Name: examining muscle regeneration in zebrafish models of muscle disease
Uploaded: 2021-01-18T15:00:00.000+00:00
Duration: 7 min 58 s
Description: Watch this Scientific Journal Video about Examining Muscle Regeneration in Zebrafish Models of Muscle Disease at JoVE.com

We would like to thank Dr. Alex Fulcher, and Monash Micro Imaging for assistance with microscope maintenance and setup. The Australian Regenerative Medicine Institute is supported by grants from the State Government of Victoria and the Australian Government. This work was funded by a Muscular Dystrophy Association (USA) project grant to P.D.C (628882).

Zebrafish maintenance was carried out as per the standard operating procedures approved by the Monash University Animal Ethics Committee under breeding colony license ERM14481.
1. Determination of the genotype of live embryos using an embryo genotyping platform.
<ol>
	<li>Anesthetize 3 days post fertilization (dpf) zebrafish embryos by adding tricaine methanesulfonate to a final concentration of 0.016% (v/v) in embryo medium (5 mM NaCl, 0.17mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4 in water). Wait for 10 minutes to ensure that the fish are completely anesthetized, evident when the fish stop swimming.</li>
	<li>Prepare the 24 chamber DNA extraction chip by peeling the clear protective film from the top surface of the chip (Figure 1A).</li>
	<li>Cut the tip of a 20 &#181;L filter tip to widen the opening. Using this, pick a single embryo in a 13 &#181;L volume of embryo medium and load into the first chamber of the chip. Repeat this process until embryos have been dispensed into each chamber. 
	NOTE: This step should be performed in an area with minimal air drafts, as high air flow may cause excessive evaporation of the fluid subsequently resulting in reduced genotyping sensitivity and survival of the embryos.</li>
	<li>Once the desired number of embryos have been loaded onto the DNA extraction chip, load it onto the zebrafish embryo genotyping platform. This is best achieved by placing in one side first, followed by the rest of it. 
	NOTE: Avoid adding too much pressure onto the platform as this may overstress and damage the underlying springs.</li>
	<li>Affix the magnetic platform lid over the chip, which will prevent evaporation of embryo media during the DNA extraction protocol, and close the platform lid.</li>
	<li>Set the base unit to 2.4 volts, 0.051 A, and 0.12 W and start the DNA extraction protocol by pressing the &#34;ON/OFF&#34; button. The platform should start vibrating, which can be assessed by gently touching the lid. The protocol should be run for 8 minutes. 
	NOTE: The vigorous vibration results in shedding of epidermal cells, from which genomic DNA is extracted.</li>
	<li>While the program is running, prepare a 24 well plate by adding 1 mL of embryo medium to each well, which is necessary to separate individual embryos while downstream genotyping assays are being performed.</li>
	<li>Additionally, appropriately label 8 well strip tubes, which will be used for collection of DNA material from each embryo.</li>
	<li>At the completion of the 8 minute protocol, press the ON/OFF button to stop the vibration of the platform.</li>
	<li>Open the lid of the platform, and gently remove the magnetic lid ensuring minimal pressure is applied to the platform.</li>
	<li>Carefully remove the DNA extraction chip from the platform and place it on a flat surface.</li>
	<li>From the first chamber, remove 10 &#181;L of embryo media surrounding the embryo and place it into the appropriate well of the 8-well strip tube. The media contains the genetic material from that embryo, which will be used for downstream assays. 
	NOTE: Avoid touching the embryo with the pipette tip during media collection, as this can damage the embryo.</li>
	<li>Using a plastic pipette, immediately add two drops of fresh embryo medium to the same chamber. Collect the embryo and move it to the first well of a 24 well plate.</li>
	<li>Repeat steps 1.12 and 1.13 for all the remaining chambers. At the completion of this, the 24 well plate containing live embryos can be moved to a 28 &#176;C incubator.</li>
	<li>Perform appropriate downstream genotyping assays to determine the genotype of the embryos. 
	NOTE: The DNA obtained from the embryo genotyping platform can be successfully amplified by PCR and can be used for subsequent analysis including sequencing, gel electrophoresis, or high-resolution melt-analysis14. To genotype the lama2 strain described in the representative results, PCR, restriction digestion and gel electrophoresis were used. Given that the concentration of DNA obtained from each embryo is low, it is suggested to utilise most, if not all, of the genetic material obtained for the downstream assay.</li>
	<li>Once the genotype of each larvae has been identified, transfer them to 90 mm Petri dishes, placing each genotype in a different dish. Fill the dish with 25 mL embryo medium and keep them in the 28 &#176;C incubator.</li>
</ol>
2. Performing muscle injury using a needle stab
<ol>
	<li>Once the larvae are 4 dpf, anesthetize them by adding tricaine methanesulfonate to a final concentration of 0.016% (v/v) in embryo medium. Wait for 10 minutes to ensure that the fish are completely anesthetized, evident when the fish stop swimming. 
	NOTE: To avoid bias, the identity of the genotype should be blinded to the investigator performing the stabs and downstream analyses.</li>
	<li>During this time, prepare 24 well plates by filling each well with 1 mL of fresh embryo medium, and set up the stereomicroscope with a black background and high-power light, to facilitate the visualization of the somite borders.</li>
	<li>Using a plastic pipette, transfer one anesthetized larva into a new Petri dish.</li>
	<li>Carefully remove excess embryo medium with a pipette, and under a dissecting microscope, orient the fish such that the head is on the left, tail on the right, dorsal region up and ventral region down (Figure 1B).</li>
	<li>Working under a dissecting microscope, use a 30-gauge needle to perform a quick but precise stab in the epaxial muscle located above the horizontal myoseptum. To ensure consistency in anterior-posterior position, aim for 1-2 somites located above the anal pore (Figure 1B). 
	NOTE: Avoid stabbing the neural tube and notochord, and work quickly to avoid the drying of the fish during the process.</li>
	<li>Using a plastic pipette, pour a drop of embryo medium on the stabbed zebrafish and carefully transfer it into a well of the 24 well plate.</li>
	<li>Repeat steps 2.3 to 2.6. until all the fish have been stabbed. 
	&#8203;NOTE: Several larvae can be stabbed together depending on the processing speed and proficiency of the investigator, as long as the fish do not dry out during the process.</li>
	<li>Once all the larvae have been stabbed, place the 24 well plate in the 28 &#176;C incubator until subsequent imaging is performed.</li>
</ol>
3. Imaging of muscle injury and recovery
<ol>
	<li>At 1 day post injury (1 dpi), anaesthetize the stabbed larvae by adding tricaine methanesulfonate to a final concentration of 0.016% (v/v) in embryo medium. Wait for 10 minutes to ensure that the fish are completely anesthetized, evident when the fish stop swimming. 
	NOTE: At 0 dpi, the wound site contains a large amount of cellular debris, which makes it difficult to quantify the extent of muscle injury (Supplementary Figure 1A-B). It takes approximately 18-20 hours for this debris to be cleared from the wound site, and as such it is more reliable to image larvae at 1 dpi, rather than 0 dpi, to determine the extent of injury elicited.</li>
	<li>Place a clean and empty glass bottom based dish on the stage of the polarizing microscope and set background using the integrated software. 
	NOTE: Do not use plastic Petri dishes to image birefringence, as they do not appropriately transmit refracted light. Depending on the polarized microscope used, additional settings may be required as per the manufacturer&#39;s guidelines.</li>
	<li>Having set the background, remove the glass bottom based dish from the microscope stage, and using a glass pipette, transfer the anesthetized, stabbed fish onto the glass bottom based dish.</li>
	<li>Carefully remove the excess of embryo medium using a pipette, and orient the fish as per 2.5 - the head is on the left, tail on the right, dorsal region up and ventral region down. 
	NOTE: Ensure the larvae is mounted as flat as possible, as uneven mounting results in different birefringence intensities across different somites.</li>
	<li>Place the glass bottom based dish with the anesthetized larvae on the microscope stage and image the muscle using polarized light (Figures 1C &#38; 1D). 
	NOTE: Depending on the type of microscope/polarized lens used, the fish orientation on its anterior-posterior axis can affect the overall birefringence15. Ensure to include at least 5 somites on either side of the injury site when imaging.</li>
	<li>Using a plastic pipette, pour a drop of embryo medium to rehydrate the fish and place the fish in a well of a 24 well plate filled up with embryo medium. 
	NOTE: It is important to perform the imaging as quickly as possible to prevent the fish from drying.</li>
	<li>Repeat steps 3.3. to 3.6. until all the fish have been imaged.</li>
	<li>When all images have been acquired, save them in .tiff format for subsequent analyses.</li>
	<li>Put the fish back into the 28 &#176;C incubator until performing subsequent imaging at 3 dpi (7 dpf).</li>
	<li>When the fish are 3 dpi, image the fish as per 3.3 to 3.8.</li>
	<li>Once all fish have been imaged, euthanize them by adding tricaine methanesulfonate to a final concentration of 0.2% (v/v) in embryo medium. Wait for at least 10 minutes to ensure that the fish are euthanized, evident by the loss of swimming ability, pumping of the gill covers, and lack of a flight response following touch.</li>
</ol>
4. Quantification of muscle regeneration
<ol>
	<li>Open a 1 dpi image on an imaging analysis software such as the freely available Image J software.</li>
	<li>Using the polygon tool, draw around the wound site, and measure the area and mean birefringence intensity of this region (Figures 1C &#38; 1D). Copy these values to cells D3 and E3 in the template provided (Supplementary Table 1).</li>
	<li>Draw two additional regions, each spanning 1-2 uninjured somites, and measure the area and mean birefringence intensities of each of these regions (Figures 1C &#38; 1D). Copy these values to cells D4-D5 and E4-E5 in the template provided (Supplementary Table 1). 
	NOTE: While it is preferable to select the same uninjured somites in the 1 dpi and 3 dpi images, the sporadic detachment of muscle fibres and subsequent reduction in birefringence in mutants may make this impossible. Therefore, in the event the same somites cannot be selected at 1 dpi and 3 dpi due to the reduction in muscle integrity in mutants, select two different but unaffected areas at each of the timepoints.</li>
	<li>Calculate the normalized birefringence for each region by dividing the mean birefringence intensity of that region by the area - displayed in column F in the template provided (Supplementary Table 1).</li>
	<li>Repeat steps 4.1-4.4 for all 1 dpi and 3 dpi images. 
	NOTE: When using the template provided (Supplementary Table 1), the 1 dpi area and mean birefringence intensity values should be inserted in columns D and E respectively, and the 3 dpi area and mean birefringence intensity values should be inserted in columns J and K respectively.</li>
	<li>For each time point, calculate the average normalized birefringence of the two uninjured regions. This value provides a reference point of uninjured muscle. 
	NOTE: In the template provided (Supplementary Table 1), this value is computed in columns G and M, for the 1 dpi and 3 dpi images respectively.</li>
	<li>Next, determine the extent of muscle injury at 1 dpi, by dividing the normalized birefringence of injury region by the average normalized birefringence of the uninjured regions (calculated in 4.6). 
	NOTE: Using the above detailed needle stab procedure, wildtype larvae typically show a normalized birefringence of 48.5 &#177; 14.3% at the wound site at 1 dpi, indicating that the birefringence has reduced by approximately 50% when compared to uninjured somites. When using the template (Supplementary Table 1), this value is computed as a percentage in column H.</li>
	<li>To determine the extent of muscle regeneration, divide the normalized birefringence of the injury region in the 3 dpi image, by the average normalized intensity of the uninjured regions at this stage. 
	NOTE: At 3 dpi, wildtype larvae typically show a normalized birefringence of 60 &#177; 15.3% at the wound site. Given that the normalized birefringence within the wound site at 1 dpi was 48.5 &#177; 14.3% (step 4.7), the increase in birefringence at 3 dpi to 60 &#177; 15.3% indicates a recovery of approximately 11.5%. When using the template (Supplementary Table 1), this value is computed as a percentage in column N.</li>
	<li>Keeping fish within each genotype separate, perform a paired t-test comparing the normalized birefringence of the wound site at 3 dpi (step 4.8) with that of 1 dpi (step 4.7). This will reveal the trajectory of muscle regeneration displayed by each fish in each genotype, and highlight if the extent of muscle regeneration displayed by each genotype has significantly altered.</li>
	<li>Finally, calculate the regenerative index by dividing the value obtained in step 4.8, which is the extent of muscle regeneration at 3 dpi, by the value obtained in step 4.7, which is the extent of muscle injury at 1 dpi. A regenerative index of 1 indicates that the injury at 1 dpi is comparable to 3 dpi and that muscle regeneration has not occurred; a value above 1 indicates that at 3 dpi new muscle has formed in the wound site highlighting that the muscle has regenerated; and a regenerative index of less than 1 highlights that the wound at 3 dpi is worse than 1 dpi, and that muscle regeneration in impaired. A t-test or a one-way ANOVA can be performed to statistically compare the extent of muscle regeneration between the different genotypes. When using the template provided (Supplementary Table 1), this value is calculated in column O. 
	NOTE: It is recommended to perform the entire experiment in triplicates, with fish from each experiment obtained from different biological parents, and each experiment performed on different days, to avoid any bias.</li>
</ol>

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The authors have nothing to disclose.

Skeletal muscle regeneration is driven by obligate tissue resident muscle stem cells, whose function is altered in many muscle diseases such as muscular dystrophy, subsequently impeding the process of muscle regeneration. Here, we describe a high throughput protocol to examine muscle regeneration in live zebrafish models of muscle disease. The first step of the pipeline utilizes a embryo genotyping platform14, which is a user-friendly and accurate method to determine the genotype of live larvae, b...

Skeletal muscle accounts for 30-50% of human body mass, and is not only indispensable for locomotion, but it also serves as a critical metabolic and storage organ1. Despite being postmitotic, skeletal muscle is highly dynamic and retains a tremendous regenerative capacity following injury. This is attributed to the presence of tissue resident stem cells (also called satellite cells), located under the basal lamina of myofibers and marked by the transcription factors paired box protein 7 (pax7) and/or paired box protein 3 (pax3), among others2,3. Following injury, the satellite cell is activated and undergoes cell proliferation to generate a pool of myoblasts, which subsequently differentiate to form new muscle fibers. The highly conserved cascade of pro-regenerative signals regulating satellite cell activation and robust muscle repair is&#160;affected in various conditions such as myopathies and homeostatic ageing4,5.
One such diverse group of myopathies is muscular dystrophy, characterized by progressive muscle wasting and degeneration6. These diseases are the consequence of genetic mutations in key proteins, including dystrophin and laminin-&#945;2 (LAMA2), responsible for the attachment of muscle fibers to the extracellular matrix7,8. Given that proteins implicated in muscular dystrophy play such a central role in maintaining muscle structure, for many years it was believed that a failure in this process was the mechanism responsible for disease pathogenesis9. However, recent studies have identified defects in the regulation of muscle stem cells and subsequent impairment in muscle regeneration as a second possible basis for the muscle pathology observed in muscular dystrophy10,11. As such, further studies are needed to investigate how an impairment in muscle stem cell function and associated niche elements contributes to muscular dystrophy.
Over the past decade, zebrafish (Danio rerio) has emerged as an important vertebrate model for disease modeling12. This is attributed to the rapid external development of the zebrafish embryo, coupled with its optical clarity, which allows the direct visualization of muscle formation, growth, and function. Additionally, not only is the development and structure of muscle highly conserved in zebrafish, they also display a highly conserved process of muscle regeneration13. Consequently, zebrafish represent an excellent system to study the pathobiology of muscle diseases, and explore how muscle regeneration is affected in it. To this end, we have developed a method that enables the timely study of skeletal muscle regeneration in zebrafish models of muscle disease. This high throughput pipeline involves a method to genotype live embryos14, following which a needle-stab injury is performed and the extent of muscle regeneration is imaged using polarizing light microscopy. The utilization of this technique will therefore reveal the regenerative capacity of muscle in zebrafish models of muscle disease.

<table><tbody><tr><td>24 well plates</td><td>Thermo Fischer</td><td>142475</td><td></td></tr><tr><td>30 gauge needles</td><td>Terumo</td><td>NN-3013R</td><td></td></tr><tr><td>90 mm Petri Dishes</td><td>Pacific Laboratory Products PT</td><td>S9014S20</td><td></td></tr><tr><td>DNA extraction chips</td><td>wFluidx</td><td>ZEG chips</td><td></td></tr><tr><td>Embryo genotyping platform</td><td>wFluidx</td><td>ZEG base unit</td><td>Zebrafish Embryo Genotyper</td></tr><tr><td>Glass pipette</td><td>Hirschmann</td><td>9260101</td><td></td></tr><tr><td>Glass plate dish</td><td>WPI</td><td>FD35-100</td><td>Commonly referred to as FluoroDish</td></tr><tr><td>Incubator</td><td>Thermoline Scientific</td><td>TEI-43L</td><td></td></tr><tr><td>Plastic pipette</td><td>Livingstone</td><td>PTP03-01</td><td></td></tr><tr><td>Polarizing microscope</td><td>Abrio</td><td>N/A</td><td></td></tr></tbody></table>

examining muscle regeneration in zebrafish models of muscle disease

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

The ability to quantify birefringence of skeletal muscle provides a non-invasive but highly reproducible method to examine and compare levels of muscle damage, and examine muscle regeneration in vivo. Birefringence results from the diffraction of polarised light through the pseudo-crystalline array of the muscle sarcomeres15, and following injury or damage to the muscle, a reduction in birefringence is evident. Likewise, the activation and differentiation ...

Watch this Scientific Journal Video about Examining Muscle Regeneration in Zebrafish Models of Muscle Disease at JoVE.com

Examining Muscle Regeneration in Zebrafish Models of Muscle Disease

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

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

examining-muscle-regeneration-in-zebrafish-models-of-muscle-disease

Universidad de Cartagena UDEC

Research

JoVE Journal

Developmental Biology

4.9K Views.  Monash University. Skeletal muscle regeneration is driven by tissue resident muscle stem cells, which are impaired in many muscle diseases such as muscular dystrophy, and this results in the inability of muscle to regenerate. Here, we describe a protocol that allows the examination of muscle regeneration in zebrafish models of muscle disease.

Video: Examining Muscle Regeneration in Zebrafish Models of Muscle Disease

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 ...

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 ...

Using Touch-evoked Response and Locomotion Assays to Assess Muscle Performance and Function in Zebrafish

Zebrafish muscle development is highly conserved with mammalian systems making them an excellent model to study muscle function and disease. Many myopathies affecting skeletal muscle function can be quickly and easily assessed in zebrafish over the first few days of embryogenesis. By 24 hr post-fertilization (hpf), wildtype zebrafish spontaneously contract their tail muscles and by 48 hpf, zebrafish exhibit controlled swimming behaviors. Reduction in the frequency of, or other alterations in, these movements may indicate a skeletal muscle dysfunction. To analyze swimming behavior and assess muscle performance in early zebrafish development, we utilize both touch-evoked escape response and locomotion assays.
Touch-evoked escape response assays can be used to assess muscle performance during short burst movements resulting from contraction of fast-twitch muscle fibers. In response to an external stimulus, which in this case is a tap on the head, wildtype zebrafish at 2 days post-fertilization (dpf) typically exhibit a powerful burst swim, accompanied by sharp turns. Our method quantifies skeletal muscle function by measuring the maximum acceleration during a burst swimming motion, the acceleration being directly proportional to the force produced by muscle contraction.
In contrast, locomotion assays during early zebrafish larval development are used to assess muscle performance during sustained periods of muscle activity. Using a tracking system to monitor swimming behavior, we obtain an automated calculation of the frequency of activity and distance in 6-day old zebrafish, reflective of their skeletal muscle function. Measurements of swimming performance are valuable for phenotypic assessment of disease models and high-throughput screening of mutations or chemical treatments affecting skeletal muscle function.

Zebrafish are an excellent model to study muscle function and disease. During early embryogenesis zebrafish begin regular muscle contractions producing rhythmic swimming behavior, which is altered when the muscle is disrupted. Here we describe a touch-evoked response and locomotion assay to examine swimming performance as a measure of muscle function.

Zebrafish muscle development is highly conserved with mammalian systems making them an excellent model to study muscle function and disease. Many ...

Normal and Malignant Muscle Cell Transplantation into Immune Compromised Adult Zebrafish

Zebrafish have become a powerful tool for assessing development, regeneration, and cancer. More recently, allograft cell transplantation protocols have been developed that permit engraftment of normal and malignant cells into irradiated, syngeneic, and immune compromised adult zebrafish. These models when coupled with optimized cell transplantation protocols allow for the rapid assessment of stem cell function, regeneration following injury, and cancer. Here, we present a method for cell transplantation of zebrafish adult skeletal muscle and embryonal rhabdomyosarcoma (ERMS), a pediatric sarcoma that shares features with embryonic muscle, into immune compromised adult rag2E450fs homozygous mutant zebrafish. Importantly, these animals lack T cells and have reduced B cell function, facilitating engraftment of a wide range of tissues from unrelated donor animals. Our optimized protocols show that fluorescently labeled muscle cell preparations from &#945;-actin-RFP transgenic zebrafish engraft robustly when implanted into the dorsal musculature of rag2 homozygous mutant fish. We also demonstrate engraftment of fluorescent-transgenic ERMS where fluorescence is confined to cells based on differentiation status. Specifically, ERMS were created in AB-strain myf5-GFP; mylpfa-mCherry double transgenic animals and tumors injected into the peritoneum of adult immune compromised fish. The utility of these protocols extends to engraftment of a wide range of normal and malignant donor cells that can be implanted into dorsal musculature or peritoneum of adult zebrafish.

Here, we present a protocol for cell transplantation of zebrafish skeletal muscle and embryonal rhabdomyosarcoma (ERMS) into adult immune compromised rag2E450fs homozygous mutant zebrafish. This protocol allows for the efficient analysis of regeneration and malignant transformation of muscle cells.

Zebrafish have become a powerful tool for assessing development, regeneration, and cancer. More recently, allograft cell transplantation protocols have ...

Immunology and Infection

Real Time and Repeated Measurement of Skeletal Muscle Growth in Individual Live Zebrafish Subjected to Altered Electrical Activity

A number of methods can be used to visualize individual cells throughout the body of live embryonic, larval or juvenile zebrafish. We show that live fish with fluorescently-marked plasma membranes can be scanned in a confocal laser scanning microscope in order to determine the volume of muscle tissue and the number of muscle fibers present. Efficient approaches for the measurement of cell number and size in live animals over time are described and validated against more arduous segmentation methods. Methods are described that permit the control of muscle electrical, and thus contractile, activity. Loss of skeletal muscle contractile activity greatly reduced muscle growth. In larvae, a protocol is described that allows reintroduction of patterned electrical-evoked contractile activity. The described methods minimize the effect of inter-individual variability and will permit analysis of the effect of electrical, genetic, drug, or environmental stimuli on a variety of cellular and physiological growth parameters in the context of the living organism. Long-term follow-up of the measured effects of a defined early-life intervention on individuals can subsequently be performed.

Optical clarity is a major advantage for cell biological and physiological work in zebrafish. Robust methods for measurement of cell growth in individual animals are described that permit novel insights into how growth of skeletal muscle and neighboring tissues are integrated with whole body growth.

A number of methods can be used to visualize individual cells throughout the body of live embryonic, larval or juvenile zebrafish. We show that live fish ...

Laser-inflicted Injury of Zebrafish Embryonic Skeletal Muscle

Various experimental approaches have been used in mouse to induce muscle injury with the aim to study muscle regeneration, including myotoxin injections (bupivacaine, cardiotoxin or notexin), muscle transplantations (denervation-devascularization induced regeneration), intensive exercise, but also murine muscular dystrophy models such as the mdx mouse (for a review of these approaches see 1). In zebrafish, genetic approaches include mutants that exhibit muscular dystrophy phenotypes (such as runzel2 or sapje3) and antisense oligonucleotide morpholinos that block the expression of dystrophy-associated genes4. Besides, chemical approaches are also possible, e.g. with Galanthamine, a chemical compound inhibiting acetylcholinesterase, thereby resulting in hypercontraction, which eventually leads to muscular dystrophy5. However, genetic and pharmacological approaches generally affect all muscles within an individual, whereas the extent of physically inflicted injuries are more easily controlled spatially and temporally1. Localized physical injury allows the assessment of contralateral muscle as an internal control. Indeed, we recently used laser-mediated cell ablation to study skeletal muscle regeneration in the zebrafish embryo6, while another group recently reported the use of a two-photon laser (822 nm) to damage very locally the plasma membrane of individual embryonic zebrafish muscle cells7.
Here, we report a method for using the micropoint laser (Andor Technology) for skeletal muscle cell injury in the zebrafish embryo. The micropoint laser is a high energy laser which is suitable for targeted cell ablation at a wavelength of 435 nm. The laser is connected to a microscope (in our setup, an optical microscope from Zeiss) in such a way that the microscope can be used at the same time for focusing the laser light onto the sample and for visualizing the effects of the wounding (brightfield or fluorescence). The parameters for controlling laser pulses include wavelength, intensity, and number of pulses.
Due to its transparency and external embryonic development, the zebrafish embryo is highly amenable for both laser-induced injury and for studying the subsequent recovery. Between 1 and 2 days post-fertilization, somitic skeletal muscle cells progressively undergo maturation from anterior to posterior due to the progression of somitogenesis from the trunk to the tail8, 9. At these stages, embryos spontaneously twitch and initiate swimming. The zebrafish has recently been recognized as an important vertebrate model organism for the study of tissue regeneration, as many types of tissues (cardiac, neuronal, vascular etc.) can be regenerated after injury in the adult zebrafish10, 11.

The method presented here comprises the precise injury of live zebrafish embryos with high-energy laser pulses and the subsequent analysis of these injuries and their recovery with time. We also show how genetically labeled single or groups of skeletal muscle cells can be tracked during and after laser light induced damage.

Various experimental approaches have been used in mouse to induce muscle injury with the aim to study muscle regeneration, including myotoxin injections ...

Biology

Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

The zebrafish has proven to be a valuable model system for exploring skeletal muscle function and for studying human muscle diseases. Despite the many advantages offered by in vivo analysis of skeletal muscle in the zebrafish, visualizing the complex and finely structured protein milieu responsible for muscle function, especially in whole embryos, can be problematic. This hindrance stems from the small size of zebrafish skeletal muscle (60 &mu;m) and the even smaller size of the sarcomere. Here we describe and demonstrate a simple and rapid method for isolating skeletal myofibers from zebrafish embryos and larvae. We also include protocols that illustrate post preparation techniques useful for analyzing muscle structure and function. Specifically, we detail the subsequent immunocytochemical localization of skeletal muscle proteins and the qualitative analysis of stimulated calcium release via live cell calcium imaging. Overall, this video article provides a straight-forward and efficient method for the isolation and characterization of zebrafish skeletal myofibers, a technique which provides a conduit for myriad subsequent studies of muscle structure and function.

Zebrafish are an emerging system for modeling human disorders of the skeletal muscle. We describe a fast and efficient method to isolate skeletal muscle myofibers from embryonic and larval zebrafish. This method yields a high-density myofiber preparation suitable for study of single skeletal muscle fiber morphology, protein subcellular localization, and muscle physiology.

The zebrafish has  proven to be a valuable model system for exploring skeletal muscle function and  for studying human muscle diseases. Despite  the many ...

Analysis of Skeletal Muscle Defects in Larval Zebrafish by Birefringence and Touch-evoke Escape Response Assays

Zebrafish (Danio rerio) have become a particularly effective tool for modeling human diseases affecting skeletal muscle, including muscular dystrophies1-3, congenital myopathies4,5, and disruptions in sarcomeric assembly6,7, due to high genomic and structural conservation with mammals8. Muscular disorganization and locomotive impairment can be quickly assessed in the zebrafish over the first few days post-fertilization. Two assays to help characterize skeletal muscle defects in zebrafish are birefringence (structural) and touch-evoked escape response (behavioral).
Birefringence is a physical property in which light is rotated as it passes through ordered matter, such as the pseudo-crystalline array of muscle sarcomeres9. It is a simple, noninvasive approach to assess muscle integrity in translucent zebrafish larvae early in development. Wild-type zebrafish with highly organized skeletal muscle appear very bright amidst a dark background when visualized between two polarized light filters, whereas muscle mutants have birefringence patterns specific to the primary muscular disorder they model. Zebrafish modeling muscular dystrophies, diseases characterized by myofiber degeneration followed by repeated rounds of regeneration, exhibit degenerative dark patches in skeletal muscle under polarized light. Nondystrophic myopathies are not associated with necrosis or regenerative changes, but result in disorganized myofibers and skeletal muscle weakness. Myopathic zebrafish typically show an overall reduction in birefringence, reflecting the disorganization of sarcomeres.
The touch-evoked escape assay involves observing an embryo&#39;s swimming behavior in response to tactile stimulation10-12. In comparison to wild-type larvae, mutant larvae frequently display a weak escape contraction, followed by slow swimming or other type of impaired motion that fails to propel the larvae more than a short distance12. The advantage of these assays is that disease progression in the same fish type can be monitored in vivo for several days, and that large numbers of fish can be analyzed in a short time relative to higher vertebrates.

The zebrafish is now an established and powerful tool for modeling muscular dystrophies, congenital myopathies, and related neuromuscular diseases. Birefringence and touch-evoked escape behavior are two common noninvasive assays used to determine the degree of muscular disorganization and locomotive impairment of zebrafish embryos during early development.

Zebrafish (Danio rerio) have become a particularly effective tool for modeling human diseases affecting skeletal muscle, including muscular ...

An Assay for Lateral Line Regeneration in Adult Zebrafish

Due to the clinical importance of hearing and balance disorders in man, model organisms such as the zebrafish have been used to study lateral line development and regeneration. The zebrafish is particularly attractive for such studies because of its rapid development time and its high regenerative capacity. To date, zebrafish studies of lateral line regeneration have mainly utilized fish of the embryonic and larval stages because of the lower number of neuromasts at these stages. This has made quantitative analysis of lateral line regeneration/and or development easier in the earlier developmental stages. Because many zebrafish models of neurological and non-neurological diseases are studied in the adult fish and not in the embryo/larvae, we focused on developing a quantitative lateral line regenerative assay in adult zebrafish so that an assay was available that could be applied to current adult zebrafish disease models. Building on previous studies by Van Trump et al.17&#160;that described procedures for ablation of hair cells in adult Mexican blind cave fish and zebrafish (Danio rerio), our assay was designed to allow quantitative comparison between control and experimental groups. This was accomplished by developing a regenerative neuromast standard curve based on the percent of neuromast reappearance over a 24 hr time period following gentamicin-induced necrosis of hair cells in a defined region of the lateral line. The assay was also designed to allow extension of the analysis to the individual hair cell level when a higher level of resolution is required.

Because many zebrafish models of neurological and non-neurological diseases are studied in the adult fish rather than the embryo/larvae, we developed a quantitative lateral line regenerative assay that can be applied to adult zebrafish disease models. The assay involved resolution at the 1) neuromast and 2) individual hair cell levels.

Due to the clinical importance of hearing and balance disorders in man, model organisms such as the zebrafish have been used to study lateral line ...

Assessment of Swim Endurance and Swim Behavior in Adult Zebrafish

Due to their renowned regenerative capacity, adult zebrafish are a premier vertebrate model to interrogate mechanisms of innate spinal cord regeneration. Following complete transection of their spinal cord, zebrafish extend glial and axonal bridges across severed tissue, regenerate neurons proximal to the lesion, and regain their swim capacities within 8 weeks of injury. Recovery of swim function is thus a central readout for functional spinal cord repair. Here, we describe a set of behavioral assays to quantify zebrafish motor capacity inside an enclosed swim tunnel. The goal of these methods is to provide quantifiable measurements of swim endurance and swim behavior in adult zebrafish. For swim endurance, zebrafish are subjected to a constantly increasing water current velocity until exhaustion, and time at exhaustion is reported. For swim behavior assessment, zebrafish are subjected to low current velocities and swim videos are captured with a dorsal view of the fish. Percent activity, burst frequency, and time spent against the water current provide quantifiable readouts of swim behavior. We quantified swim endurance and swim behavior in wild-type zebrafish before injury and after spinal cord transection. We found that zebrafish lose swim function after spinal cord transection and gradually regain that capacity between 2 and 6 weeks post-injury. The methods described in this study could be applied to neurobehavioral, musculoskeletal, skeletal muscle regeneration, and neural regeneration studies in adult zebrafish.

Capable of functional recovery after spinal cord injury, adult zebrafish is a premier model system to elucidate innate mechanisms of neural regeneration. Here, we describe swim endurance and swim behavior assays as functional readouts of spinal cord regeneration.

Due to their renowned regenerative capacity, adult zebrafish are a premier vertebrate model to interrogate mechanisms of innate spinal cord regeneration. ...