We thank Viewpoint for their kind sponsorship of this manuscript. This work was funded by an Australian National Health and Medical Research Council (NHMRC) Project Grant (APP1010110).

1. Touch-evoked Response Assay
<ol>
	<li>Preparation of 2 dpf Embryos for Touch-evoked Response Assay
	<ol>
		<li>Ensure that the time of day at which the test is conducted is consistent between experiments because activity can vary dramatically throughout the day17,18. 
		&#8203;NOTE: The experiment should be performed blinded and the order of testing randomized to minimize experimental artifacts.</li>
		<li>Assign the fish strains a number, which is unknown to the individual carrying out the experiment. Following this, using freely available online tools generate a random list that dictates the order of testing.</li>
		<li>At least one hour prior to testing, dechorionate embryos by ripping a hole in the chorion and gently pulling the chorion apart using a pair of fine tweezers. Remove any debris from the petri dish before returning to the 28 &#176;C incubator.</li>
	</ol>
	</li>
	<li>Performing Touch-evoked Response Assay
	<ol>
		<li>Heat stage to 28 &#176;C at least 15 min prior to starting the testing. 
		&#8203;NOTE: This stage is temperature controlled and will remain at 28&#160;&#176;C for the duration of the testing. Temperature will affect activity and it is therefore important to maintain a constant temperature. If a heated stage is not available then the temperature of the water should be monitored and all experiments should be conducted at the same temperature.</li>
		<li>Place a petri dish filled with embryo medium (5 mM NaCl, 0.17mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4 in water) onto an illuminated stage and mount the high-speed camera over the petri dish.</li>
		<li>Launch the video camera recording software (such as Stream Pix 5, described here) and under the &#34;workspace&#34; tab select 1,000 frame per sec (1,000 fps) as the capture speed to ensure that fast swimming action of the fish is recorded.</li>
		<li>Working with one embryo at a time, place the embryo in the middle of the petri dish with the zebrafish clearly visible in the field of view. 
		&#8203;NOTE: If the embryo swims away prior to commencement of the experiment replace it with another, as recapturing and positioning of the embryo may result in it becoming desensitized to the stimulus and repeated burst responses may promote muscle weakness in some disease models.</li>
		<li>Begin recording by clicking on the &#34;record&#34; button and, deliver the mechanosensory stimulus to the embryo by touching it gently with a blunt needle on the top of the head.</li>
		<li>Stop the recording after the embryo has swum out of the field of view or returned to rest. 
		&#8203;NOTE: The acceleration peaks within the first 0.2 sec of the burst escape response following the mechanical stimulus. Therefore, ensure that at least during the first 0.2 sec of the escape response recording the fish is in the field of view. Using the software described in step 1.2.3, the data will automatically be saved as a .avi video file. Alternative video capture software such as Free Video Capture or Softonic, both of which are freely available for download, could also be used.</li>
		<li>Return the embryo to a new petri dish and select another embryo for testing. Perform testing on a minimum of 15 fish.</li>
	</ol>
	</li>
	<li>Quantification of Swimming Behavior
	<ol>
		<li>To quantify the swimming behavior, launch the software and select the &#34;Single Larvae Locomotion without background subtraction&#34; module to open the saved .avi video file.</li>
		<li>Using the &#34;freehand&#34; or &#34;polygon&#34; tool from the menu bar select areas of the movie to be used for the analysis. Ensure that the region encompasses both the original position of the fish and the area that the fish will swim into. Make sure that the probe is excluded from the area to be analyzed. The software will automatically track the trajectory of the fish within the desired area.</li>
		<li>To perform the analysis, click on &#34;experiment&#34; from the menu bar and select &#34;execute&#34;. When prompted save the raw data analysis file (.phr format) in the desired location. Once saved, click on &#34;start&#34; to begin the analysis. End the analysis by clicking on &#34;stop&#34; under the &#34;experiment&#34; drop down menu. A window containing the results will be displayed.</li>
		<li>Scroll to the right to obtain the &#34;maximum acceleration&#34; value. If desired, export this data by closing the results window and clicking on the &#34;export instantaneous results&#34; button under the &#34;results&#34; drop down menu. Select the appropriate raw data analysis file and click on open. A text file that can be opened in a spreadsheet program will be saved in the destination folder.</li>
		<li>Repeat this process for each individual fish and average to obtain the mean maximum acceleration for each strain (see Figure 1). 
		NOTE: As an alternative to using the software described here, similar packages such as the freely available ImageJ software can be used to extract the relevant movement data. The 3D Particle Tracker plugin can be used to track swimming trajectories.</li>
	</ol>
	</li>
</ol>
2. Locomotion Assay &#8212; 10-min Swim Test
<ol>
	<li>Preparation of 6 dpf Embryos for Swimming Analysis
	<ol>
		<li>If required, sort embryos for the required genotype, for example by examining expression of a fluorescent protein or by phenotype, and place into a separate petri dish (25-30 embryos per dish). Alternatively, the genotype can be determined following completion of the locomotion assay.</li>
		<li>At 3 dpf, re-examine petri dishes and remove any unhatched embryos and debris. Return petri dishes to 28 &#176;C incubator until 6 dpf.</li>
		<li>Perform testing of all strains between 9 am and 12 pm, which is the time at which zebrafish larvae are most active. Randomize the order of testing and position in the plate of wildtype and mutant samples to minimize the effects of circadian differences and other experimental bias. 
		&#8203;NOTE: It is important that the testing time is consistent between experiments because activity can vary dramatically throughout the day.</li>
		<li>At least 30 min prior to testing, place larvae into a 48-well plate with one larva per well. After transfer, fill the wells so that the water surface is just below the top of the well, ensuring there are no bubbles. Return plates to 28 &#176;C incubator.</li>
		<li>Take plates out of the incubator and acclimatize in the light for five minutes prior to testing.</li>
	</ol>
	</li>
	<li>Performing Locomotion Assay
	<ol>
		<li>Place the 48-well plate into the recording chamber, which is equipped with an infrared digital camera, capturing up to 60 frames/sec, so that larvae can be detected in the dark. Check that all of the wells are placed inside the circular grid on the locomotion software and that all larvae are clearly detectable.</li>
		<li>Launch the software and select the &#8220;tracking&#8221; module. Under &#8220;file&#8221; click on &#8220;generate new protocol&#8221; and edit the number of wells used for the experiment. Set both the experiment duration and the integration period to 10 min by clicking on the &#34;parameters&#34; drop down menu and selecting the &#34;protocol parameters&#34; and subsequently the &#34;time&#34; tab. On the same &#34;protocol parameters&#34; dialogue box click on the &#34;options&#34; tab and ensure the &#34;Numeriscope&#34; checkbox is clicked following which, the &#34;protocol parameters&#34; dialogue box can be closed.</li>
		<li>To set the recording areas highlight the entire grid and double click on one of the wells. Click on the &#8220;draw areas&#8221; button and draw around the top left, top right, and bottom left wells and click &#8220;build&#8221;, which allows the software to automatically determine the position of each well. Also draw in a scale bar and click on &#8220;apply to group&#8221;. Once completed, click on the &#8220;draw areas&#8221; button.</li>
		<li>Visually determine the detection threshold by sliding the &#34;detection threshold&#34; bar to a level at which only the fish movements are highlighted with no background signals. 
		&#8203;NOTE: The detection threshold will vary between strains and thus the thresholds have to be determined whenever a new strain is tested. In the representative data presented a detection threshold of 25 mm/sec was used.</li>
		<li>Before commencing testing enter movement thresholds for detection of inactivity, and small and large movements. 
		NOTE: In the representative data presented an inactivity threshold of 6 mm/sec and an activity burst threshold of 30 mm/sec was used. The thresholds determine the minimum movement to be considered active and the level required to be considered burst activity and allow classification of activity into small (active but below burst activity thresholds) and large (greater than the burst activity threshold) movements. The thresholds can be altered depending on the activity of the particular fish strains analyzed. 
		&#8203;NOTE: Although the assay can be performed in either light or dark conditions, zebrafish larvae have been shown to be more active in the dark18.</li>
		<li>Set the light intensity inside the chamber to be at 0% by clicking on the &#34;light driving settings&#34; button under the &#34;parameters&#34; drop down menu. On the resulting dialogue box, add the required light settings. 
		&#8203;NOTE: The light intensity inside the chamber can be triggered to turn on and off during the testing time to stimulate larval activity</li>
		<li>Close the door of the recording chamber and start video recording.</li>
	</ol>
	</li>
	<li>Quantification of Swimming Behavior
	<ol>
		<li>After the experiment is complete, click on &#34;stop&#34; under the &#34;experiment&#34; drop down menu. A dialogue box with all the results will be displayed.</li>
		<li>To access these results in excel click on &#34;open containing folder&#34; and open the excel file that appears in the resulting folder. The important parameters are &#34;smlct&#34; (small movement count), &#34;larct&#34; (large movement count), &#34;smldist&#34; (total distance covered by the fish in small movements) and &#34;lardist&#34; (total distance covered by the fish in large movements). 
		NOTE: Following the recording, the software also returns two additional output files in the form of an .avi file (containing a video of the 10 min recording) and a .png image file (containing a visual representation of the locomotion during the 10-min experiment; see Figure 2).</li>
		<li>Once the locomotion values are calculated, replay the .avi and .png files to review whether the locomotion values calculated accurately depict the swimming movements of the fish (see Figure 3). 
		NOTE: As an alternative to using the software described here, packages such as the freely available ImageJ software can be used to track the locomotory behavior.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Many different animal models including mice, dogs, zebrafish, flies and worms have contributed towards our understanding of the genetic and molecular basis of muscle diseases, and assisted in the development of therapeutic approaches to combat them. The zebrafish boasts several advantages for the study of muscle disease. The zebrafish provides a genetically manipulable system to assess complex muscle patterning in a suitable physiological environment, which is not possible in in vitro culture systems. Unlike other vertebrate animal models, the large number of fish produced, together with its optical clarity, facilitates rapid, high-throughput in vivo chemical and genetic screening.
Here we describe the development of zebrafish movement assays to provide a high-throughput and automated method to assess muscle performance during zebrafish embryogenesis. For both assays it must be acknowledged that circadian rhythms and external environmental stimuli will significantly affect zebrafish swimming behavior17,18. Repeated testing of the same zebrafish will also lead to habituation causing a decrease in response to the tactile stimulus23. Therefore, in order to achieve reproducible results between experiments each zebrafish embryo should only be tested once, the time of the day and lighting conditions should be standardized, and water temperature needs to be tightly regulated.
Using the touch evoked analysis at 2 dpf we can directly measure the maximum acceleration of a burst swimming action, which is proportional to muscle force. Previous techniques in zebrafish have examined muscle force by tying both ends of the embryos to experimental equipment following which muscle contraction is stimulated using an electric field and the force-generating capability of the muscle14 is measured. Whilst this method measures the force generating capacity of the larval muscle, it does not measure the actual force generated by the larval muscle during swimming. We therefore developed a method to indirectly assess the force generated during the normal larval swimming motion to provide an overall measure of muscle health. The high-speed video system, capable of recording individual zebrafish movements at a frame rate of 1,000 frames/sec can be used to identify small but significant differences in muscle function, which are not directly distinguishable by eye. It will be of interest to see how previously reported changes in electrically stimulated force-generation correlate with changes in swimming performance.
In addition the touch evoked response assays can also be used to assess swimming kinematics, such as the shape and speed of the body wave during the swimming motion24, to give a quantitative measurement of the locomotory behavior.
Due to the spontaneous movement of zebrafish larvae after 3 dpf, we were not able to perform the touch-evoke assays to measure muscle function. Conversely, we measured muscle performance over a longer period by determining the distance swum by zebrafish larvae at 6 dpf. This test, although an indirect measure of muscle function, can be used to identify fish displaying impaired muscle performance8 or neurodegeneration25,26. This test not only provides a measurement analogous to the 6 min walk test but is also suitable for automated high-throughput in vivo drug or mutagenesis screens.

Over the past decade zebrafish have been increasingly used to study muscle cell biology and disease. The rapid external development of the zebrafish embryo, coupled with its optical clarity, allows the direct visualization of muscle formation, growth, and function. The process of muscle development is highly conserved in zebrafish and this has allowed the successful modeling of a range of muscle diseases including muscular dystrophies and congenital myopathies1-8. Detailed examination of zebrafish models has not only provided novel insights into the pathobiology of these conditions but also provided a platform for the testing of suitable therapies6,9-13.
The analysis of zebrafish models of muscle diseases relies on reliable and reproducible assays to measure muscle performance. Previous studies have successfully measured the force generating capability of the zebrafish trunk muscle in fish between 3 and 7 dpf by electrically stimulating contraction of an immobilized fish attached to a force transduction system14. This can provide detailed measurements of force but are not ideally suited to higher throughput experiments and there are advantages to measuring muscle performance during swimming. At 2 dpf zebrafish muscle is fully functional and the fish can elicit burst swimming movements in response to stimuli. The touch-evoke escape response assay is used to measure acceleration during a burst swimming motion, which can be used as a measure of contractile force.
One of the most utilized measures of muscle function in myopathy patients is the 6 min walk test, which records the total distance walked on a hard flat surface15,16. We have applied a comparable test to measure muscle function in 6 dpf zebrafish larvae, whereby we monitor the total distance swum, and the total number of movements made by each larva over a 10 min period. This is performed using an automated tracking system, which provides reliable and high-throughput measurements of muscle performance. Both muscle tests are highly reproducible and have been used to quantify differences in muscle performance in zebrafish myopathy models8.

<table><tbody><tr><td>21 G x 1&#039; Blunt Needle</td><td>Terumo/Admiral Medical Supplies</td><td>TE2125</td><td></td></tr><tr><td>48-well plates</td><td>Sigma</td><td>M8937</td><td></td></tr><tr><td>90 mm Petri Dishes</td><td>Pacific Laboratory Products PT</td><td>S90001</td><td></td></tr><tr><td>High Speed Camera</td><td>Baumer</td><td>HXC20</td><td></td></tr><tr><td>http://www.randomization.com</td><td>N/A</td><td></td><td>Steps 1.1.2, 2.1.3</td></tr><tr><td>Incubator</td><td>Thermoline Scientific</td><td>TEI-43L</td><td></td></tr><tr><td>Plastic Pipette</td><td>VWR</td><td>16001-188</td><td></td></tr><tr><td>StreamPix5</td><td>NorPix</td><td></td><td>Step 1.2.3</td></tr><tr><td>Temperature Control Unit</td><td>Viewpoint</td><td></td><td></td></tr><tr><td>Tweezers, style 8</td><td>ProSciTech</td><td>T04-821</td><td></td></tr><tr><td>Zebrabox System</td><td>Viewpoint</td><td></td><td></td></tr><tr><td>Zebralab</td><td>Viewpoint</td><td></td><td>Steps 1.3.1, 2.2.1</td></tr></tbody></table>

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.

Touch evoked response assay can be used to determine the speed and acceleration of swimming movements which is a proportional measure of muscle force. In response to a mechanical stimulus, such as a small tap on the head 2 dpf wild type zebrafish exhibit a fast swimming action. Videos were captured and analyzed for two different zebrafish myopathy models: Tg(ACTA1D286G-eGFP), a model of nemaline myopathy that has been shown to have significant muscle weakness, and a model of Duchenne muscular dystrophy in which severe muscle defects have been described at 5 dpf19,20. Images from a video of a typical touch evoked assay are represented in Figure 1A. Acceleration of the zebrafish was examined and found to peak within the first 0.2 sec of the burst swimming escape response (Figure 1B). This peak maximum acceleration provides a measure that is proportional to the force generating capacity of the skeletal muscle. The maximum acceleration values were averaged to obtain a mean maximum acceleration value (&#177; standard error of the mean) for each strain: Tg(ACTA1D286G-eGFP): mean = 276.0 &#177; 28.8 m/sec2, n = 3 independent replicate experiments comprising 15 individual fish; wildtype control: mean = 500.8 &#177; 50.28 m/sec2, n = 3 independent replicate experiments comprising 15 individual fish; dmdpc2-/- mutant: mean = 249.9 &#177; 19.1 m/sec2, n=3 independent replicate experiments comprising 12-19 individual fish; dmdpc2+/- heterozygotes: mean = 235.9 &#177; 8.7 m/sec2, n = 3 independent replicate experiments comprising 16-27 individual fish; dmdpc2+/+ wildtype homozygotes: mean = 230.9 &#177; 8.7 m/sec2, n = 3 independent replicate experiments comprising 8-27 individual fish (Figure 1C). As expected, the Tg(ACTA1D286G-eGFP) fish were found to have a significant decrease in maximum acceleration indicating reduced muscle function, which is consistent with mouse models and patient data8,21,22. The dmdpc2-/- mutant fish however, showed no difference in maximum acceleration, at 2 dpf, consistent with the detection of muscle defects from 3 dpf20 (Figure 1D).
Locomotion assays were performed at 6 dpf to determine the activity and distance swum by zebrafish strains as an indication of muscle performance. Following testing, a diagrammatic representation of the swimming movements over the ten-minute testing period was generated, with red and green lines representing periods of slow and fast movement respectively and black lines representing periods of inactivity (Figure 2). Individual wildtype zebrafish show high activity with relatively no periods of inactivity as opposed to Tg(ACTA1D286G-eGFP) fish, which are less active over the testing period (Figure 2B).
The swimming behavior was quantified by averaging the individual values of the number of movements and the distance swum by each fish (Figure 3). Both, Tg(ACTA1D286G-eGFP) fish (Figure 3A and 3B) and dmdpc2-/- mutant fish (Figure 3C and 3D) were found to have a significant decrease in the mean number of movements and distance swum compared to their respective controls: Tg(ACTA1D286G-eGFP) fish: mean number of movements = 94.3 &#177; 13.6, mean distance swum = 112.9 &#177; 18.4 mm, n = 3 independent replicate experiments comprising 45 fish; wild type controls: mean number of movements = 177.4 &#177; 14.0, mean distance swum = 300.2 &#177; 22.8 mm, n = 3 independent replicate experiments comprising 45 fish; dmdpc2-/- mutant: mean number of movements = 163.3 &#177; 30.0, mean distance swum: 298.4 &#177; 60.37 mm, n = 3 independent replicate experiments comprising 12-20 fish; dmdpc2+/- heterozygotes: mean number of movements = 362.3 &#177; 38.8, mean distance swum: 660.3 &#177; 86.1mm n = 3 independent replicate experiments comprising 17-27 fish; dmdpc2+/+ wildtype homozygotes: mean number of movements = 341.9 &#177; 91.6, mean distance swum = 574.3 &#177; 170.9mm n = 3 independent replicate experiments comprising 8-25 fish.
<img alt="Figure 1" src="/files/ftp_upload/54431/54431fig1.jpg" /> 
Figure 1: Quantification of touch-evoke response assay for 2 dpf zebrafish embryos. (A) Snapshot images of a control zebrafish during touch-evoke assays at 2 dpf. (B) Acceleration profile for the first 0.2 sec of a single Tg(ACTA1D286G-eGFP) (red) and single control (blue) zebrafish following application of the touch stimulus. The maximum acceleration is represented by the dotted lines. (C, D) Quantification of the maximum acceleration (m/sec2) recorded from touch-evoked response assays of (C) Tg(ACTA1D286G-eGFP) zebrafish and (D) dmdpc2-/- mutant fish compared to control zebrafish at 2 dpf. Error bars represent &#177; SEM for 3 replicate experiments, *p &#60;0.05. <a href="http://cloudflare.jove.com/files/ftp_upload/54431/54431fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54431/54431fig2.jpg" /> 
Figure 2: Representation of locomotion assays for zebrafish embryos. (A) Zebrafish embryos are placed in 48-well plates and locomotion is recorded from above using an infrared digital camera. (B) Schematic of zebrafish movement during the testing period with red lines depicting fast movements, green lines depicting slow movements and black lines depicting inactivity (as determined by the detection thresholds entered in the software). <a href="http://cloudflare.jove.com/files/ftp_upload/54431/54431fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54431/54431fig3.jpg" /> 
Figure 3: Quantification of locomotion assays for 6 dpf zebrafish larvae. Quantification of the (A) number of movements and (B) distance travelled by Tg(ACTA1D286G-eGFP) zebrafish compared to control zebrafish at 6 dpf. Quantification of the (C) number of movements and (D) distance travelled by dmdpc2-/- mutant fish compared to control zebrafish at 6 dpf. Error bars represent &#177; SEM for 3 replicate experiments, *p &#60;0.05, **p &#60;0.01. <a href="http://cloudflare.jove.com/files/ftp_upload/54431/54431fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Using Touch-evoked Response and Locomotion Assays to Assess Muscle Performance and Function in Zebrafish at JoVE.com

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

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 ...

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JoVE Journal

Developmental Biology

12.7K Views.  Monash University. 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. 

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

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 ...

A Behavioral Assay to Measure Responsiveness of Zebrafish to Changes in Light Intensities 

The optokinetic reflex (OKR) is a basic visual reflex exhibited by most vertebrates and plays an important role in stabilizing the eye relative to the visual scene. However, the OKR requires that an animal detect moving stripes and it is possible that fish that fail to exhibit an OKR may not be completely blind. One zebrafish mutant, the no optokinetic response c (nrc) has no OKR under any light conditions tested and was reported to be completely blind. Previously, we have shown that OFF-ganglion cell activity can be recorded in these mutants. To determine whether mutant fish with no OKR such as the nrc mutant can detect simple light increments and decrements we developed the visual motor behavioral assay (VMR). In this assay, single zebrafish larvae are placed in each well of a 96-well plate allowing the simultaneous monitoring of larvae using an automated video-tracking system. The locomotor responses of each larva to 30 minutes light ON and 30 minutes light OFF were recorded and quantified. WT fish have a brief spike of motor activity upon lights ON, known as the startle response, followed by return to lower-than baseline activity, called a freeze. WT fish also sharply increase their locomotor activity immediately following lights OFF and only gradually (over several minutes) return to baseline locomotor activity. The nrc mutants respond similarly to light OFF as WT fish, but exhibit a slight reduction in their average activity as compared to WT fish. Motor activity in response to light ON in nrc mutants is delayed and sluggish. There is a slow rise time of the nrc mutant response to light ON as compared to WT light ON response. The results indicate that nrc fish are not completely blind. Because teleosts can detect light through non-retinal tissues, we confirmed that the immediate behavioral responses to light-intensity changes require intact eyes by using the chokh (chk) mutants, which completely lack eyes from the earliest stages of development. In our VMR assay, the chk mutants exhibit no startle response to either light ON or OFF, showing that the lateral eyes mediate this behavior. The VMR assay described here complements the well-established OKR assay, which does not test the ability of zebrafish larvae to respond to changes in light intensities. Additionally, the automation of the VMR assay lends itself to high-throughput screening for defects in light-intensity driven visual responses.

A Behavioral Assay to Measure Responsiveness of Zebrafish to Changes in Light Intensities

We developed the Visual-Motor Response to quantitate the motor output of larval zebrafish in response to light increments and decrements. We also examined zebrafish vision mutants, including the no optokinetic response (nrc) mutants, which were thought to be completely blind when tested by another vision assay, the optokinetic reflex.

The optokinetic reflex (OKR) is a basic visual reflex exhibited by most vertebrates and plays an important role in stabilizing the eye relative to the ...

Biology

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.

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, ...

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 ...

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 ...

Force Measurement During Contraction to Assess Muscle Function in Zebrafish Larvae

Zebrafish larvae provide models of muscle development, muscle disease and muscle-related chemical toxicity, but related studies often lack functional measures of muscle health. In this video article, we demonstrate a method to measure force generation during contraction of zebrafish larval trunk muscle. Force measurements are accomplished by placing an anesthetized larva into a chamber filled with a salt solution. The anterior end of the larva is tied to a force transducer and the posterior end of the larva is tied to a length controller. An isometric twitch contraction is elicited by electric field stimulation and the force response is recorded for analysis. Force generation during contraction provides a measure of overall muscle health and specifically provides a measure of muscle function. Although we describe this technique for use with wild-type larvae, this method can be used with genetically modified larvae or with larvae treated with drugs or toxicants, to characterize muscle disease models and evaluate treatments, or to study muscle development, injury, or chemical toxicity.

Force measurements can be used to demonstrate changes in muscle function due to development, injury, disease, treatment or chemical toxicity. In this video, we demonstrate a method to measure force during a maximal contraction of zebrafish larval trunk muscle.

Zebrafish larvae provide models of muscle development, muscle disease and muscle-related chemical toxicity, but related studies often lack functional ...

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 ...

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. ...

Neuroscience

Startle Assay for Toxicity Evaluation in Zebrafish Embryo

Evaluating Toxicity of Chemicals using a Zebrafish Vibration Startle Response Screening System

We developed a simple screening system for the evaluation of neuromuscular and general toxicity in zebrafish embryos. The modular system consists of electrodynamic transducers above which tissue culture dishes with embryos can be placed. Multiple such loudspeaker-tissue culture dish pairs can be combined. Vibrational stimuli generated by the electrodynamic transducers induce a characteristic startle and escape response in the embryos. A belt-driven linear drive sequentially positions a camera above each loudspeaker to record the movement of the embryos. In this way, alterations to the startle response due to lethality or neuromuscular toxicity of chemical compounds can be visualized and quantified. We present an example of the workflow for chemical compound screening using this system, including the preparation of embryos and treatment solutions, operation of the recording system, and data analysis to calculate benchmark concentration values of compounds active in the assay. The modular assembly based on commercially available simple components makes this system both economical and flexibly adaptable to the needs of particular laboratory setups and screening purposes.

Author Spotlight: High-Throughput Toxicity Screening Using Zebrafish Embryo Startle Response Assay

We describe a screening system's workflow and data analysis for evaluating chemical compound toxicity based on the zebrafish embryo vibration startle response. The system records the movements of zebrafish embryos upon exposure to a vibration stimulus and allows for an integrated evaluation of general toxicity/lethality and neuromuscular toxicity.

We developed a simple screening system for the evaluation of neuromuscular and general toxicity in zebrafish embryos. The modular system consists of ...

Escape Response Assay: A Method to Study the Response of Zebrafish Larva to Touch Stimuli

Source: Sztal, T. E., et al. <a href="https://jove.unicartagenaproxy.elogim.com/v/54431/" target="_blank">Using Touch-evoked Response and Locomotion Assays to Assess Muscle Performance and Function in Zebrafish</a>.&#160;J. Vis. Exp.&#160;(2016)
This video describes the touch-evoke response assay, which measures the response of the zebrafish larva to touch stimuli.

Test reakcji ucieczki: metoda badania reakcji larw danio pręgowanego na bodźce dotykowe

Source: Sztal, T. E., et al. Using Touch-evoked Response and Locomotion Assays to Assess Muscle Performance and Function in Zebrafish.&#160;J. Vis. ...

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

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Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

A Behavioral Assay to Measure Responsiveness of Zebrafish to Changes in Light Intensities

Examining Muscle Regeneration in Zebrafish Models of Muscle Disease

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

Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

Force Measurement During Contraction to Assess Muscle Function in Zebrafish Larvae

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

Assessment of Swim Endurance and Swim Behavior in Adult Zebrafish

Evaluating Toxicity of Chemicals using a Zebrafish Vibration Startle Response Screening System

Test reakcji ucieczki: metoda badania reakcji larw danio pręgowanego na bodźce dotykowe