Preparation of Single-Somite Explants from Zebrafish Embryos

This protocol involves use of live vertebrate embryos younger than 1-day post-fertilization. All experiments were carried out using embryos derived from freely mating adults, and thus are covered under the general animal experiment license of the EPFL granted by the Service de la Consommation et des Affaires Vétérinaires of the canton of Vaud – Switzerland (authorization number VD-H23).

1. Before dissection

1. Obtain embryos from a cross of heterozygous transgenic lines of interest. For explanting two to three somites, only a few embryos are typically needed. In this protocol, one embryo was used for explanting one somite.
2. Prepare 1x E3 buffer from two 50x stocks: stock 1 contains 0.458 mM Na₂HPO₄, 0.042 mM KH₂PO₄, 4.084 mM NaCl, 0.128 mM KCl and stock 2 contains 0.33 mM CaCl₂, 0.33 mM MgSO₄. Prepare 500 mL of 1x E3 buffer by mixing 480 mL of distilled water with 10 mL each of the two 50x stocks.
3. If dissecting on the same day, raise embryos in 25 mL of E3 medium in a Petri dish at 33 °C until three-somite stage. If dissecting the following morning, raise embryos in E3 medium at 28 °C until shield stage before transferring to 19 °C for overnight incubation.
4. For identifying the shield stage or the different somite stages, use a standard lab stereomicroscope with an adjustable magnification range from approximately 0.67x to 4.5x.
5. Assemble tools and reagents needed for dissection.
   1. Aliquot Leibovitz's L-15 medium in 50 mL tubes and store in fridge. We recommend using the same aliquot from the fridge for a maximum of two rounds.
   2. Prepare 50 mL of 2% agarose in L-15 medium by using a magnetic stirrer and keeping the temperature at 85 °C. Once dissolved, store the solution at room temperature.
   3. Prepare 25 mL of 2% low-melting agarose in L-15 medium by using a magnetic stirrer and keeping the temperature at 85 °C. Once dissolved, aliquot the solution in 1.5 mL tubes and store at 4 °C. This will be used for preparing the imaging chamber.
   4. Pour molten 2% agarose into a 20 mm Petri dish until one-third of the depth of the dish is filled and allow the agarose solution to solidify. Dissections will be performed in these Petri dishes. Prepare the Petri dishes with agarose coating well in advance and store at 4 °C.
   5. Assemble the following tools: Pasteur pipette for transfer of embryos, a pair of forceps for dechorionating embryos, a pair of fine forceps for removal of skin of embryos, a micro knife for making incisions in embryos, fire-polished glass pipette for transferring explants to imaging chamber.
   6. Sterilize the glass pipette in 100% ethanol for 15 min and rinse twice by pipetting 5 mL of L-15 medium up and down.
   7. Prepare an eye-lash tool by sticking an eyelash or hair from the eyebrow to a glass capillary.
6. Use a fluorescent stereoscope with an adjustable magnification range from approximately 0.63x to 6.3x and standard GFP and RFP filters to sort positive transgenic embryos and transfer to a separate Petri dish with E3 medium.

2. Single-somite explant preparation

1. Transfer 2 or 3 embryos to an agarose-coated dish (made in step 1.5.4) filled with 10 mL of L-15 medium. Ensure that the dish along with L-15 medium is pre-warmed at 28 °C in an incubator for 30 min before transferring embryos.
2. Carefully dechorionate (Figure 1 A,B) an embryo with a pair of forceps. Hold the chorion with one of the forceps and use the second forceps to pinch and pull on the chorion close to where the first forceps is positioned.
3. Repeat this step one more time to open the chorion widely after which the embryo can be gently pushed out. Ensure that the embryo is not damaged or squeezed during dechorionation.
4. Orient the embryo on its lateral side and hold on to the skin in between the head and tail of the embryo with one of the fine forceps. Use the second fine forceps to pinch on the skin close to where the first forceps is still positioned.
5. Move the two forceps away from each other, while holding on to the skin, thus creating a large opening. Release one of the forceps from the skin and move it close to where the second forceps is held and repeat the skin removal process until the skin is removed in somite-forming regions.
   NOTE: It is critical to completely remove the skin surrounding the somites that will be explanted, else it gets difficult to do finer dissections in the later steps of the protocol.
6. Use a micro knife to scrape off the yolk (Figure 1C - explant with skin removed and yolk scraped off) and make a series of incisions as follows:
   1. Slice through the embryo with the first cut anterior to the first morphological somite in the embryo and the second cut posterior to the most recently formed somite (Figure 1D). Slice once more posterior and anterior to somites of interest (Figure 1E).
   2. Use the micro knife to scrape off any remaining yolk from the explanted regions of the embryo. Try and remove as much of the yolk as possible to reduce the possibility of explants getting stuck to the micro knife. If this happens, use an eye-lash tool to move the explant back to solution while keeping the micro knife immersed in the medium.
   3. Position the explant dorsally and make a slice along the anteroposterior axis through the notochord, close to somites on one side (Figure 1F,G).
   4. Take the explant without the notochord and slice through the lateral plate mesoderm (LPM) close to the somites.
      NOTE: This step can be exchanged with the previous step i.e., it works equally well to slice through the LPM first followed by slicing through the notochord.
7. Choose the somite to be explanted and make two slices posterior and anterior to the somite respectively (Figure 1H).
8. Finally, rotate the somite by 90° using the eye-lash tool to visualize the neural plate still attached to the somite. Make a slice in between the somite and the neural plate, thus releasing a single somite (Figure 1I).
   NOTE: The neural plate can also be removed immediately after step 2.6.4, i.e., after removal of the notochord and the LPM and before dissecting a somite from neighboring somites.

3. Imaging single-somite explants

NOTE: Here, the procedure for imaging single-somite explants using a single-view light-sheet microscope is described. As an alternative, the explanted single somites can also be imaged in a confocal microscope, which is more widely available or even in a widefield microscope if exclusively interested in following overall morphology of explants.

1. Prepare an imaging chamber as follows:
   1. Fill a 1 mL syringe with silicone rubber formulation. Cut a 200 µL pipette tip about 1 cm away from the mouth and insert the syringe into the pipette tip.
   2. Manually wrap the membrane holder with a pre-cut fluorinated ethylene propylene (FEP) membrane strip (Figure 2A-C) and fix the imaging chamber on to the membrane holder, which fit snugly into each other (Figure 2D).
   3. Add silicone rubber formulation along the intersection of the chamber and the membrane and dry it overnight, which allows the membrane to stick to the chamber. Remove the membrane holder 1 h before the explant experiment.
   4. Heat an aliquot containing 0.5 mL of 2% low-melting agarose made with L-15 medium at 75 °C for 10 min. Add 160 µL of the heated low-melting agarose to the center of the chamber and place an imaging mold in the agarose (Figure 2E,F) and hold it upright until the agarose solidifies.
   5. Add another 160 µL of heated low-melting agarose to the two ends of the mold and transfer the entire unit to 4 °C for 30 min.
   6. Add 600 µL of L-15 medium to the chamber and remove the mold from the chamber with a forceps. Wait until the chamber with L-15 medium returns to room temperature.
2. Transfer the explanted somite with a polished glass pipette by pipetting to the imaging chamber and use an eye-lash tool to position the explant in the center of the chamber.
3. Align the imaging lasers as per manufacturer's instructions and carefully transfer the imaging chamber to the microscope equipped with an incubation chamber to maintain the explants at 28 °C.
4. Perform a second round of laser alignment with regions of interest from the transgenic line, followed by setting up the desired z-stack, time interval and duration for time-lapse imaging. For this experiment, acquire 70 z-slices with a spacing of 2 µm and a frame interval of 2 min.
5. Perform dual-colored light-sheet imaging (Figure 1J) of the explants with 561 nm (10% laser power, 100 ms exposure) and 488 nm lasers (10% laser power, 100 ms exposure). In this protocol, the signal was collected with a 25x/1.1 NA objective and through 561/25 nm and 525/50-25 bandpass filters respectively onto a sCMOS camera.