The authors wish to thank the members of the Dowling lab (Aaron Reifler, Trent Waugh, Angela Busta, and William Telfer) that contributed to the development of the technique and to the production of the manuscript. This work was funded by the Taubman Institute, the Department of Pediatrics at the University of Michigan, and in part from grants from the Muscular Dystrophy Association (JJD MDA186999) and the National Institutes of Health (JJD 1K08AR054835).

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).
<ol>
	<li>Cut and place Parafilm on the bottom of a 60 mm Petri dish (any brand).</li>
	<li>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.</li>
</ol>
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.
<ol start="3">
	<li>Pipette 50-200 &#956;l of poly-L-lysine solution (0.01%) on each cover glass in the Petri dish.</li>
	<li>Allow the coverslips to sit at least one hour at 37 &#176;C. Longer incubations will not negatively influence results.</li>
	<li>Remove poly-L-lysine solution from the cover glass and wash twice with a minimum of 100 &#956;l 1x PBS.</li>
	<li>Allow coverslips to dry.</li>
	<li>Coverslips are now ready for myofibers.</li>
</ol>
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 &#176;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.
<ol>
	<li>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.
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>Add 900 &#956;l of CO2&#160;independent media to the tube containing the embryos.</li>
	<li>Add 100 &#956;l 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.</li>
	<li>Rotate embryos on an orbital shaker, and triturate every 30 min at RT using a P1000 Pipetman for the trituration.</li>
</ol>
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.
<ol start="5">
	<li>Stop digestion when no whole embryos are visible, yet solid pieces are still visible -&#160;this is essential to prevent overdigestion.</li>
	<li>Centrifuge tubes with dissociated embryos at 0.8-2.3 x g for 3-5 min to pellet cells.</li>
	<li>Remove supernatant from pelleted cells and wash 2x with CO2&#160;independent media.</li>
	<li>Add fresh CO2&#160;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.</li>
	<li>Using a P1000 pipette, pass embryo suspension through a 70 &#956;m filter . This helps remove unwanted debris from the prep.</li>
</ol>
Note 4: Embryo suspension can be filtered a second time through a 40 &#956;m filter to further remove unwanted debris. From a double filtration, recovery is approximately 800 &#956;l from a starting volume of 1 ml.
<ol start="10">
	<li>Add approximately 50-100 &#956;l of myofiber suspension to each poly-L-lysine coated coverslip.</li>
</ol>
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.
<ol start="11">
	<li>Allow myofibers to settle approximately 1 hr onto the coated coverslips at room temperature.</li>
</ol>
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.
<ol start="12">
	<li>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.</li>
</ol>
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. Fluorescence Immunostaining of Dissociated Zebrafish Myofibers (Time: approximately 1 day)
3.1.&#160;Immunolabeling
<ol>
	<li>Remove a portion of media from myofibers adhered to the coated coverslip</li>
	<li>Fix cells using 4% paraformaldehyde or methanol. For PFA, remove &#189; 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 &#956;l of PFA, and fix for an additional 10 min. For alcohol fixation, ice cold methanol or acetone can be applied at 4 &#176;C for 10 min or 5 min, respectively.</li>
	<li>Wash myofibers at least 3-5x with 1x PBS or 1x PBS plus 0.1% TWEEN. Average wash volume is 50-100 &#956;l per coverslip.</li>
	<li>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.</li>
	<li>Remove blocking solution and add primary antibody diluted in either blocking solution or PBS. Incubate myofibers overnight at 4 &#176;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.</li>
	<li>Remove primary antibody from coverslips and wash coverslips 3-5x for 5 min with PBS or blocking solution.</li>
	<li>Add secondary antibody at appropriate concentration diluted in 1x PBS or blocking solution for 1-2 hr at RT.</li>
	<li>Remove secondary antibody and wash myofibers 3-5x in PBS for 5 min each at RT.</li>
</ol>
3.2.&#160;Mounting coverslips
<ol start="9">
	<li>Apply 1-2 drops of antifade reagent to a microscope slide.</li>
	<li>Carefully pick-up the coated and immunolabled coverslip using forceps and place (myofibers down) the coverslip on the antifade reagent on the microscope slide.</li>
</ol>
Note 1: Attention to the orientation of the coated coverslip is critical.
<ol start="11">
	<li>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.</li>
	<li>Apply light pressure to the corners of the microscope slide to remove excess antifade and form a tight seal between the slide and coverslip</li>
	<li>Allow the antifade remaining to dry for 5-10 min at RT, then the myofibers are ready to image (Figures 2A and&#160;B) or store at 4 &#176;C.</li>
</ol>
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 &#945;-actin promoter (pSKM)23. Alternatively, this technique can be readily adapted to use calcium indicator dyes such as Fura-2.
<ol>
	<li>Inject embryos with pSKM: GCaMP3 construct at the 1-2 cell stage</li>
	<li>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.</li>
	<li>Carefully remove any excess media, and add 300 &#956;l of fresh CO2&#160;independent media at RT.</li>
	<li>Observe cells under an inverted microscope. GCaMP3 positive myofibers should be visible under green fluorescence.</li>
	<li>Set up camera for recording, if desired.</li>
	<li>Prepare a 30 mM caffeine solution in CO2&#160;independent media. To stimulate cells, gently pipette 300 &#956;l 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&#160;and ryanodine4, can be used to study inducible calcium release.</li>
</ol>

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E., 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=Novel+deletion+of+lysine+7+expands+the+clinical,+histopathological+and+genetic+spectrum+of+TPM2-related+myopathies.">Novel deletion of lysine 7 expands the clinical, histopathological and genetic spectrum of TPM2-related myopathies.</a> Brain. 136, 508-521 (2013).</li><li>Schredelseker, J., Shrivastav, M., Dayal, A., Grabner, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Non-Ca2+-conducting+Ca2++channels+in+fish+skeletal+muscle+excitation-contraction+coupling.">Non-Ca2+-conducting Ca2+ channels in fish skeletal muscle excitation-contraction coupling.</a> Proc. Natl. Acad. Sci. U.S.A. 107, 5658-5663 (2010).</li></ol>

The authors declare no conflicting interests.

Zebrafish are a powerful vertebrate model system for studying muscle development and function&#160;in vivo25,27,28&#160;. They have also emerged as a valuable asset for modeling human muscle diseases14,15,20,29&#160;. 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, behavio...

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 stores1,2 . 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 coupling4,5 , and actin-myosin dependent contractions6 continue to be the ongoing subject of intense research. In addition, proteins that stabilize the muscle membrane during contraction7,8 and that mediate signaling between the myofiber and the extracellular matrix7,9 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 (<a href="http://www.musclegenetable.org/" target="_blank">http://www.musclegenetable.org/</a>). 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 mortality10,11 . The zebrafish has proven to be an outstanding system for modeling and studying human skeletal muscle diseases8,12,13 . It has been employed to validate new gene mutations8, define new aspects of disease pathophysiology14,15 , and identify new therapeutic approaches15,16 . 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 others12,18,19 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.

<table border="1">
	<tbody>
		<tr>
			<td>Name of the reagent</td>
			<td>Company</td>
			<td>Catalogue number</td>
			<td>Comments (optional)</td>
		</tr>
		<tr>
			<td>24-well culture plate</td>
			<td>Corning</td>
			<td>3524</td>
			<td></td>
		</tr>
		<tr>
			<td>10x PBS</td>
			<td>Invitrogen Gibco</td>
			<td>70011</td>
			<td></td>
		</tr>
		<tr>
			<td>CO2 Independent medium</td>
			<td>Invitrogen Gibco</td>
			<td>18045</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase Type II</td>
			<td>Worthington Biochemical</td>
			<td>LS004186</td>
			<td>Lot 41H12764</td>
		</tr>
		<tr>
			<td>Collagenase Type IV</td>
			<td>Worthington Biochemical</td>
			<td>L5004188</td>
			<td></td>
		</tr>
		<tr>
			<td>8% Paraformaldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>157-8</td>
			<td></td>
		</tr>
		<tr>
			<td>Methanol</td>
			<td>Sigma</td>
			<td>322415</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma</td>
			<td>X100</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Sigma</td>
			<td>A2153</td>
			<td></td>
		</tr>
		<tr>
			<td>Sheep serum</td>
			<td>Sigma</td>
			<td>S3772</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat serum</td>
			<td>Sigma</td>
			<td>G9023</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass coverslips</td>
			<td>Fischerbrand</td>
			<td>12-545-82 12CIR-1D</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly-L-Lysine</td>
			<td>Sigma</td>
			<td>P4707</td>
			<td></td>
		</tr>
		<tr>
			<td>Pronase</td>
			<td>Sigma</td>
			<td>P5147</td>
			<td></td>
		</tr>
		<tr>
			<td>40 &#956;m Filter</td>
			<td>BD Biosciences</td>
			<td>352340</td>
			<td></td>
		</tr>
		<tr>
			<td>70 &#956;m Filter</td>
			<td>BD Biosciences</td>
			<td>352350</td>
			<td></td>
		</tr>
		<tr>
			<td>Prolong Gold antifade reagent</td>
			<td>Invitrogen</td>
			<td>P36931</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-&#945;-Actinin antibody</td>
			<td>Sigma</td>
			<td>A5044</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-RYR antibody</td>
			<td>Abcam</td>
			<td>34C</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor antibody</td>
			<td>Invitrogen</td>
			<td>A-21425</td>
			<td></td>
		</tr>
		<tr>
			<td>TWEEN 20</td>
			<td>Sigma</td>
			<td>P1379</td>
			<td></td>
		</tr>
		<tr>
			<td>60 mm Petri dish</td>
			<td>Fischerbrand</td>
			<td>0875713A</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly-L-Ornithine</td>
			<td>Sigma</td>
			<td>P4957</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope slide</td>
			<td>Fischerbrand</td>
			<td>12-550-15</td>
			<td></td>
		</tr>
		<tr>
			<td>Caffeine</td>
			<td>Sigma</td>
			<td>C0750</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

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-&#945;-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...

Watch this Scientific Journal Video about Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations at JoVE.com

Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

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 ...

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11.5K vues.  University of Michigan.  Poisson zèbre sont d'un nouveau système pour la modélisation de maladies humaines du muscle squelettique. Nous décrivons une méthode rapide et efficace pour isoler les fibres musculaires des muscles squelettiques de poisson zèbre embryonnaire et larvaire. Cette méthode produit une préparation de haute densité myofiber approprié pour l'étude de la morphologie unique de fibres de muscle squelettique, la localisation subcellulaire des protéines, et la ph...

Vidéo: Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

Primary Dissociated Midbrain Dopamine Cell Cultures from Rodent Neonates

The ability to create primary cell cultures of dopamine neurons allows for the study of the presynaptic characteristics of dopamine neurons in isolation from systemic input from elsewhere in the brain. In our lab, we use these neurons to assess dopamine release kinetics using carbon fiber amperometry, as well as expression levels of dopamine related genes and proteins using quantitative PCR and immunocytochemistry. In this video, we show you how we generate these cultures from rodent neonates.
The process involves several steps, including the plating of cortical glial astrocytes, the conditioning of neuronal cell culture media by the glial substrate, the dissection of the midbrain in neonates, the digestion, extraction and plating of dopamine neurons and the addition of neurotrophic factors to ensure cell survival.
The applications suitable for such a preparation include electrophysiology, immunocytochemistry, quantitative PCR, video microscopy (i.e., of real-time vesicular fusion with the plasma membrane), cell viability assays and other toxicological screens.

Primary dissociated midbrain dopamine cell cultures allow for the study of presynaptic characteristics of dopamine neurons. They can be used to monitor real-time dopamine release kinetics and protein/mRNA levels of regulators of dopamine exocytosis. Here, we show you how to generate these cultures from rodent neonates.

The ability to create primary cell cultures of dopamine neurons allows for the study of the presynaptic characteristics of dopamine neurons in isolation ...

RNA Isolation from Embryonic Zebrafish and cDNA Synthesis for Gene Expression Analysis

Many important and complex laboratory procedures require an input of high quality, intact RNA.  A degraded sample or the presence of impurities can lead to disastrous results in downstream experimental applications.  It is therefore, of utmost importance to use solid techniques with numerous safeguards and quality control checks to ensure a superior sample.  Herein, we detail a protocol to isolate total RNA from whole zebrafish embryos using a commercially available chemical denaturant and subsequent cleanup to remove traces of DNA and impurities using a commercial RNA isolation kit. As RNA is relatively unstable and easily prone to cleavage by RNAses, most protocols assay gene expression using a cDNA product that is directly synthesized from an RNA template. We detail a procedure to convert RNA into the more stable cDNA product using a commercially available kit. Throughout these procedures there are numerous quality control checks to ensure that the sample is not degraded or contaminated.  The end product of these protocols is cDNA that is suitable for microarray analysis, RT-PCR or long-term storage.    

The isolation of high quality, intact RNA is an essential step in many laboratory protocols. Here, we demonstrate RNA extraction from whole zebrafish embryos and cDNA synthesis for subsequent application in various experimental procedures including gene expression microarray analysis.

Many important and complex laboratory procedures require an input of high quality, intact RNA.  A degraded sample or the presence of impurities can lead ...

Microdissection of Zebrafish Embryonic Eye Tissues

Zebrafish is a popular animal model for research on eye development because of its rapid ex utero development and good fecundity. By 3 days post fertilization (dpf), the larvae will show the first visual response. Many genes have been identified to control a proper eye development, but we are far from a complete understanding of the underlying genetic architecture. Whole genome gene expression profiling is a useful tool to elucidate genetic regulatory network for eye development. However, the small size of the embryonic eye in zebrafish makes it challenging to obtain intact and pure eye tissues for expression analysis. For example, the anterior-posterior length of the eye between day 2 and 3 is only approximately 200-300 &mu;m, while the diameter of the lens is less 100 &mu;m. Also, the retinal pigment epithelium (RPE) underlying the retina is just a single-layer epithelium. While gene expression profiles can be obtained from the whole embryo, they do not accurately represent the expression of these tissues. Therefore pure tissue must be obtained for a successful gene expression profiling of eye development. To address this issue, we have developed an approach to microdissect intact retina and retina with RPE attached from 1-3 dpf, which cover major stages of eye morphogenesis. All procedures can be done with fine forceps and general laboratory supplies under standard stereomicroscopes. For retinal dissection, the single-layer RPE is removed and peeled off by brushing action and the preferential adherence of the RPE remnants to the surface of the culture plate for dissection. For RPE-attached retinal dissection, the adherence of RPE to the dissection plate is removed before the dissection so that the RPE can be completely preserved with the retina. A careful lifting action of this tissue can efficiently separate the presumptive choroid and sclera. The lens can be removed in both cases by a chemically etched tungsten needle. In short, our approach can obtain intact eye tissues and has been successfully utilized to study tissue-specific expression profiles of zebrafish retina1, 2 and retinal pigment epithelium3.

This article describes an approach to microdissect zebrafish retinas with and without retinal pigment epithelium attached, from one to three days postfertilization embryos.

Zebrafish is a popular animal model for research on eye development because of its rapid ex utero development and good fecundity. By 3 days post ...

Adult and Embryonic Skeletal Muscle Microexplant Culture and Isolation of Skeletal Muscle Stem Cells

Cultured embryonic and adult skeletal muscle cells have a number of different uses. The micro-dissected explants technique described in this chapter is a robust and reliable method for isolating relatively large numbers of proliferative skeletal muscle cells from juvenile, adult or embryonic muscles as a source of skeletal muscle stem cells. The authors have used micro-dissected explant cultures to analyse the growth characteristics of skeletal muscle cells in wild-type and dystrophic muscles. Each of the components of tissue growth, namely cell survival, proliferation, senescence and differentiation can be analysed separately using the methods described here. The net effect of all components of growth can be established by means of measuring explant outgrowth rates. The micro-explant method can be used to establish primary cultures from a wide range of different muscle types and ages and, as described here, has been adapted by the authors to enable the isolation of embryonic skeletal muscle precursors.
Uniquely, micro-explant cultures have been used to derive clonal (single cell origin) skeletal muscle stem cell (SMSc) lines which can be expanded and used for in vivo transplantation. In vivo transplanted SMSc behave as functional, tissue-specific, satellite cells which contribute to skeletal muscle fibre regeneration but which are also retained (in the satellite cell niche) as a small pool of undifferentiated stem cells which can be re-isolated into culture using the micro-explant method.

The micro-dissected explants technique is a robust and reliable method for isolating proliferative skeletal muscle cells from juvenile, adult or embryonic muscles as a source of skeletal muscle stem cells. Uniquely, these cells have been clonally derived to produce skeletal muscle stem cell lines used for in vivo transplantation.

Cultured embryonic and adult skeletal muscle cells have a number of different uses. The micro-dissected explants technique described in this chapter is a ...

Heart Dissection in Larval, Juvenile and Adult Zebrafish, Danio rerio

Zebrafish have become a beneficial and practical model organism for the study of embryonic heart development (see recent reviews1-6), however, work examining post-embryonic through adult cardiac development has been limited7-10. Examining the changing morphology of the maturing and aging heart are restricted by the lack of techniques available for staging and isolating juvenile and adult hearts. In order to analyze heart development over the fish's lifespan, we dissect zebrafish hearts at numerous stages and photograph them for further analysis11. The morphological features of the heart can easily be quantified and individual hearts can be further analyzed by a host of standard methods. Zebrafish grow at variable rates and maturation correlates better with fish size than age, thus, post-fixation, we photograph and measure fish length as a gauge of fish maturation. This protocol explains two distinct, size dependent dissection techniques for zebrafish, ranging from larvae 3.5mm standard length (SL) with hearts of 100&mu;m ventricle length (VL), to adults, with SL of 30mm and VL 1mm or larger. Larval and adult fish have quite distinct body and organ morphology. Larvae are not only significantly smaller, they have less pigment and each organ is visually very difficult to identify. For this reason, we use distinct dissection techniques.
We used pre-dissection fixation procedures, as we discovered that hearts dissected directly after euthanization have a more variable morphology, with very loose and balloon like atria compared with hearts removed following fixation. The fish fixed prior to dissection, retain in vivo morphology and chamber position (data not shown). In addition, for demonstration purposes, we take advantage of the heart (myocardial) specific GFP transgenic Tg(myl7:GFP)twu34 (12), which allows us to visualize the entire heart and is particularly useful at early stages in development when the cardiac morphology is less distinct from surrounding tissues. Dissection of the heart makes further analysis of the cell and molecular biology underlying heart development and maturation using in situ hybridization, immunohistochemistry, RNA extraction or other analytical methods easier in post-embryonic zebrafish. This protocol will provide a valuable technique for the study of cardiac development maturation and aging.

Heart Dissection in Larval, Juvenile and Adult Zebrafish, Danio rerio

A clear, standardized method for dissection and isolation of the zebrafish heart at multiple developmental stages are described. Annotation and quantification techniques are also discussed.

Zebrafish have become a beneficial and practical model organism for the study of embryonic heart development (see recent reviews1-6), however, work ...

Immunostaining of Dissected Zebrafish Embryonic Heart

Zebrafish embryo becomes a popular in vivo vertebrate model for studying cardiac development and human heart diseases due to its advantageous embryology and genetics 1,2. About 100-200 embryos are readily available every week from a single pair of adult fish. The transparent embryos that develop ex utero make them ideal for assessing cardiac defects 3. The expression of any gene can be manipulated via morpholino technology or RNA injection 4. Moreover, forward genetic screens have already generated a list of mutants that affect different perspectives of cardiogenesis 5. 
Whole mount immunostaining is an important technique in this animal model to reveal the expression pattern of the targeted protein to a particular tissue 6. However, high resolution images that can reveal cellular or subcellular structures have been difficult, mainly due to the physical location of the heart and the poor penetration of the antibodies. 
 Here, we present a method to address these bottlenecks by dissecting heart first and then conducting the staining process on the surface of a microscope slide. To prevent the loss of small heart samples and to facilitate solution handling, we restricted the heart samples within a circle on the surface of the microscope slides drawn by an immEdge pen. After the staining, the fluorescence signals can be directly observed by a compound microscope. 
Our new method significantly improves the penetration for antibodies, since a heart from an embryonic fish only consists of few cell layers. High quality images from intact hearts can be obtained within a much reduced procession time for zebrafish embryos aged from day 2 to day 6. Our method can be potentially extended to stain other organs dissected from either zebrafish or other small animals.

A rapid way to conduct immunostaining of zebrafish embryonic heart is described. Compared to the whole mount immunostaining approach, this method dramatically increases the penetration of the antibodies, which allows obtaining high resolution images that reveal cellular/subcellular structures in the heart within a much reduced processing time.

Zebrafish embryo becomes a popular in vivo vertebrate model for studying cardiac development and human heart diseases due to its advantageous embryology ...

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 ...

Isolation and Culture of Individual Myofibers and their Satellite Cells from Adult Skeletal Muscle

Muscle regeneration in the adult is performed by resident stem cells called satellite cells. Satellite cells are defined by their position between the basal lamina and the sarcolemma of each myofiber. Current knowledge of their behavior heavily relies on the use of the single myofiber isolation protocol. In 1985, Bischoff described a protocol to isolate single live fibers from the Flexor Digitorum Brevis (FDB) of adult rats with the goal to create an in vitro system in which the physical association between the myofiber and its stem cells is preserved 1. In 1995, Rosenblattmodified the Bischoff protocol such that myofibers are singly picked and handled separately after collagenase digestion instead of being isolated by gravity sedimentation 2, 3. The Rosenblatt or Bischoff protocol has since been adapted to different muscles, age or conditions 3-6. The single myofiber isolation technique is an indispensable tool due its unique advantages. First, in the single myofiber protocol, satellite cells are maintained beneath the basal lamina. This is a unique feature of the protocol as other techniques such as Fluorescence Activated Cell Sorting require chemical and mechanical tissue dissociation 7. Although the myofiber culture system cannot substitute for in vivo studies, it does offer an excellent platform to address relevant biological properties of muscle stem cells. Single myofibers can be cultured in standard plating conditions or in floating conditions. Satellite cells on floating myofibers are subjected to virtually no other influence than the myofiber environment. Substrate stiffness and coating have been shown to influence satellite cells' ability to regenerate muscles 8, 9 so being able to control each of these factors independently allows discrimination between niche-dependent and -independent responses. Different concentrations of serum have also been shown to have an effect on the transition from quiescence to activation. To preserve the quiescence state of its associated satellite cells, fibers should be kept in low serum medium 1-3. This is particularly useful when studying genes involved in the quiescence state. In serum rich medium, satellite cells quickly activate, proliferate, migrate and differentiate, thus mimicking the in vivo regenerative process 1-3. The system can be used to perform a variety of assays such as the testing of chemical inhibitors; ectopic expression of genes by virus delivery; oligonucleotide based gene knock-down or live imaging. This video article describes the protocol currently used in our laboratory to isolate single myofibers from the Extensor Digitorum Longus (EDL) muscle of adult mice (6-8 weeks old).

Isolation and culture of myofibers is the gold standard in vitro system to study the transition of satellite cells through quiescence, activation and differentiation. Importantly, the single myofiber culture system preserves the myofiber/stem cell association, which is an essential component of the muscle stem cell niche.

Muscle  regeneration in the adult is performed by resident stem cells called satellite  cells. Satellite cells are defined by  their position between the ...

Imaging and 3D Reconstruction of Cerebrovascular Structures in Embryonic Zebrafish

Zebrafish are a powerful tool to study developmental biology and pathology in vivo. The small size and relative transparency of zebrafish embryos make them particularly useful for the visual examination of processes such as heart and vascular development. In several recent studies transgenic zebrafish that express EGFP in vascular endothelial cells were used to image and analyze complex vascular networks in the brain and retina, using confocal microscopy. Descriptions are provided to prepare, treat and image zebrafish embryos that express enhanced green fluorescent protein (EGFP), and then generate comprehensive 3D renderings of the cerebrovascular system. Protocols include the treatment of embryos, confocal imaging, and fixation protocols that preserve EGFP fluorescence. Further, useful tips on obtaining high-quality images of cerebrovascular structures, such as removal the eye without damaging nearby neural tissue are provided. Potential pitfalls with confocal imaging are discussed, along with the steps necessary to generate 3D reconstructions from confocal image stacks using freely available open source software.

Imaging of cerebrovascular development in larval zebrafish is described. Techniques to facilitate 3D imaging and modify cerebrovascular development using chemical treatments are also provided.

Zebrafish are a powerful tool to study developmental biology and pathology in vivo.  The small size and relative transparency of zebrafish embryos make ...

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.