Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

Abstract

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 μm) 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.

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Introduction

Skeletal muscle is a highly specialized tissue responsible for generating the contractile forces necessary for motility. Contraction is initiated through a process known as excitation-contraction (EC) coupling that converts electric signals to calcium release from intracellular stores^1,2 . Intracellular calcium release activates the sarcomere to shorten and generate force. The many specific components of the molecular machinery responsible for mediating neuromuscular junction transmission³, EC coupling^4,5 , and actin-myosin dependent contractions⁶ continue to be the ongoing subject of intense research. In addition, proteins that stabilize the muscle membrane during contraction^7,8 and that mediate signaling between the myofiber and the extracellular matrix^7,9 have been identified and studied in great detail.

Mutations in a number of genes important for muscle structure and function have been identified as causes of human skeletal muscle diseases (http://www.musclegenetable.org/). These diseases, classified broadly as skeletal myopathies and muscular dystrophies based on clinical and histopathologic features, are associated with muscle weakness, lifelong disability, and early mortality^10,11 . The zebrafish has proven to be an outstanding system for modeling and studying human skeletal muscle diseases^8,12,13 . It has been employed to validate new gene mutations⁸, define new aspects of disease pathophysiology^14,15 , and identify new therapeutic approaches^15,16 . The power of the zebrafish for studying human muscle disease relates to the large number of offspring, the rapid development of muscle structure and function, the optical clarity of the zebrafish embryo, and the ease of genetic and pharmacologic manipulation of the developing zebrafish¹⁷.

We and others^12,18,19 have recently developed a simple technique for the rapid and efficient isolation of myofibers from the developing zebrafish. This methodology has enabled the examination of myofibers in greater detail than can be provided by whole embryo analysis. The technique has been exploited for the characterization of protein subcellular localization²⁰ as well as for the identification of important histopathologic characteristics as part of validation studies in newly developed disease models²¹. Furthermore, isolated myofibers can additionally be used for live imaging and for electrophysiological studies²², techniques that allow for the interrogation of key aspects of muscle function. The specific protocol for myofiber isolation, along with two examples of subsequent analytic experiments, is detailed in the remaining parts of this manuscript.

Protocol

1. Preparation of Poly-L-Lysine Coated Coverslips (Time: 1 hr)

Coating coverslips allows for rapid myofiber settling and adhesion. This may be performed during the dissociation step of the myofiber isolation (step 2 below).

1. Cut and place Parafilm on the bottom of a 60 mm Petri dish (any brand).

2. Place microscope cover glass slips (12 mm diameter) on Parafilm in a 60 mm tissue culture dish or place them individually in single wells of 24-well plates.

Note 1: These small round coverslips help to concentrate myofiber numbers and reduce excess antibody usage.

Note 2: Other size coverslips will work; however, the volumes of reagents and embryos used will need to be adjusted accordingly.

1. Pipette 50-200 μl of poly-L-lysine solution (0.01%) on each cover glass in the Petri dish.

2. Allow the coverslips to sit at least one hour at 37 °C. Longer incubations will not negatively influence results.

3. Remove poly-L-lysine solution from the cover glass and wash twice with a minimum of 100 μl 1x PBS.

4. Allow coverslips to dry.

5. Coverslips are now ready for myofibers.

Note 3: To ensure sterility, coating can be done in a hood and on autoclaved coverslips.

Note 4: Keep the 60 mm Petri dish with Parafilm on the bottom. This setup will be used during myofiber plating and immunolabeling (later steps).

Alternative 1: Instead of coating coverslips immediately prior to use, a coated coverslip stock can be used. A 60 mm Petri dish containing a minimum of 2 ml poly-L-lysine can be stored at 4 °C containing numerous coverslips. With this alternative start at step 1.5 to process the coverslips for myofibers.

Alternative 2: We also coat coverslips in poly-L-ornithine. This is more labor intensive, but is useful for longer term culturing because poly-L-ornithine coated coverslips can be UV treated. With UV treatment and careful sterile technique live myofibers can typically be maintained in culture from 4-7 days.

2. Dissociation of Zebrafish Embryos and Plating of Myofibers (Time: 1 to 3 hr)

Note: the standard protocol applies best to 3 dpf (days post fertilization) embryos.

1. Transfer zebrafish embryos into a standard 1.5 ml centrifuge tube and remove as much excess fish water as possible. Typically, 10-20 embryos per tube, though less can be used. Using more than 20-25 embryos often results in a preparation of excessive density.

   1. Remove chorions prior to dissociation, manually using #5 forceps. Alternatively, chemical chorion removal is achieved with a 10-15 min Pronase treatment. Typically, the previous removal of the chorion is only needed when prior confirmation of a mutant phenotype or morphology is required, or when using stages where embryos are naturally hatched from their chorion.

2. Add 900 μl of CO₂ independent media to the tube containing the embryos.

3. Add 100 μl of collagenase type II, final concentration 3.125 mg/ml (Stock collagenase solution at 31.25 mg/ml diluted in 1x PBS) to begin dissociation. Collagenase IV can also be used for the dissociation.

4. Rotate embryos on an orbital shaker, and triturate every 30 min at RT using a P1000 Pipetman for the trituration.

Note 2: Carefully monitor embryo dissociation; over or under dissociation (especially over) are the most common reasons for protocol failure. Times for digestion will vary depending on intensity of trituration, number of embryos per tube, and age of embryos. It is also often less (in comparison to wild types) for embryos from skeletal muscle mutants.

Note 3: Average digestion time per embryo age: 1 dpf = 1 hr, 2 dpf = 1.5 hr, 3 dpf = 2 hr, and 4 dpf = 2-2.5 hr.

1. Stop digestion when no whole embryos are visible, yet solid pieces are still visible - this is essential to prevent overdigestion.

2. Centrifuge tubes with dissociated embryos at 0.8-2.3 x g for 3-5 min to pellet cells.

3. Remove supernatant from pelleted cells and wash 2x with CO₂ independent media.

4. Add fresh CO₂ independent media to resuspend cells. 1 ml is typically used for preps of 10-20 embryos. The volume can be scaled based on embryo number.

5. Using a P1000 pipette, pass embryo suspension through a 70 μm filter . This helps remove unwanted debris from the prep.

Note 4: Embryo suspension can be filtered a second time through a 40 μm filter to further remove unwanted debris. From a double filtration, recovery is approximately 800 μl from a starting volume of 1 ml.

1. Add approximately 50-100 μl of myofiber suspension to each poly-L-lysine coated coverslip.

Note 5: Perform step 2.9 in the Petri dishes with Parafilm on the bottom, previously prepared. The Parafilm keeps the myofiber suspension from running off the coated coverslips. Keep Petri dishes covered to prevent evaporation.

1. Allow myofibers to settle approximately 1 hr onto the coated coverslips at room temperature.

Note 6: Myofibers will begin to settle within 5-10 min. However, for good myofiber attachment, a minimum of 1 hr (1-2 hr) is recommended. Allowing myofibers to settle for longer will not harm the prep. For longer incubations (including overnight), antibiotics can be added to the media. With antibiotics and sterile technique live cultures can typically be maintained for 1-2 days.

1. Live myofibers can be observed at this point. Myofibers from embryos 2 dpf and later will be seen as striated and elongated cells (Figure 1). At this point, myofibers are now ready for live analysis or for immunolabeling.

Note 7: Skeletal muscle from 1 dpf embryos does not plate as elongated and clearly striated fibers. Instead, large myoballs are visible. In addition, during the pelleting phase post dissociation (step 2.6), 1 dpf myoballs need to be centrifuged for a minimum of 8 min at 5,000 x g to achieve a pellet. For analysis of embryos at this stage, it is recommended to use the transgenic line expressing EGFP specifically in skeletal muscle²³. This will allow identification of cells from muscle origin versus other sources.

3.1. Immunolabeling

1. Remove a portion of media from myofibers adhered to the coated coverslip

2. Fix cells using 4% paraformaldehyde or methanol. For PFA, remove ½ the volume of media and replace with the same amount of PFA. Fix for 10 min at RT, then remove total volume, replace with 50-100 μl of PFA, and fix for an additional 10 min. For alcohol fixation, ice cold methanol or acetone can be applied at 4 °C for 10 min or 5 min, respectively.

3. Wash myofibers at least 3-5x with 1x PBS or 1x PBS plus 0.1% TWEEN. Average wash volume is 50-100 μl per coverslip.

4. Add blocking solution to myofibers (working stock) and incubate 20-60 min at room temperature. Blocking solution will vary depending on the primary and secondary antibodies. The two most common used in the lab are (1) 1x PBS, 2 mg/ml BSA, 1% sheep serum, 0.25% Triton X-100 final and (2) 0.2% Triton X-100, 2 mg/ml (0.2% BSA) and 5% sheep/goat serum.

5. Remove blocking solution and add primary antibody diluted in either blocking solution or PBS. Incubate myofibers overnight at 4 °C. Make sure to either Parafilm or wrap in saran wrap the Petri dish containing the myofiber-loaded coverslips coated with antibody. Small pieces of moistened paper towel can be added to create a humidified chamber.

6. Remove primary antibody from coverslips and wash coverslips 3-5x for 5 min with PBS or blocking solution.

7. Add secondary antibody at appropriate concentration diluted in 1x PBS or blocking solution for 1-2 hr at RT.

8. Remove secondary antibody and wash myofibers 3-5x in PBS for 5 min each at RT.

3.2. Mounting coverslips

1. Apply 1-2 drops of antifade reagent to a microscope slide.

2. Carefully pick-up the coated and immunolabled coverslip using forceps and place (myofibers down) the coverslip on the antifade reagent on the microscope slide.

Note 1: Attention to the orientation of the coated coverslip is critical.

1. Lay 1-2 Kimwipes on a hard solid surface. Quickly invert the slide with the coated coverslips resting on the antifade reagent and place it (coverslip down) on the Kimwipes.

2. Apply light pressure to the corners of the microscope slide to remove excess antifade and form a tight seal between the slide and coverslip

3. Allow the antifade remaining to dry for 5-10 min at RT, then the myofibers are ready to image (Figures 2A and B) or store at 4 °C.

4. Live Cell Calcium Imaging Using GCaMP3

Live cell experiments can be performed on myofibers prior to fixation (following step 2.10). The following protocol describes live imaging in myofibers expressing GCaMP3²⁴, a genetically encoded calcium indicator, expressed by the skeletal muscle specific zebrafish α-actin promoter (pSKM)²³. Alternatively, this technique can be readily adapted to use calcium indicator dyes such as Fura-2.

1. Inject embryos with pSKM: GCaMP3 construct at the 1-2 cell stage

2. At 3 dpf, collect larvae and prepare myofibers as described in sections 1 and 2. Allow myofibers to adhere for a minimum of 1 hr. For this technique, preparing coverslips in 24-well dishes is required.

3. Carefully remove any excess media, and add 300 μl of fresh CO₂ independent media at RT.

4. Observe cells under an inverted microscope. GCaMP3 positive myofibers should be visible under green fluorescence.

5. Set up camera for recording, if desired.

6. Prepare a 30 mM caffeine solution in CO₂ independent media. To stimulate cells, gently pipette 300 μl of caffeine solution into the well under magnification. Within 5-10 sec, GCaMP positive myofibers should respond to caffeine with a rapid increase in fluorescence (Figure 3). Myofibers may also contract. Of note, caffeine induces calcium release from the sarcoplasmic reticulum stores²⁵. Other agents, such as KCl²⁶ and ryanodine⁴, can be used to study inducible calcium release.

Representative Results

Fluorescent immunolabeling of myofibers (Figure 2)

Images showing expected fluorescent labeling pattern from myofibers immunostained after successful isolation and plating. The myofibers have been labeled with either anti-ryanodine receptor (1:100) (Figure 2A) or anti-α-actinin (1:100) (Figure 2B) antibodies, and reveal immunostaining of the triad and the Z-band respectively. Secondary antibodies used were Alexa Fluor 555 (1:500). Images captured using confocal microscopy.

Induced calcium release from a myofiber (Figure 3)

Panel series showing the representative calcium release from an isolated myofiber treated with caffeine. In brief, zebrafish embryos were injected at the one cell stage with the pSKM: GCaMP3 construct. At 3 dpf, embryos were dissociated and plated as described in the protocol. For analysis, GCaMP3 expressing myofibers were selected, as indicated by the presence of green fluorescence at baseline. Using a micropipette for drug administration, the selected fiber was bathed in a 30 mM caffeine solution to induce calcium release. Video recording was started prior to caffeine application and continued until peak fluorescence was reached. This figure demonstrates normal caffeine induced calcium release, and the technique can be used to interrogate the excitation-contraction coupling apparatus via live imaging.

Figure 1. Schematic Timeline of Embryo Dissociation. Successive panels (top to bottom) illustrate individual steps, timing, and equipment required for embryo dissociation. A) Selection of embryos for dissociation. Scale bar 500 μm. B) Image of dissociation setup. Embryos are in media and collagenase while being rotated until sufficiently dissociated (Protocol step 2). C) Image of both myofiber plating techniques; round coverslips or 24-well plates (Protocol steps 1 and 4, respectively). D) A bright-field image of live dissociated zebrafish myofibers plated on poly-L Lysine coated coverslips. Arrows denote elongated myofibers from 48 hpf zebrafish. Scale bar 30 μm.

Figure 2. Immunolabeling of individual myofibers isolated from 48 hpf zebrafish embryos. A) Myofiber labeled with anti-ryanodine receptor (RyR1), revealing a banded pattern along the fiber consistent with localization to the triad/EC coupling apparatus. B) Myofiber labeled with anti-α-actinin, visualizing striations (red) along the myofiber localizing to the sarcomere and highlighting the Z bands. In both images, nuclei are labeled with DAPI, seen as blue ovals. Inserts show higher magnification images of the myofiber in the large image. A) and B) scale bar 30 μm.

Figure 3. Representative induced calcium release response from a single isolated zebrafish myofiber. All panels show a pseudocolored zebrafish myofiber expressing GCaMP3. The time course shows the increase of GCaMP3 fluorescence (red color) in response to application of 30 mM caffeine. Recording started prior to caffeine application (0 msec) and continued to maximum fluorescence response (992 msec). Scale bar 30 μm. Click here to view larger figure.

Discussion

Zebrafish are a powerful vertebrate model system for studying muscle development and function in vivo^25,27,28 . They have also emerged as a valuable asset for modeling human muscle diseases^14,15,20,29 . While great strides have been taken to advance the use and application of zebrafish for the study of muscle function and muscle disease, there is a constant critical need to develop tools that will allow more in depth analysis that compliments the genetic, morphologic, behavioral and functional advantages zebrafish already offer¹⁷. We have therefore adapted and developed a simple and robust technique for the characterization of zebrafish myofibers from dissociated whole embryos.

The use of individual myofibers is commonplace in studies using the murine model system. In mice this has allowed for investigations and analyses that are impractical or impossible in whole animals, including subcellular protein localization studies³⁰, live cell imaging³¹, and electrophysiologic measurements³². In the zebrafish, similar dissociation techniques have been previously developed to specifically identify and examine motorneurons³³ as well as Rohon-Beard sensory neurons³⁴, and these techniques have enabled detailed analysis of these neuron subtypes.

The broad applicability of isolated myofibers, combined with the many unique advantages the zebrafish offers as a vertebrate model¹⁷, are what motivated us to develop a technique for isolating and characterizing embryonic and larval zebrafish myofibers. We have utilized isolated myofibers in previous studies to determine subcellular localization of muscle proteins^20,21 , to study localization of chimerically expressed transgenes³⁵ and to define specific myofiber parameters including especially size^20,21 . In addition, we have used the fibers as a source of validation for fish models of human muscle diseases^20,21 . For example, we have identified nemaline bodies by immunofluorescent analysis in a model of nemaline myopathy²⁰. We have also used myofibers as a means of interrogating specific intracellular pathways, including apoptosis and oxidative stress¹⁵, and for the application and testing of specific chemical modulators¹⁵.

Grabner and colleagues have also used the myofiber system to great success^19,22,36 . They are the only group to report prolonged culturing of isolated fibers. In addition, they have been able to use the fibers to measure the electrophysiological properties of the muscle. Specifically, they have tested the impact of the loss of the beta subunit of the dihydropyridine receptor on calcium release and measured the ability of a set of transgenes to rescue defects in this release. These studies illuminate some of the very powerful potential applications of isolated myofibers.

Another group to report use of the myofiber prep for studying muscle disease is Nixon et al. who used myofibers to study the impact of morpholino mediated caveolin 3 knockdown on muscle development and muscle structure¹⁸. These authors not only examined parameters by light microscopy, but also studied the fibers using electron microscopy. Their work identifies yet another advantage of the isolated myofiber system: the ability to study the ultrastructure of the myofiber in an isolated and controlled setting.

These studies, along with our work, point to the potential wide-ranging utility of this technique. Additional applications are certain to be developed, such as measuring rapid calcium spikes using indicators like Fura-2. Additionally, this preparation offers the opportunity to observe electrical activity, such as action potentials via voltage sensitive dyes (Di-4-ANEPPS) or direct electrophysiological recording. It seems clear that all current techniques used on murine myofibers should be applicable to zebrafish myofibers. In addition, the ability to express different transgenes in the developing zebrafish, as well as the availability of a large and growing number of transgenic zebrafish expressing various marker proteins (www.zfin.org), adds an additional layer of complexity and applicability to the system. We demonstrate this fact with the myofibers cultured from GCaMP3 expressing zebrafish, where we take advantage of the ability to express GCaMP3 in muscle to study induced calcium imaging in real time in isolated fibers (Figure 3).

As with most techniques, there are also some clear limitations to the isolated myofiber system. These issues are in addition to the standard shortcomings of studying a cell type in vitro that functions as a three dimensional syncytia in vivo and of performing electrophysiologic experiments with nonphysiologic stimuli. Using our methodology, one is not able to obtain a pure preparation of myofibers. This is not a problem for immunostaining or for electrophysiological measurements, but it does preclude certain experimental approaches. For example, we have yet to determine a suitable methodology for identifying a pure population of myofibers for transcriptomic analysis. We have attempted FACS sorting of GFP expressing myofibers followed by myofiber preparation, but this has not resulted in a pure culture with a suitable number of myofibers for either RNA or protein analysis.

Another challenge is the long term culturing of myofibers. We have been able to keep cultures for approximately 24 hr, and Grabner and colleagues report a preparation that has been suitable for multiple days of experimentation¹⁹. The challenge related to this aspect of the technique is preventing contamination, as these cultures (much like murine myofiber preps) are often difficult to keep sterile. Great care is needed if one is to attempt prolonged cultures. We also recommend the use of antimicrobials including antifungals.

In all, we demonstrate a practical method for the isolation of zebrafish myofibers as well as outline how to perform fluorescent immunolabeling and real time calcium imaging on the isolated fiber preparations. Continuing modification and development of this technique will offer an ever-expanding repertoire of tools to experiment on specific, isolated cell types in zebrafish. By dissociating zebrafish embryos into isolated individual myofibers, we can further enhance our understanding of muscle development, function and disease.

Disclosures

The authors declare no conflicting interests.