Examining Muscle Regeneration in Zebrafish Models of Muscle Disease

Summary

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.

Abstract

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.

Introduction

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 others^2,3. 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 affected in various conditions such as myopathies and homeostatic ageing^4,5.

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-α2 (LAMA2), responsible for the attachment of muscle fibers to the extracellular matrix^7,8. 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 dystrophy^10,11. 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.

Protocol

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.

1. 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.
2. Prepare the 24 chamber DNA extraction chip by peeling the clear protective film from the top surface of the chip (Figure 1A).
3. Cut the tip of a 20 µL filter tip to widen the opening. Using this, pick a single embryo in a 13 µL 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.
4. 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.
5. 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.
6. Set the base unit to 2.4 volts, 0.051 A, and 0.12 W and start the DNA extraction protocol by pressing the "ON/OFF" 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.
7. 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.
8. Additionally, appropriately label 8 well strip tubes, which will be used for collection of DNA material from each embryo.
9. At the completion of the 8 minute protocol, press the ON/OFF button to stop the vibration of the platform.
10. Open the lid of the platform, and gently remove the magnetic lid ensuring minimal pressure is applied to the platform.
11. Carefully remove the DNA extraction chip from the platform and place it on a flat surface.
12. From the first chamber, remove 10 µL 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.
13. 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.
14. 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 °C incubator.
15. 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.
16. 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 °C incubator.

2. Performing muscle injury using a needle stab

1. 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.
2. 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.
3. Using a plastic pipette, transfer one anesthetized larva into a new Petri dish.
4. 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).
5. 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.
6. 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.
7. Repeat steps 2.3 to 2.6. until all the fish have been stabbed.
   ​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.
8. Once all the larvae have been stabbed, place the 24 well plate in the 28 °C incubator until subsequent imaging is performed.

3. Imaging of muscle injury and recovery

1. 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.
2. 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's guidelines.
3. 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.
4. 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.
5. Place the glass bottom based dish with the anesthetized larvae on the microscope stage and image the muscle using polarized light (Figures 1C & 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.
6. 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.
7. Repeat steps 3.3. to 3.6. until all the fish have been imaged.
8. When all images have been acquired, save them in .tiff format for subsequent analyses.
9. Put the fish back into the 28 °C incubator until performing subsequent imaging at 3 dpi (7 dpf).
10. When the fish are 3 dpi, image the fish as per 3.3 to 3.8.
11. 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.

4. Quantification of muscle regeneration

1. Open a 1 dpi image on an imaging analysis software such as the freely available Image J software.
2. Using the polygon tool, draw around the wound site, and measure the area and mean birefringence intensity of this region (Figures 1C & 1D). Copy these values to cells D3 and E3 in the template provided (Supplementary Table 1).
3. 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 & 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.
4. 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).
5. 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.
6. 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.
7. 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 ± 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.
8. 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 ± 15.3% at the wound site. Given that the normalized birefringence within the wound site at 1 dpi was 48.5 ± 14.3% (step 4.7), the increase in birefringence at 3 dpi to 60 ± 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.
9. 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.
10. 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.

Results

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 ...

Discussion

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...

Materials

List of materials used in this article

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