, but their progeny are scored and a value is then assigned to the parents. Borrowing from this strategy, the protocol presented here involves scoring the fixed and stained mutant offspring from a pair of zebrafish that are heterozygous for a mutant gene of interest. The penetrance of a phenotype in the homozygous lethal mutant offspring is assigned to the parents when deciding which individuals will produce the next generation in the line. We find that this method is an effective means of shifting penetrance in zebrafish lethal skeletal mutants1.
Similar to other studies, this selective breeding protocol takes under consideration criteria like clutch size, survival of offspring, normal development of embryos, and sex ratio9. However, these factors are all considered in the context of a mutant background with the objective of shifting the mutant penetrance. Therefore, this protocol extends previous selective breeding paradigms by offering a method to strengthen developmental mutant analyses as well as increase background homogeneity.
This protocol requires extensive genotyping, so it is important to develop a reliable, rapid genotyping protocol in advance. There are many genotyping protocols available10,11, however we find the KASP genotyping12,13,14 is faster, more cost efficient, and more reliable than methods based on restriction enzyme cleavage of amplified sequences10. Therefore, we include a KASP protocol in this work. Additionally, we focus on skeletal mutant phenotypes in this protocol and include a procedure for Alcian Blue/Alizarin Red staining modified from previous protocols15.
The method described here is a straightforward strategy for shifting lethal mutant penetrance upward or downward. While this protocol focuses on skeletal mutant phenotypes, we believe it will be a useful strategy for husbandry of all mutant zebrafish lines. In fact, the utility of this breeding strategy likely extends beyond zebrafish. We predict that this protocol can be modified to shift penetrance in a broad range of organisms. Shifting lethal penetrance by progeny testing can help push forward the progress of any developmental geneticist.

<table border="0" cellpadding="0" cellspacing="0" width="627">
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Paraformaldehyde, pelleted, solid</td>
			<td style="width:125px;">Ted Pella Co.</td>
			<td style="width:119px;">18501</td>
			<td style="width:167px;">Pelleted PFA is a safer alternative to powdered PFA</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Magnesium Chloride, solid</td>
			<td style="width:125px;">Acros Organics</td>
			<td>223210010</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">10x PBS, Aqueous</td>
			<td style="width:125px;">Fisher</td>
			<td style="width:119px;">BP3994</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">190 proof Ethanol</td>
			<td style="width:125px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Alcian Blue, solid</td>
			<td style="width:125px;">Anatech Ltd.</td>
			<td style="width:119px;">867</td>
			<td>Must be from Anatech</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Alizarin Red, solid</td>
			<td style="width:125px;">Sigma</td>
			<td style="width:119px;">A5533-25G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Glycerol, liquid</td>
			<td style="width:125px;">Fisher</td>
			<td style="width:119px;">BP229 1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Hydrogen peroxide, liquid</td>
			<td style="width:125px;">Fisher</td>
			<td style="width:119px;">BP263500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Potassium hydroxide,&#160; solid</td>
			<td style="width:125px;">Fisher</td>
			<td style="width:119px;">P250 500</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">StepOnePlus Real-time PCR Machine</td>
			<td style="width:125px;">Applied Biosystems</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:216px;">MicroAmp Fast Optical 96-well Reaction Plate with Barcode (0.1mL)</td>
			<td style="width:125px;">Applied Biosystems</td>
			<td style="width:119px;">4346906</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Microseal &#39;B&#39; seal</td>
			<td style="width:125px;">BioRad</td>
			<td style="width:119px;">MSB1001</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">KASP Master Mix, High ROX</td>
			<td style="width:125px;">LGC</td>
			<td>KBS-1016-022</td>
			<td>https://www.lgcgroup.com/products/kasp-genotyping-chemistry/#.WOPX41UrKUk</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">KASP By Design Primer Mix</td>
			<td style="width:125px;">LGC</td>
			<td style="width:119px;">KBS-2100-100</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Tris HCl, solid</td>
			<td style="width:125px;">Fisher</td>
			<td style="width:119px;">BP153 500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">potassium chloride, solid</td>
			<td style="width:125px;">Fisher</td>
			<td style="width:119px;">BP366 500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Tween-20, liquid</td>
			<td style="width:125px;">Fisher</td>
			<td style="width:119px;">BP337 100</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Nonidet P40</td>
			<td style="width:125px;">ThermoFisher</td>
			<td>28324</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Tricaine-S</td>
			<td style="width:125px;">Western Chemicals</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Proteinase K</td>
			<td style="width:125px;">Fisher</td>
			<td style="width:119px;">BP1700 100</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">T100 Thermal Cycler</td>
			<td style="width:125px;">BioRad</td>
			<td>1861096</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Controlled Drop Pasteur Pipets</td>
			<td style="width:125px;">Fisher</td>
			<td>13-678-30</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Nanodrop</td>
			<td style="width:125px;">ThermoFisher</td>
			<td style="width:119px;"></td>
			<td>for DNA quantitation</td>
		</tr>
	</tbody>
</table>

shifting zebrafish lethal skeletal mutant penetrance by progeny testing

Zebrafish mutant phenotypes are often incompletely penetrant, only manifesting in some mutants. Interesting phenotypes that inconsistently appear can be difficult to study, and can lead to confounding results. The protocol described here is a straightforward breeding paradigm to increase and decrease penetrance in lethal zebrafish skeletal mutants. Because lethal mutants cannot be selectively bred directly, the classic selective breeding strategy of progeny testing is employed. This method also includes protocols for Kompetitive Allele Specific PCR (KASP) genotyping zebrafish and staining larval zebrafish cartilage and bone. Applying the husbandry strategy described here can increase the penetrance of an interesting skeletal phenotype enabling more reproducible results in downstream applications. In addition, decreasing the mutant penetrance through this selective breeding strategy can reveal the developmental processes that most crucially require the function of the mutated gene. While the skeleton is specifically considered here, we propose that this methodology will be useful for all zebrafish mutant lines.

This protocol is a long-term husbandry technique useful for understanding zebrafish skeletal mutants (Figure 1). Selective breeding by progeny testing should yield a shift in overall penetrance both downward and upward in a few generations (Figure 2). In our previous work, two rounds of selective breeding drove the average penetrance downward from 17% to 3%1. Similarly, in our upward line, we shifted the a...

Watch this Scientific Journal Video about Shifting Zebrafish Lethal Skeletal Mutant Penetrance by Progeny Testing at JoVE.com

Shifting Zebrafish Lethal Skeletal Mutant Penetrance by Progeny Testing

The goal of this protocol is to alter the penetrance of lethal skeletal mutant phenotypes in zebrafish by selective breeding. Lethal mutants cannot be grown to adulthood and bred themselves, therefore this protocol describes a method for tracking and selecting penetrance through multiple generations by progeny testing.

Zebrafish mutant phenotypes are often incompletely penetrant, only manifesting in some mutants. Interesting phenotypes that inconsistently appear can be ...

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7.5K Views.  University of Colorado Anschutz Medical Campus. The goal of this protocol is to alter the penetrance of lethal skeletal mutant phenotypes in zebrafish by selective breeding. Lethal mutants cannot be grown to adulthood and bred themselves, therefore this protocol describes a method for tracking and selecting penetrance through multiple generations by progeny testing. 

Video: Shifting Zebrafish Lethal Skeletal Mutant Penetrance by Progeny Testing

Kidney Regeneration in Adult Zebrafish by Gentamicin Induced Injury

The kidney is essential for fluid homeostasis, blood pressure regulation and filtration of waste from the body. The fundamental unit of kidney function is the nephron. Mammals are able to repair existing nephrons after injury, but lose the ability to form new nephrons soon after birth. In contrast to mammals, adult fish produce new nephrons (neonephrogenesis) throughout their lives in response to growth requirements or injury. Recently, lhx1a has been shown to mark nephron progenitor cells in the adult zebrafish kidney, however mechanisms controlling the formation of new nephrons after injury remain unknown. Here we show our method for robust and reproducible injury in the adult zebrafish kidney by intraperitoneal (i.p.) injection of gentamicin, which uses a noninvasive visual screening process to select for fish with strong but nonlethal injury. Using this method, we can determine optimal gentamicin dosages for injury and go on to demonstrate the effect of higher temperatures on kidney regeneration in zebrafish.

Here we present a reliable method to study adult kidney regeneration by inducing acute kidney injury by gentamicin injection. We show that injury is dependent on gentamicin dosage and environmental temperature using in situ hybridization to label lhx1a+ developing new nephrons.

The kidney is essential for fluid homeostasis, blood pressure regulation and filtration of waste from the body. The fundamental unit of kidney function is ...

Mechanical Vessel Injury in Zebrafish Embryos

Zebrafish (Danio rerio) embryos have proven to be a powerful model for studying a variety of developmental and disease processes. External development and optical transparency make these embryos especially amenable to microscopy, and numerous transgenic lines that label specific cell types with fluorescent proteins are available, making the zebrafish embryo an ideal system for visualizing the interaction of vascular, hematopoietic, and other cell types during injury and repair in vivo. Forward and reverse genetics in zebrafish are well developed, and pharmacological manipulation is possible. We describe a mechanical vascular injury model using micromanipulation techniques that exploits several of these features to study responses to vascular injury including hemostasis and blood vessel repair. Using a combination of video and timelapse microscopy, we demonstrate that this method of vascular injury results in measurable and reproducible responses during hemostasis and wound repair. This method provides a system for studying vascular injury and repair in detail in a whole animal model.

This article describes a method for creating a mechanical vessel injury in zebrafish embryos. This injury model provides a platform for studying hemostasis, injury-related inflammation, and wound healing in an organism ideally suited for real-time microscopy.

Zebrafish (Danio rerio) embryos have proven to be a powerful model for studying a variety of developmental and disease processes.  External development ...

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

Generation of Parabiotic Zebrafish Embryos by Surgical Fusion of Developing Blastulae

Surgical parabiosis of two animals of different genetic backgrounds creates a unique scenario to study cell-intrinsic versus cell-extrinsic roles for candidate genes of interest, migratory behaviors of cells, and secreted signals in distinct genetic settings. Because parabiotic animals share a common circulation, any blood or blood-borne factor from one animal will be exchanged with its partner and vice versa. Thus, cells and molecular factors derived from one genetic background can be studied in the context of a second genetic background. Parabiosis of adult mice has been &#160;used extensively to research aging, cancer, diabetes, obesity, and brain development. More recently, parabiosis of zebrafish embryos has been used to study the developmental biology of hematopoiesis. In contrast to mice, the transparent nature of zebrafish embryos permits the direct visualization of cells in the parabiotic context, making it a uniquely powerful method for investigating fundamental cellular and molecular mechanisms. The utility of this technique, however, is limited by a steep learning curve for generating the parabiotic zebrafish embryos. This protocol provides a step-by-step method on how to surgically fuse the blastulae of two zebrafish embryos of different genetic backgrounds to investigate the role of candidate genes of interest. In addition, the parabiotic zebrafish embryos are tolerant to heat shock, making temporal control of gene expression possible. This method does not require a sophisticated set-up and has broad applications for studying cell migration, fate specification, and differentiation in vivo during embryonic development.

This protocol provides step-by-step instruction on how to generate parabiotic zebrafish embryos of different genetic backgrounds. When combined with the unparalleled imaging capabilities of the zebrafish embryo, this method provides a uniquely powerful means to investigate cell-autonomous versus non-cell-autonomous functions for candidate genes of interest.

Surgical parabiosis of two animals of different genetic backgrounds creates a unique scenario to study cell-intrinsic versus cell-extrinsic roles for ...

Induction of Acute Skeletal Muscle Regeneration by Cardiotoxin Injection

Skeletal muscle regeneration is a physiological process that occurs in adult skeletal muscles in response to injury or disease. Acute injury-induced skeletal muscle regeneration is a widely used, powerful model system to study the events involved in muscle regeneration as well as the mechanisms and different players. Indeed, a detailed knowledge of this process is essential for a better understanding of the pathological conditions that lead to skeletal muscle degeneration, and it aids in identifying new targeted therapeutic strategies. The present work describes a detailed and reproducible protocol to induce acute skeletal muscle regeneration in mice through a single intramuscular injection of cardiotoxin (CTX). CTX belongs to the family of snake venom toxins and causes myolysis of myofibers, which eventually triggers the regeneration events. The dynamics of skeletal muscle regeneration is evaluated by histological analysis of muscle sections. The protocol also illustrates the experimental procedures for dissecting, freezing, and cutting the Tibialis Anterior muscle, as well as the routine Hematoxylin &#38; Eosin staining that is widely used for subsequent morphological and morphometric analysis.

This manuscript describes a detailed protocol to induce acute skeletal muscle regeneration in adult mice and subsequent manipulations of the muscles, such as dissection, freezing, cutting, routine staining, and myofiber cross-sectional area analysis.

Skeletal muscle regeneration is a physiological process that occurs in adult skeletal muscles in response to injury or disease. Acute injury-induced ...

Identification Of Erythromyeloid Progenitors And Their Progeny In The Mouse Embryo By Flow Cytometry

Macrophages are professional phagocytes from the innate arm of the immune system. In steady-state, sessile macrophages are found in adult tissues where they act as front line sentinels of infection and tissue damage. While other immune cells are continuously renewed from hematopoietic stem and progenitor cells (HSPC) located in the bone marrow, a lineage of macrophages, known as resident macrophages, have been shown to be self-maintained in tissues without input from bone marrow HSPCs. This lineage is exemplified by microglia in the brain, Kupffer cells in the liver and Langerhans cells in the epidermis among others. The intestinal and colon lamina propria are the only adult tissues devoid of HSPC-independent resident macrophages. Recent investigations have identified that resident macrophages originate from the extra-embryonic yolk sac hematopoiesis from progenitor(s) distinct from fetal hematopoietic stem cells (HSC). Among yolk sac definitive hematopoiesis, erythromyeloid progenitors (EMP) give rise both to erythroid and myeloid cells, in particular resident macrophages. EMP are only generated within the yolk sac between E8.5 and E10.5 days of development and they migrate to the fetal liver as early as circulation is connected, where they expand and differentiate until at least E16.5. Their progeny includes erythrocytes, macrophages, neutrophils and mast cells but only EMP-derived macrophages persist until adulthood in tissues. The transient nature of EMP emergence and the temporal overlap with HSC generation renders the analysis of these progenitors difficult. We have established a tamoxifen-inducible fate mapping protocol based on expression of the macrophage cytokine receptor Csf1r promoter to characterize EMP and EMP-derived cells in vivo by flow cytometry.

While infiltrating macrophages are continuously recruited to adult tissues from circulating precursors, resident macrophages seed their tissue during development, where they are maintained without further input from progenitors. The progenitors for resident macrophages were recently identified. Here, we present methods for the genetic fate mapping of the resident macrophage progenitors.

Macrophages are professional phagocytes from the innate arm of the immune system. In steady-state, sessile macrophages are found in adult tissues where ...

Culture of Adult Transgenic Zebrafish Retinal Explants for Live-cell Imaging by Multiphoton Microscopy

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The M&#252;ller glia re-enter the cell cycle and produce neuronal progenitor cells that undergo subsequent rounds of cell divisions and differentiate into the lost neuronal cell types. Both M&#252;ller glia and neuronal progenitor cell nuclei replicate their DNA and undergo mitosis in distinct locations of the retina, i.e. they migrate between the basal Inner Nuclear Layer (INL) and the Outer Nuclear Layer (ONL), respectively, in a process described as Interkinetic Nuclear Migration (INM). INM has predominantly been studied in the developing retina. To examine the dynamics of INM in the adult regenerating zebrafish retina in detail, live-cell imaging of fluorescently-labeled M&#252;ller glia/neuronal progenitor cells is required. Here, we provide the conditions to isolate and culture dorsal retinas from Tg[gfap:nGFP]mi2004 zebrafish that were exposed to constant intense light for 35 h. We also show that these retinal cultures are viable to perform live-cell imaging experiments, continuously acquiring z-stack images throughout the thickness of the retinal explant for up to 8 h using multiphoton microscopy to monitor the migratory behavior of gfap:nGFP-positive cells. In addition, we describe the details to perform post-imaging analysis to determine the velocity of apical and basal INM. To summarize, we established conditions to study the dynamics of INM in an adult model of neuronal regeneration. This will advance our understanding of this crucial cellular process and allow us to determine the mechanisms that control INM.

Zebrafish retinal regeneration has mostly been studied using fixed retinas. However, dynamic processes such as interkinetic nuclear migration occur during the regenerative response and require live-cell imaging to investigate the underlying mechanisms. Here, we describe culture and imaging conditions to monitor Interkinetic Nuclear Migration (INM) in real-time using multiphoton microscopy.

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The ...

Isolation of Type I and Type II Pericytes from Mouse Skeletal Muscles

Pericytes are perivascular multipotent cells that show heterogeneity in different organs or even within the same tissue. In skeletal muscles, there are at least two pericyte subpopulations (called type I and type II), which express different molecular markers and have distinct differentiation capabilities. Using NG2-DsRed and Nestin-GFP double-transgenic mice, type I (NG2-DsRed+Nestin-GFP-) and type II (NG2-DsRed+Nestin-GFP+) pericytes have been successfully isolated. However, the availability of these double-transgenic mice prevents the widespread use of this purification method. This work describes an alternative protocol that allows for the easy and simultaneous isolation of type I and type II pericytes from skeletal muscles. This protocol utilizes the fluorescence-activated cell sorting (FACS) technique and targets PDGFR&#946;, rather than NG2, together with the Nestin-GFP signal. Following isolation, type I and type II pericytes show distinct morphologies. In addition, type I and type II pericytes isolated with this new method, like those isolated from the double-transgenic mice, are adipogenic and myogenic, respectively. These results suggest that this protocol can be used to isolate pericyte subpopulations from skeletal muscles and possibly from other tissues.

This work describes a FACS-based protocol that allows for easy and simultaneous isolation of type I and type II pericytes from skeletal muscles.

Pericytes are perivascular multipotent cells that show heterogeneity in different organs or even within the same tissue. In skeletal muscles, there are at ...

Identification of Skeletal Muscle Satellite Cells by Immunofluorescence with Pax7 and Laminin Antibodies

Immunofluorescence is an effective method that helps to identify different cell types on tissue sections. In order to study the desired cell population, antibodies for specific cell markers are applied on tissue sections. In adult skeletal muscle, satellite cells (SCs) are stem cells that contribute to muscle repair and regeneration. Therefore, it is important to visualize and trace the satellite cell population under different physiological conditions. In resting skeletal muscle, SCs reside between the basal lamina and myofiber plasma membrane. A commonly used marker for identifying SCs on the myofibers or in cell culture is the paired box protein Pax7. In this article, an optimized Pax7 immunofluorescence protocol on skeletal muscle sections is presented that minimizes non-specific staining and background. Another antibody that recognizes a protein (laminin) of the basal lamina was also added to help identify SCs. Similar protocols can also be used to perform double or triple labeling with Pax7 and antibodies for additional proteins of interest.

The precise identification of satellite cells is essential for studying their functions under various physiological and pathological conditions. This article presents a protocol to identify satellite cells on adult skeletal muscle sections by immunofluorescence-based staining.

Immunofluorescence is an effective method that helps to identify different cell types on tissue sections. In order to study the desired cell population, ...

Performing Human Skeletal Muscle Xenografts in Immunodeficient Mice

Treatment effects observed in animal studies often fail to be recapitulated in clinical trials. While this problem is multifaceted, one reason for this failure is the use of inadequate laboratory models. It is challenging to model complex human diseases in traditional laboratory organisms, but this issue can be circumvented through the study of human xenografts. The surgical method we describe here allows for the creation of human skeletal muscle xenografts, which can be used to model muscle disease and to carry out preclinical therapeutic testing. Under an Institutional Review Board (IRB)-approved protocol, skeletal muscle specimens are acquired from patients and then transplanted into NOD-Rag1null IL2r&gamma;null (NRG) host mice. These mice are ideal hosts for transplantation studies due to their inability to make mature lymphocytes and are thus unable to develop cell-mediated and humoral adaptive immune responses. Host mice are anaesthetized with isoflurane, and the mouse tibialis anterior and extensor digitorum longus muscles are removed. A piece of human muscle is then placed in the empty tibial compartment and sutured to the proximal and distal tendons of the peroneus longus muscle. The xenografted muscle is spontaneously vascularized and innervated by the mouse host, resulting in robustly regenerated human muscle that can serve as a model for preclinical studies.

Complex human diseases can be challenging to model in traditional laboratory model systems. Here, we describe a surgical approach to model human muscle disease through the transplantation of human skeletal muscle biopsies into immunodeficient mice.

Treatment effects observed in animal studies often fail to be recapitulated in clinical trials. While this problem is multifaceted, one reason for this ...