The authors thank Angela Busta for assistance with zebrafish husbandry. This work was supported by the National Institutes of Health (AG-020591 to S.V.B. and 1K08AR054835 to J.J.D.).

Note: all procedures involving zebrafish should be performed in accordance with relevant guidelines, regulations, and regulatory agencies. All animal use procedures shown in this article were approved by the University of Michigan Committee on the Use and Care of Animals (UCUCA).
1. Make Suture Loops
<ol>
	<li>Use forceps to separate non-sterile suture (USP 10/0 monofilament nylon, 3 ply) into three strands.</li>
	<li>Begin to tie a double overhand knot in one of the strands. Stop before tightening the knot completely to make a small (~1 mm diameter) loop instead of a knot.</li>
	<li>Use scissors to cut excess suture from the loop tails. An example of a finished loop is shown in Figure 1.</li>
	<li>Place the loop on the sticky side of a Post-it Note for later use. The suture loops will be used to hold the larvae in place during force testing.</li>
	<li>Repeat steps 1.1-1.4 as necessary. Make two suture loops for each larva that will be tested.</li>
</ol>
2. Make Testing Solution
<ol>
	<li>Make Tyrodes solution by adding 7.977 g sodium chloride, 0.373 g potassium chloride, 0.265 g calcium chloride dihydrate, 0.102 g magnesium chloride hexahydrate, 0.048 g sodium phosphate monobasic, 1.000 g sodium bicarbonate, and 0.037 g ethylenediaminetetraacetic acid disodium salt dihydrate to 1,000 ml of purified water.</li>
	<li>Stir the solution until the salts are completely dissolved. This solution can be stored for a month at 4 &deg;C.</li>
	<li>Add 2.1 ml of 4 mg/ml tricaine, prepared according to The Zebrafish Book24, to 47.9 ml Tyrodes solution and mix. Protect this solution from light by storing it in a dark glass bottle or in a glass bottle covered with aluminum foil. This solution should be stored at room temperature and made fresh each day.</li>
</ol>
3. Tie Aanesthetized Larva into Experimental Chamber
<ol>
	<li>Place the testing apparatus (Figure 2) on the stage of a stereo microscope.</li>
	<li>Connect the force transducer and length controller cables to the testing apparatus. Turn on the force transducer. Turn on the length controller so that it remains rigid. (Note: the length controller provides the ability to stretch or shorten a muscle preparation during a contraction. However, this feature of the length controller is not used in the method described herein. Therefore, the length controller can be thought of as a rigid attachment point mounted to an XYZ positioning system).</li>
	<li>With a disposable transfer pipette, fill the experimental chamber with testing solution.</li>
	<li>Use forceps to pick up a suture loop by one of the tails and hang it on the force transducer tube. Hang a second suture loop on the tube attached to the length controller. (Note: gripping a suture loop on the curved part can kink the suture and cause it to break during subsequent steps).</li>
	<li>With a disposable transfer pipette, transfer a zebrafish larva to a small Petri dish filled with testing solution. Wait for the anesthetic in the testing solution (tricaine) to take effect (~1 min). With a forceps, gently nudge the tail and verify that the larva is anesthetized by a lack of touch-evoked swimming.</li>
	<li>Use a glass pipette to transfer the larva to the experimental chamber.</li>
	<li>By gently nudging the larva with closed forceps, guide the anterior portion of the larva through the suture loop on the force transducer tube. Guide the anterior portion of the larva through the suture loop on the tube. Grasp both suture loop tails with forceps and pull them simultaneously to tighten the suture loop posterior to the yolk sac or swimbladder (Figure 3A).</li>
	<li>With a forceps, hold one suture loop tail and pull, causing the larva to swivel 90&deg; around the tube until the lateral side of the larva faces up (Figure 3B). If the loop was tightened enough, there will be some resistance to the pull; the larva should not swivel easily. If the loop was tightened too much, the larva will not swivel around the tube.</li>
	<li>Using the XYZ positioning device attached to the length controller, move the length controller tube along the X-axis (axis definitions in Figure 2A) and under the trunk and tail of the larva. Leave space between the ends of the length controller tube and the force transducer tube.</li>
	<li>Guide the suture loop over the tail of the larva and tighten the suture loop as previously described (Figure 3C). You may need to swivel the posterior part of the larva so that the lateral side faces up. Trim the suture loop tails (Figure 3D).</li>
</ol>
4. Position Larva in Experimental Chamber
<ol>
	<li>Move the larva to an appropriate distance from the chamber bottom to ensure the larva will be within "working distance" of an inverted microscope objective during subsequent steps. To accomplish this, use the XYZ positioning devices to slowly lower the tubes (with attached larva) along the Z-axis until the tubes just touch the bottom of the chamber. Then, raise the tubes until the larva is an appropriate distance from the chamber bottom (~100 &mu;m).</li>
	<li>Using the XYZ positioning device attached to the length controller, adjust the length controller tube along the Y-axis to align the long axis of the larva with the long axis of the force transducer tube.</li>
</ol>
5. Record Force During a Maximal Twitch Contraction
<ol>
	<li>Move the testing apparatus to the stage of an inverted microscope.</li>
	<li>Adjust the chamber temperature to a desired value. To begin, connect the water bath circulator, thermometer, and temperature controller to the testing apparatus. Turn on the necessary components and adjust the setting on the temperature controller until the thermometer reports the desired value. Data included in this article were collected at 25 &deg;C, but measurements may also be made at RT or at 28.5 &deg;C.</li>
	<li>Connect cables from the stimulator to the testing apparatus. Turn on the power to the stimulator but do not stimulate the larva until step 5.6.</li>
	<li>Make sure the larva is parallel to the chamber bottom. Through a 40X objective, view the portion of the larva between the ends of the tubes. If parallel to the bottom, both ends of the larva will be in focus. If needed, adjust the force transducer tube along the Z-axis until both ends are in focus.</li>
	<li>Verify that the larva length is shorter than optimal. Turn on the video sarcomere length system and rotate the video camera such that the striations are parallel to the sides of the video frame. This system monitors striation spacing by analyzing variations in pixel intensity along each horizontal row of pixels within a user-defined region of interest (ROI). The results for all rows within the ROI are averaged and reported with a frequency equivalent to the video frame-rate (&ge;80 sec-1). The striation spacing is used as an indicator of sarcomere length.</li>
	<li>Adjust the microscope to focus on peripheral fibers and note the indicated sarcomere length. If necessary, use the XYZ positioning device attached to the length controller to adjust length of the larva (X-axis) until the sarcomere length is less than optimal (e.g. 1.90 &mu;m).</li>
	<li>Adjust the stimulation current to optimize twitch force. To begin, set the output current on the stimulator to a low magnitude (e.g. 100 mA). The stimulator can be triggered manually or by a computer running a custom LabVIEW program. Elicit a twitch of the larval muscles with a current pulse of 0.2 msec in duration.</li>
	<li>Use an oscilloscope to record the force output and measure the peak twitch force using the oscilloscope's cursors. Increase the current by 50 mA increments and measure the peak twitch force at each current level. Wait 30 sec between twitches to prevent fatigue. As stimulation current increases, peak twitch force typically increases to a maximum and then gradually decreases. The current at which the larva generates the greatest force is the optimal stimulation current. Set the current amplitude to the optimal stimulation current.</li>
	<li>Using the XYZ positioning device, adjust the length of the larva (and thus, sarcomere length) in order to elicit maximum twitch force. Wild-type zebrafish larvae (3-7 dpf) generate maximum twitch force at sarcomere lengths of 2.10 &mu;m or 2.15 &mu;m. However, the sarcomere length can be set to 2.08 &mu;m to avoid excess strain on the larva.</li>
	<li>Elicit a twitch of the larval muscles. Use the oscilloscope to record the force response and save the record for subsequent analysis.</li>
</ol>
6. Measure Musculature Dimensions with Larva at Optimal Length
<ol>
	<li>Move the testing apparatus back to the stereo microscope.</li>
	<li>Using the eyepiece scale, measure the height of the musculature as viewed from the side. Then, taking care not to change the length of the larva, swivel the larva by 90&deg; using the suture loop tails in order to view the larva from the bottom. Measure the width of the musculature as viewed from the bottom. Take the measurements at an anatomical landmark (e.g. urogenital opening) (Figure 4).</li>
	<li>Cut the suture loops with a microblade to release the larva from the testing equipment.</li>
</ol>

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Open access fees have been paid for by Aurora Scientific.

This method measures force generation during a twitch to assess muscle function in trunk muscles of zebrafish larvae. Although tetanic contractions can be elicited in zebrafish larvae (e.g. by 200 stimulation pulses/sec for a duration of 0.2 sec), the maximum tetanic force is only 10-15% greater than the maximum twitch force. Therefore, the force generated during a twitch is a reasonable measure of force generating capability. Twitches are preferred over tetanic contractions because twitches are less likely to cause ripping or slipping at the suture ties.
In order to generate meaningful data with this technique, maximum twitch force should be achieved for each larva and the variability between experimental groups should be minimized. With these goals in mind, we offer the following suggestions. First, take care when tying the larva to the force transducer and the length controller tubes. If the suture loops are tightened too much, the suture will cut through the muscle tissue. If the suture loops are not tightened enough, the force generated by the larva will not be fully transmitted to the force transducer. Both situations, but especially the latter, underestimate maximum twitch force. Second, since testing multiple experimental groups can take several hours (20-30 min/larva), alternate between groups because larvae will continue to develop during the testing period.
While some of the mentioned equipment is essential for measurement of maximum twitch force (e.g. force transducer, current stimulator), other items are not absolutely necessary. The video sarcomere length system is desirable but not required. As an alternative, a series of twitches can be used to find optimal length, during which the length of the larva is adjusted until maximum twitch force is achieved. A temperature control system is also not absolutely necessary. Temperature control is critical when measuring twitch kinetics, which are highly sensitive to temperature, whereas maximum twitch force is not particularly sensitive to small changes in temperature and could be measured at room temperature. Note that regardless of the temperature in the chamber during force testing, the larvae should be maintained at the optimal growth temperature of 28.5 &deg;C24 prior to force testing for accurate staging.
The larvae are tested in a Tyrodes solution containing tricaine. We use 0.02% (w/v) tricaine, the concentration recommended for anesthesia24, to eliminate spontaneous contractions evoked by the nervous system and thus prevent fatigue during force testing. Tricaine also facilitates the tie-on step and reduces overall testing time. However, we observe that including tricaine in the testing solution consistently reduces the maximum twitch force by approximately 30%. A similar effect has also been observed in tadpole tail muscle, where tricaine reduced force generation after neuromuscular transmission was blocked, suggesting that tricaine has a direct effect on muscle25. Tricaine may reduce muscle cell excitability by reducing sodium conductance across the cell membrane, as it does in nerve cells26. Other options for blocking activation by motoneurons are d-tubocurarine and &alpha;-bungarotoxin but, unlike tricaine, these compounds are not skin-permeable and must be injected directly into the head, spinal cord, or heart27. Individual investigators will need to assess whether or not tricaine is desirable for their specific application. If tricaine is included in the testing solution, the concentration should be consistent between experiments and researchers should verify that the effect of the tricaine does not vary between experimental groups.
We describe this method for larvae as young as 3 dpf and as old as 7 dpf. Although muscle fibers appear to be functional as early as 17 hr post-fertilization, when spontaneous tail movements begin27, the short length of the tail before 3 dpf hinders tying the larva to the testing equipment. We typically do not test larva after 7 dpf since many disease models do not survive much longer than this time. If testing larvae beyond 5 dpf, the larvae should be fed. We have observed that unfed larvae have smaller muscles and generate less maximum twitch force than fed larvae, likely due to the diminishing yolk sac. Thus it may be desirable to test larvae between 3-5 dpf, to avoid the additional variable of external feeding.
In summary, we describe a quantitative and reliable method for measuring force generation during a maximal twitch contraction of zebrafish larval trunk muscle. This method can be used to assess the overall health of zebrafish larval muscle and specifically provides information about muscle function. In addition to providing information about the magnitude of force generation, this technique can be used to study the kinetics of force generation or be adapted to study muscle fatigue22. Although we describe this technique for use with wild-type larvae, this method can be used for genetically modified larvae or for larvae treated with drugs or toxicants, to characterize muscle disease models and evaluate treatments, or to study muscle development, muscle injury, or muscle-related chemical toxicity.

Young zebrafish (Danio rerio) larvae, 3-7 days post-fertilization (dpf), are increasingly recognized as a useful organism for skeletal muscle research. Young larvae are used to model human muscle disease1-9, evaluate drugs and therapeutic strategies10-11, study muscle injury12, understand muscle development13-16, and investigate muscle-related chemical toxicity17-19. Typical studies in these areas examine the degree to which healthy muscle is rendered abnormal by genetic manipulation or exposure to toxicants, and some studies examine the degree to which abnormal muscle responds to treatment. Critical to the success of these studies is the ability to accurately assess muscle health.
While there are a variety of methods available to assess muscle health in zebrafish larvae, few provide direct information about muscle function. Muscle health is usually evaluated by appearance, as assessed by histological staining6,8,11, immunostaining9,15,16,18, light microscopy3,13, electron microscopy3,4,14,16, or birefringence7,9,11, but these techniques provide morphological information only. Trunk and tail displacements and swimming speed4,17 evaluate motor function, but these are not direct measures of muscle function since they also reflect neural input, energy metabolism, and other processes.
In contrast, measuring force generation during contraction provides a direct assessment of muscle function and represents a measure of overall muscle health. Added benefits of this approach include straightforward data analysis and quantitative results. In this video article, we provide a detailed procedure for measuring force generation by larval muscles, in the hope that more researchers will use this method to complement existing measures of muscle health in their research.
The overall goal of this method is to measure force generation during contraction of zebrafish larval trunk muscle. To accomplish this goal, a zebrafish larva is anesthetized and placed 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. Muscle activation is accomplished by electric field stimulation, and the stimulation current and the length of the larva are adjusted to produce maximum twitch force. An isometric twitch contraction is elicited and the force response is recorded for analysis.
To be clear, this technique does not measure forces generated by larval muscles during swimming. Because both ends of the larva are tied to equipment and because the larva remains anesthetized, it cannot initiate movement during testing. Furthermore, field stimulation activates all the muscle fibers at the same time to induce a bilateral contraction, which is not what naturally occurs20. Therefore, rather than measuring actual forces generated during swimming, this technique determines the force generating capability of the larval muscles.
We have used this technique to demonstrate muscle weakness in a zebrafish model of nemaline myopathy21, as well as to evaluate the effect of antioxidant treatment on muscle function in a zebrafish model of multi-minicore disease22. Others have used a similar technique23 to examine the effects of an environmental pollutant on muscle function19.

<table border="1">
		<tbody>
		 <tr>
			<td></td>
			<td></td>
			<td></td>
			<td>REAGENTS</td>
		 </tr>
		 <tr>
			<td>Tricaine powder</td>
			<td>Sigma-Aldrich</td>
			<td>A5040</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Sodium chloride</td>
			<td>Sigma-Aldrich</td>
			<td>S7653</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Potassium chloride</td>
			<td>Sigma-Aldrich</td>
			<td>P9541</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Calcium chloride dihydrate</td>
			<td>Sigma-Aldrich</td>
			<td>223506</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Magnesium chloride hexahydrate</td>
			<td>Sigma-Aldrich</td>
			<td>M2670</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Sodium phosphate monobasic</td>
			<td>Sigma-Aldrich</td>
			<td>S0751</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Sodium bicarbonate</td>
			<td>Sigma-Aldrich</td>
			<td>S6297</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Ethylenediaminetetraacetic acid disodium salt dihydrate</td>
			<td>Sigma-Aldrich</td>
			<td>E5134</td>
			<td></td>
		 </tr>
		 <tr>
			<td></td>
			<td></td>
			<td></td>
			<td>EQUIPMENT</td>
		 </tr>
		 <tr>
			<td>Nonsterile-suture</td>
			<td>Ashaway Line &amp; Twine</td>
			<td>S30002</td>
			<td>USP 10/0 monofilament nylon (3 ply)</td>
		 </tr>
		 <tr>
			<td>Forceps</td>
			<td>Fine Science Tools</td>
			<td>11251-20</td>
			<td>Dumont #5</td>
		 </tr>
		 <tr>
			<td>Spring scissors</td>
			<td>Fine Science Tools</td>
			<td>15000-08</td>
			<td>Vannas</td>
		 </tr>
		 <tr>
			<td>Stereo microscope</td>
			<td>Leica Microsystems</td>
			<td>MZ8</td>
			<td>Illuminated with Fostec EKE ACE I light source</td>
		 </tr>
		 <tr>
			<td>Force transducer</td>
			<td>Aurora Scientific</td>
			<td>400A</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Length controller</td>
			<td>Aurora Scientific</td>
			<td>318B</td>
			<td></td>
		 </tr>
		 <tr>
			<td>XYZ positioning devices</td>
			<td>Parker Hannifin</td>
			<td>3936M</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Thermometer</td>
			<td>Physitemp</td>
			<td>BAT-12</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Disposable transfer pipette</td>
			<td>Fisher Scientific</td>
			<td>13-711-9AM</td>
			<td>Cut end to widen opening and facilitate larva transfer</td>
		 </tr>
		 <tr>
			<td>Petri dish</td>
			<td>Fisher Scientific</td>
			<td>08-757-11YZ </td>
			<td></td>
		 </tr>
		 <tr>
			<td>Glass pipette</td>
			<td>Fisher Scientific</td>
			<td>13-678-8B</td>
			<td>Cut end (and fire-polish) to widen opening and facilitate larva transfer</td>
		 </tr>
		 <tr>
			<td>Inverted microscope</td>
			<td>Carl Zeiss Microscopy</td>
			<td>Axiovert 100</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Water bath circulator</td>
			<td>Neslab Instruments</td>
			<td>RTE-111</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Temperature controller</td>
			<td>Alpha Omega Instruments</td>
			<td>Series 800</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Stimulator</td>
			<td>Aurora Scientific</td>
			<td>701C</td>
			<td>High-power, follow stimulator</td>
		 </tr>
		 <tr>
			<td>Video sarcomere length system</td>
			<td>Aurora Scientific</td>
			<td>900B-5A</td>
			<td></td>
		 </tr>
		 <tr>
			<td>LabVIEW software</td>
			<td>National Instruments</td>
			<td></td>
			<td></td>
		 </tr>
		 <tr>
			<td>Oscilloscope</td>
			<td>Nicolet Technologies</td>
			<td>ACCURA 100</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Microblade</td>
			<td>Fine Science Tools</td>
			<td>10050-00</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Microblade holder</td>
			<td>Fine Science Tools</td>
			<td>10053-13</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Data analysis software (Signo)</td>
			<td>Alameda Applied Sciences</td>
			<td></td>
			<td></td>
		 </tr>
		</tbody>
 </table>

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.

In healthy wild-type zebrafish larvae, the muscle fibers should be parallel to one another without large gaps between them and have evident striations (Figure 5A). Wild-type zebrafish larvae that do not exhibit these features, or with evident damage such as detached fibers (Figure 5B), should be discarded.
A representative plot of peak twitch force versus stimulation current for a single zebrafish larva is shown in Figure 6. For wild-type zebrafish larvae between 3-7 dpf, the optimal stimulation current is typically between 400-600 mA, with 3 dpf larvae generally requiring greater stimulation current than 6-7 dpf larvae.
The raw force data (collected during step 5.8) has to be processed and analyzed with data analysis software. First, the baseline of the force record is set to zero. Second, the voltage output of the force transducer is converted to force (mN) (see manufacturer's instructions to generate a calibration curve for the force transducer). A representative force response collected during a maximal twitch contraction of a single larva is shown in Figure 7. Data analysis software can be used to measure peak force and other features of the force response.
A representative set of peak force data from maximal twitch contractions is shown in Figure 8A. Typical peak twitch force values for wild-type 3-7 dpf larvae range from 0.9 to 1.7 mN, with older larvae generating more force than younger larvae. Differences in peak twitch force can be due to normal processes like growth and development (Figure 8) or abnormal processes such as gene mutation-related pathology21,22.
Normalization by muscle cross-sectional area (CSA) can be used to determine the degree to which differences in peak twitch force are simply due to differences in size of the musculature21,22. Muscle CSA can be estimated using the formula: CSA = &pi;(A/2)(B/2), where A is the height of the musculature as viewed from the side, B is the width of the musculature as viewed from the bottom, and an elliptical cross-section is assumed. Typical CSA values for wild-type 3-7 dpf larvae range from 0.027 to 0.034 mm2, with 3-4 dpf larvae generally showing smaller CSA values than 5-7 dpf larvae. A representative set of normalized peak force data from maximal twitch contractions is shown in Figure 8B. Typical normalized peak twitch force values for wild-type 3-7 dpf larvae range from 34 to 51 mN/mm2, with 4-7 dpf larvae generally showing greater values than 3 dpf larvae.
<img alt="Figure 1" src="/files/ftp_upload/50539/50539fig1.jpg" /> 
 Figure 1. Suture loop. Arrows point to the suture loop tails.
<img alt="Figure 2" fo:content-width="5in" fo:src="/files/ftp_upload/50539/50539fig2highres.jpg" src="/files/ftp_upload/50539/50539fig2.jpg" /> 
Figure 2. (A) Testing apparatus with labeled components. (B) Close-up views of the experimental chamber. (a) Experimental chamber with transparent bottom. (b) Force transducer. (c) Length controller. (d) XYZ positioning devices. The X, Y, and Z axes are defined in the upper right corner. (e) Temperature control system utilizing thermoelectric modules. Tubing accommodates water flow for cooling of thermoelectric modules. (f) Stainless steel tube attached to force transducer. (g) Stainless steel tube attached to length controller. (h) Thermometer microprobe. (i) Platinum parallel plate electrodes, spanning the length of the chamber. Platinum plates are 2.5 mm high and 0.255 mm thick.
<img alt="Figure 3" fo:content-width="6in" fo:src="/files/ftp_upload/50539/50539fig3highres.jpg" src="/files/ftp_upload/50539/50539fig3.jpg" /> 
 Figure 3. Tying larva into experimental chamber. (A) Larva tied on at anterior end but not yet swiveled 90&deg;. (B) Larva after swiveled 90&deg;. (C) Larva tied on at posterior end but not yet swiveled. (D) Larva after swiveled and suture loop tails are trimmed.
<img alt="Figure 4" fo:content-width="6in" fo:src="/files/ftp_upload/50539/50539fig4highres.jpg" src="/files/ftp_upload/50539/50539fig4.jpg" /> 
 Figure 4. Measurements for cross-sectional area estimation. Musculature as viewed from the (A) side and (B) bottom. Placement of the red bars indicate the location of the urogenital opening. The length of the red bars indicate the height and width of the musculature as viewed from the side and bottom respectively.
<img alt="Figure 5" fo:content-width="6in" fo:src="/files/ftp_upload/50539/50539fig5highres.jpg" src="/files/ftp_upload/50539/50539fig5.jpg" /> 
 Figure 5. Lateral view of zebrafish larvae trunk musculature. (A) Healthy tissue. (B) Tissue with evident damage. Contractures resulting from fiber detachments are marked with asterisks.
<img alt="Figure 6" src="/files/ftp_upload/50539/50539fig6.jpg" /> 
 Figure 6. Representative plot of peak twitch force versus stimulation current. The optimal stimulation current is 500 mA.
<img alt="Figure 7" src="/files/ftp_upload/50539/50539fig7.jpg" /> 
 Figure 7. Representative force record for a single twitch contraction. This contraction was elicited with a stimulus pulse at 0 msec. The peak force is 1.56 mN.
<img alt="Figure 8" fo:content-width="6in" fo:src="/files/ftp_upload/50539/50539fig8highres.jpg" src="/files/ftp_upload/50539/50539fig8.jpg" /> 
 Figure 8. Representative force data from 3-7 dpf larvae. (A) Peak force data from maximal twitch contractions. (B) Peak force data from maximal twitch contractions normalized to CSA. Older larvae (6-7 dpf) were fed Hatchfry Encapsulon Grade 0 with Spirulina (Argent Laboratories) starting on 5 dpf. Means + standard deviations are reported with N=5 in each group. Groups significantly different from 3 dpf larvae (*) and 4 dpf larvae (#) are indicated (ANOVA, P&lt;0.05). The significant increase in normalized force between 3 and 4 dpf (B) indicates an increase in the intrinsic force generating capability during this time period, whereas the increase in force between 4 and 6-7 dpf (A) is attributed to growth based on no change from 4 to 7 dpf in normalized force.

Watch this Scientific Journal Video about Force Measurement During Contraction to Assess Muscle Function in Zebrafish Larvae at JoVE.com

Force Measurement During Contraction to Assess Muscle Function in Zebrafish Larvae

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 ...

force-measurement-during-contraction-to-assess-muscle-function

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15.6K Views.  University of Michigan. 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.

Video: Force Measurement During Contraction to Assess Muscle Function in Zebrafish Larvae

High-resolution Measurement of Odor-Driven Behavior in Drosophila Larvae

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral paradigm, the experimenter usually assumes that the rapid diffusion of odorant molecules from the source leads to the creation of a stable gradient in the dish. To establish a quantitative correlation between sensory inputs and behavioral responses, it is necessary to achieve a more thorough characterization of the odorant stimulus conditions. In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral ...

Microinjection of Zebrafish Embryos to Analyze Gene Function

One of the advantages of studying zebrafish is the ease and speed of manipulating protein levels in the embryo.  Morpholinos, which are synthetic oligonucleotides with antisense complementarity to target RNAs, can be added to the embryo to reduce the expression of a particular gene product.  Conversely, processed mRNA can be added to the embryo to increase levels of a gene product.  The vehicle for adding either mRNA or morpholino to an embryo is microinjection.  Microinjection is efficient and rapid, allowing for the injection of hundreds of embryos per hour.  This video shows all the steps involved in microinjection.  Briefly, eggs are collected immediately after being laid and lined up against a microscope slide in a Petri dish.  Next, a fine-tipped needle loaded with injection material is connected to a microinjector and an air source, and the microinjector controls are adjusted to produce a desirable injection volume.  Finally, the needle is plunged into the embryo's yolk and the morpholino or mRNA is expelled.

This video shows how morpholino or mRNA can be injected into zebrafish embryos at the one-cell stage to decrease or increase the level of specific gene products during subsequent development.

One of the advantages of studying zebrafish is the ease and speed of manipulating protein levels in the embryo.  Morpholinos, which are synthetic ...

Using the optokinetic response to study visual function of zebrafish

Optokinetic response (OKR) is a behavior that an animal vibrates its eyes to follow a rotating grating around it. It has been widely used to assess the visual functions of larval zebrafish1-5. Nevertheless, the standard protocol for larval fish is not yet readily applicable in adult zabrafish. Here, we introduce how to measure the OKR of adult zebrafish with our simple custom-built apparatus using a new protocol which is established in our lab. Both our apparatus and step-by-step procedure of OKR in adult zebrafish are illustrated in this video. In addition, the measurements of the larval OKR, as well as the optomotor response (OMR) test of adult zebrafish, are also demonstrated in this video. This OKR assay of adult zebrafish in our experiment may last for up to 4 hours. Such OKR test applied in adult fish will benefit to visual function investigation more efficiently when the adult fish vision system is manipulated.
Su-Qi Zou and Wu Yin contributed equally to this paper.

Optokinetic response has been widely used to assess the visual functions of larval zebrafish. Nevertheless, the standard protocol for larval fish is not yet readily applicable in adults1-5. Here, we introduce how to measure the OKR of adult zebrafish using a new protocol which is established in our lab.

Optokinetic response (OKR) is a behavior that an animal vibrates its eyes to follow a rotating grating around it. It has been widely used to assess the ...

Intravenous Microinjections of Zebrafish Larvae to Study Acute Kidney Injury

In this video article we describe a zebrafish model of AKI using gentamicin as the nephrotoxicant.  The technique consists of intravenous microinjections on 2 dpf zebrafish. This technique represents an efficient and rapid method to deliver soluble substances into the bloodstream of zebrafish larvae, allowing for the injection of 15-20 fish per hour. In addition to AKI studies, this microinjection technique can also be used for other types of experimental studies such as angiography.  We provide a detailed protocol of the technique from equipment required to visual measures of decreased kidney function. In addition, we also demonstrate the process of fixation, whole mount immunohistochemistry with a kidney tubule marker, plastic embedding and sectioning of the larval zebrafish. We demonstrate that zebrafish larvae injected with gentamicin show morphological features consistent with AKI: edema, loss of cell polarity in proximal tubular epithelial cells, and morphological disruption of the tubule.

We describe a technique of microinjecting the aminoglycoside, gentamicin, into 2 days post-fetilization (dpf) zebrafish larvae to induce acute kidney injury (AKI). We also describe a method for whole mount immunohistochemistry, plastic embedding and sectioning of zebrafish larvae to visualize the AKI mediated damage.

In this video article we describe a zebrafish model of AKI using gentamicin as the nephrotoxicant.  The technique consists of intravenous microinjections ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Microgavage of Zebrafish Larvae

The zebrafish has emerged as a powerful model organism for studying intestinal development1-5, physiology6-11, disease12-16, and host-microbe interactions17-25. Experimental approaches for studying intestinal biology often require the in vivo introduction of selected materials into the lumen of the intestine. In the larval zebrafish model, this is typically accomplished by immersing fish in a solution of the selected material, or by injection through the abdominal wall. Using the immersion method, it is difficult to accurately monitor or control the route or timing of material delivery to the intestine. For this reason, immersion exposure can cause unintended toxicity and other effects on extraintestinal tissues, limiting the potential range of material amounts that can be delivered into the intestine. Also, the amount of material ingested during immersion exposure can vary significantly between individual larvae26. Although these problems are not encountered during direct injection through the abdominal wall, proper injection is difficult and causes tissue damage which could influence experimental results. We introduce a method for microgavage of zebrafish larvae. The goal of this method is to provide a safe, effective, and consistent way to deliver material directly to the lumen of the anterior intestine in larval zebrafish with controlled timing. Microgavage utilizes standard embryo microinjection and stereomicroscopy equipment common to most laboratories that perform zebrafish research. Once fish are properly positioned in methylcellulose, gavage can be performed quickly at a rate of approximately 7-10 fish/ min, and post-gavage survival approaches 100% depending on the gavaged material. We also show that microgavage can permit loading of the intestinal lumen with high concentrations of materials that are lethal to fish when exposed by immersion. To demonstrate the utility of this method, we present a fluorescent dextran microgavage assay that can be used to quantify transit from the intestinal lumen to extraintestinal spaces. This test can be used to verify proper execution of the microgavage procedure, and also provides a novel zebrafish assay to examine intestinal epithelial barrier integrity under different experimental conditions (e.g. genetic manipulation, drug treatment, or exposure to environmental factors). Furthermore, we show how gavage can be used to evaluate intestinal motility by gavaging fluorescent microspheres and monitoring their subsequent transit. Microgavage can be applied to deliver diverse materials such as live microorganisms, secreted microbial factors/toxins, pharmacological agents, and physiological probes. With these capabilities, the larval zebrafish microgavage method has the potential to enhance a broad range of research fields using the zebrafish model system.

We present a novel method for microgavage of larval zebrafish utilizing standard embryo microinjection and stereomicroscopy equipment. We demonstrate that microgavage is a safe and efficient technique useful for delivering controlled amounts of diverse materials specifically into the larval zebrafish intestinal lumen.

The zebrafish has emerged as a powerful model organism for studying intestinal development1-5, physiology6-11, disease12-16, and host-microbe ...

Methods to Assess Subcellular Compartments of Muscle in C. elegans

Muscle is a dynamic tissue that responds to changes in nutrition, exercise, and disease state. The loss of muscle mass and function with disease and age are significant public health burdens. We currently understand little about the genetic regulation of muscle health with disease or age. The nematode C. elegans is an established model for understanding the genomic regulation of biological processes of interest. This worm&#8217;s body wall muscles display a large degree of homology with the muscles of higher metazoan species. Since C. elegans is a transparent organism, the localization of GFP to mitochondria and sarcomeres allows visualization of these structures in vivo. Similarly, feeding animals cationic dyes, which accumulate based on the existence of a mitochondrial membrane potential, allows the assessment of mitochondrial function in vivo. These methods, as well as assessment of muscle protein homeostasis, are combined with assessment of whole animal muscle function, in the form of movement assays, to allow correlation of sub-cellular defects with functional measures of muscle performance. Thus, C. elegans provides a powerful platform with which to assess the impact of mutations, gene knockdown, and/or chemical compounds upon muscle structure and function. Lastly, as GFP, cationic dyes, and movement assays are assessed non-invasively, prospective studies of muscle structure and function can be conducted across the whole life course and this at present cannot be easily investigated in vivo in any other organism.

Methods to Assess Subcellular Compartments of Muscle in C. elegans

Skeletal muscle is essential for locomotion and is the bodies&#8217; main protein store. Muscle health measurements within C. elegans are described. Prospective changes to muscle structure and function are assessed using localized GFP and cationic dyes.

Muscle is a dynamic tissue that responds to changes in nutrition, exercise, and disease state. The loss of muscle mass and function with disease and age ...

Measurement of Smooth Muscle Function in the Isolated Tissue Bath-applications to Pharmacology Research

Isolated tissue bath assays are a classical pharmacological tool for evaluating concentration-response relationships in a myriad of contractile tissues. While this technique has been implemented for over 100 years, the versatility, simplicity and reproducibility of this assay helps it to remain an indispensable tool for pharmacologists and physiologists alike. Tissue bath systems are available in a wide array of shapes and sizes, allowing a scientist to evaluate samples as small as murine mesenteric arteries and as large as porcine ileum &#8211; if not larger. Central to the isolated tissue bath assay is the ability to measure concentration-dependent changes to isometric contraction, and how the efficacy and potency of contractile agonists can be manipulated by increasing concentrations of antagonists or inhibitors. Even though the general principles remain relatively similar, recent technological advances allow even more versatility to the tissue bath assay by incorporating computer-based data recording and analysis software. This video will demonstrate the function of the isolated tissue bath to measure the isometric contraction of an isolated smooth muscle (in this case rat thoracic aorta rings), and share the types of knowledge that can be created with this technique. Included are detailed descriptions of aortic tissue dissection and preparation, placement of aortic rings in the tissue bath and proper tissue equilibration prior to experimentation, tests of tissue viability, experimental design and implementation, and data quantitation. Aorta will be connected to isometric force transducers, the data from which will be captured using a commercially available analog-to-digital converter and bridge amplifier specifically designed for use in these experiments. The accompanying software to this system will be used to visualize the experiment and analyze captured data.

This protocol describes the measurement of isometric contraction in an isolated smooth muscle preparation, using an isolated tissue bath system and computer-based data acquisition.

Isolated tissue bath assays are a classical pharmacological tool for evaluating concentration-response relationships in a myriad of contractile tissues.  ...

Design and Development of Aptamer&#8211;Gold Nanoparticle Based Colorimetric Assays for In-the-field Applications

The design and development of an aptamer&#8211;gold nanoparticle (AuNP) colorimetric assay for the detection of small molecules for in-the-field applications was examined. Target selective AuNP based color assays have been developed in controlled proof-of-concept laboratory settings. However, these schemes have not been exerted to a point of failure to determine their practical use beyond laboratory settings. This work describes a generic approach to design, develop, and troubleshoot an aptamer-AuNP colorimetric assay for small molecule analytes and using the assay for in-the-field settings. The assay is advantageous because adsorbed aptamers passivate the nanoparticle surfaces and provide a means to reduce and eliminate false positive responses to non-target analytes. Transitioning this system to practical uses required defining not only the shelf-life of the aptamer-AuNP assay, but establishing methods and procedures for extending the long-term storage capabilities. Also, one of the recognized concerns with colorimetric readout is the burden placed on analysts to accurately identify often subtle changes in color. To lessen the responsibility on analysts in the field, a color analysis protocol was designed to perform the color identification duties without the need for performing this task on laboratory grade equipment. The method for creating and testing the data analysis protocol is described. However to understand and influence the design of adsorbed aptamer assays, the interactions associated with the aptamer, target, and AuNPs require further study. The knowledge gained could lead to tailoring aptamers for improved functionality.

The design and development of an aptamer&#8211;gold nanoparticle colorimetric assay for the detection of small molecules for in-the-field applications was examined. Additionally, a smart-device colorimetric application (app) was validated and long-term storage of the assay was established for use in the field.

The design and development of an aptamer&#8211;gold nanoparticle (AuNP) colorimetric assay for the detection of small molecules for in-the-field ...

Nephrotoxin Microinjection in Zebrafish to Model Acute Kidney Injury

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid decline in renal function and development of the clinical syndrome known as acute kidney injury (AKI). Pharmacological agents used to treat medical circumstances ranging from bacterial infection to cancer, when administered individually or in combination with other drugs, can initiate AKI. Zebrafish are a useful animal model to study the chemical effects on renal function in vivo, as they form an embryonic kidney comprised of nephron functional units that are conserved with higher vertebrates, including humans. Further, zebrafish can be utilized to perform genetic and chemical screens, which provide opportunities to elucidate the cellular and molecular facets of AKI and develop therapeutic strategies such as the identification of nephroprotective molecules. Here, we demonstrate how microinjection into the zebrafish embryo can be utilized as a paradigm for nephrotoxin studies.

Renal injuries incurred from nephrotoxins, which include drugs ranging from antibiotics to chemotherapeutics, can result in complex disorders whose pathogenesis remains incompletely understood. This protocol demonstrates how zebrafish can be used for disease modeling of these conditions, which can be applied to the identification of renoprotective measures.

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid ...