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

Our protocol allows us to study how tissues, such as skeletal muscle, regulate their size, which remains a key question in cell and developmental biology with important physiological and medical implications. The main advantage of our technique is that cellular growth can be observed and measured in live animals in vivo with a high degree of spatial and temporal accuracy. Progress in the basic understanding of early muscle growth can highlight the developmental origins of early muscle problems and lead to the discovery of Novo Therapeutics to protect against muscle loss in elderly and patients with muscle-wasting disorders.

Prepare 1%LMA in a 1.5 milliliter tube and keep it in a 37 degrees Celsius heat block for repeated use. Select fish to be mounted and transiently anesthetize each fish. To avoid heat shock, remove the LMA aliquot from the heat block and let it cool to just above setting.

Take a 60 milliliter Petri dish that has been coated with a layer of 1%LMA and place it on the stage of a dissecting microscope. Transfer the larva with a one milliliter plastic Pasteur pipette onto a 60 millimeter coated Petri dish and remove as much transferred medium as possible. Then, still using the Pasteur pipette, place a few drops of LMA onto the fish and rapidly orient the larva near the upper surface of the LMA with its anteroposterior and dorsoventral axis within 10 degrees of the horizontal, using forceps or a fire-polished, fine glass needle before the LMA sets.

Alternatively, use a one milliliter plastic Pasteur pipette. Collect the larvae with minimum fish medium and transfer the larvae into the aliquot of cooled LMA. Allow the larva to sink for five seconds to become entirely surrounded by LMA.

Then retrieve the larva and transfer it in a drop of LMA onto the agarose-coated Petri dish. Quickly position the larva as demonstrated earlier. If the larvae is not correctly mounted horizontally near the surface of the agarose drop, remove and re-embed, larvae can be easily retrieved by gentle suction using a one milliliter plastic Pasteur pipette and LMA can be gently removed using chem wipes.

Wait for about 10 minutes until LMA sets. Flood the dish with around 10 milliliters of tricaine-containing fish medium. To capture confocal stacks, let the mounted fish rest for at least 10 minutes before scanning, as agarose swelling occurs.

Load the sample dish to the stage of the confocal system. Locate the larvae and focus on the desired somite. Somite 17 may be chosen because of its ease of localization near the anal vent and ease of imaging.

Check by counting somites from the anterior. To capture YZ images, set up as if to capture a Z stack. Start by defining the top and bottom points of the fish.

Both left and right sides can be captured as desired, ensuring that all the rapid YZ scans capture the desired regions. Orient the scan area for the fish as the confocal software permits. Position the fish with the anteroposterior axis, paralleled to the image dex axis and the dorsoventral axis parallel to the Y-axis with somite 17 in the center of the field.

Focus on a mid-level plane in the uppermost myotome, in which the whole epaxial and hypaxial somite halves, together with the vertical and horizontal myosepta are visible and capture a high resolution XY image. Name the file and save the image. To capture the YZ images, draw a precise dorsal to ventral line across the chosen somite, perpendicular to the anteroposterior axis of the fish.

Add a selected anteroposterior position, then perform a Z stack line scan. Repeat the YZ line scan three times at defined anteroposterior positions along the selected myotome to capture YZA, YZM and YZP. Name and save these images with the related XY image.

This protocol has been termed as the four slice method and the images can be analyzed using Zen software or ImageJ. To create a stimulation chamber, take a six by 35 millimeter well plate and create two small openings less than five millimeters in diameter, one centimeter apart on each side of each well using a narrow soldering iron. Once created, the stimulation chamber can be reused.

Thread a pair of silver or platinum wires through the openings of each well. The reusable adhesive material can be applied near the openings to keep the wires in place and ensure a one centimeter separation between the wires. At three days post-fertilization, split fish into three conditions, fish medium control, inactive and inactive plus stem.

For inactive and inactive plus stem groups, anesthetize larvae at 72 hours post-fertilization with 0.6 millimolar tricaine. For the fish medium control fish, leave them unanesthetized. Prepare 60 milliliters of 2%agarose in fish medium.

Melt it thoroughly using a microwave and let it cool. Then add tricaine and pour at least four milliliters into each well of the stimulation chamber. Immediately add custom-made, four-well combs between the electrodes.

Allow 10 minutes for the gel to set. Remove combs carefully to create four rectangular wells. Fill each well with tricaine water and place a single anesthetized, inactive plus stem larva in each well using a micro pipette, with their anteroposterior access perpendicular to the electrodes.

Check under the dissecting fluorescent microscope whether each fish is fully anesthetized within each chamber well. Connect an adjustable electrophysiological, pattern-generating stimulator to the chamber through a polarity controller, using crocodile clips connected to each electrode on one side of the chamber. Then stimulate the fish as described in the text manuscript.

Regularly check under the microscope to confirm the fish are being stimulated. The electrical stimulus should induce a visible bilateral contraction and slight movement once every five seconds following the described parameters. After stimulation, carefully remove fish from each well by gently flushing them out with a plastic pipette and return to the incubator in a fresh, tricaine-containing fish medium.

Pour away the tricaine water from within the chamber and use forceps to cut around and remove the agarose from each well, rinse the wells with tap water and allow them to dry. For the analysis of muscle size, termed the four slice method, XY and YZ images of zebrafish larval muscle were obtained and somite volume was calculated. A strong correlation was observed between the four slice and full stack methods, although the four slice method gave a slightly larger volume estimate.

Volumetric analysis of the adjacent myotomes of somites 16 and 18 revealed that each somite was about 7%larger than the one behind. Further investigation revealed that a two slice method, requiring only a single YZ measurement, gave a reasonably accurate estimate of myotome volume. The myotome grew detectably in length and the cross-sectional area between two and five days post-fertilization, leading to a steady increase in volume.

Muscle made inactive by anesthetic tricaine between the third and fourth day post-fertilization had significantly reduced volume, indicating that larval muscle growth was activity-dependent. Larger variation in the reduction of myotome size was observed when measuring myotome volume at four days post-fertilization, compared to measuring each individual fish at three and four days post-fertilization. Nonetheless, no significant difference was observed in myotome volume between either methods of myotome measurements.

Measurement of the number of fibers within the myotome, as well as average fiber volume, revealed that activity controls both cellular aspects of growth. Ensure that the larvae are completely anesthetized using freshly-prepared tricaine. Partial anesthesia would lead to variable result.

As we know, larvae respond more vigorously to electrical stimulation. This method, combined with downstream transcriptomics and pharmacological experiments, has revealed novel insights into how force sensing drives growth in skeletal muscle.