Stab Wound Injury Model of the Adult Optic Tectum Using Zebrafish and Medaka for the Comparative Analysis of Regenerative Capacity

All experimental protocols were approved by the Institutional Animal Care and Use Committee at the National Institute of Advanced Industrial Science and Technology. Zebrafish and medaka were maintained according to standard procedures²⁸.

1. Stab wound injury in the adult optic tectum

1. Prepare a 0.4% (w/v) tricaine stock solution for anesthesia. For a 100 mL stock solution, dissolve 400 mg tricaine methanesulfonate (see Table of Materials) in 90 mL of distilled water and adjust the pH to 7.0 using 1 M Tris HCl buffer (pH 9.0). Once the pH is adjusted, add water up to 100 mL, then make appropriate aliquots and store them at -20 °C.
2. To anesthetize the adult zebrafish or medaka, prepare a 0.02 % (w/v) tricaine solution by diluting the 0.4 % tricaine stock solution with fish facility water.
3. Anesthetize the fish with 0.02% tricaine solution; they become anesthetized when they do not move or respond to touch (1-2 min).
4. Place the anesthetized fish on a Styrofoam tray (see Table of Materials) and fix the fish upright between two 30 G needles inserted vertically into the Styrofoam.
5. Hold the fish body to prevent the fish head from moving and vertically insert a 30 G needle through the skull into the medial region of one of the optic tectum hemispheres (Figure 1A,B). The insertion depth is ~0.75 mm, almost equal to half the length of the 30 G bevel. This insertion depth induces brain injury on zebrafish and medaka because of the similar body size of these fish.
   NOTE: Medaka has scales on the skull. Scratch and remove the scales using the 30 G needle to efficiently and accurately induce the stab wound injury (Figure 1C).
6. Transfer the injured fish into a fish tank with fresh fish facility water and move the tank to the fish breeding system once they have fully recovered from the anesthesia.
   NOTE: Fish will recover and start freely moving within a few minutes.
7. For lineage analysis after the injury, prepare fresh fish facility water containing 5 mM bromodeoxyuridine (BrdU) (see Table of Materials) and then incubate the fish with this 5 mM BrdU solution for a predetermined amount of time^13,19 (as determined by an experimental design). Then dissect these fish as per step 2 and evaluate BrdU incorporation and cell lineage identity as appropriate.
   ​NOTE: This step is an optional step for cell lineage analysis. To detect BrdU signals, perform antigen retrieval as shown in step 4.3.

2. Brain dissection

1. Prepare 0.02% tricaine solution, a Styrofoam tray for dissection, and a 10 mL syringe containing phosphate-buffered saline (1x PBS) with an extension tube and 30 G needles (see Table of Materials) for intracardial perfusion.
2. Anesthetize the injured fish at selected time points.
3. Place paper towels on the Styrofoam tray and place the anesthetized fish on the paper towels.
4. Hold the fish body in place by vertically inserting two 30 G needles on either side of the anal fin (Figure 2A).
5. Make a ventral incision from the anus to the heart (Figure 2B) and keep the heart visible by inserting another two 30 G needles on each side of this organ (Figure 2C).
   NOTE: The heart is covered in the silvery epithelial layer of the hypodermis in both zebrafish and medaka (Figure 2C).
6. Gently remove the silver epithelial layer with the tip of the forceps (Figure 2D).
7. Insert the 30 G needle into the ventricle, keeping the bevel up, and make an incision into the atrium using the forceps (Figure 2E). Carefully press down on the syringe to push the 1x PBS and confirm that the atrium flushed the blood.
   NOTE: To prevent the needle from penetrating the ventricle, carefully insert the needle to the depth of the half bevel and then adjust the insertion depth. If the perfusion is well done, gills turn white (Figure 2F,G).
8. Stop the perfusion after the blood is drained and the gills turn white.
9. Remove the needles fixing the fish body in place and fix the fish ventral side down (Figure 2H). Cut the spinal cord and remove the skull from the optic tectum and telencephalon (Figure 2I). Then, cut the optic nerves and dissect the brain carefully.
   NOTE: Watch out for the optic nerves during the dissection because the nerves are connected to the tectum. When the optic nerve is pulled, the optic tectum can be detached from the brain. If the blood removal is not complete, the brain looks light pink (the right brain in Figure 2J).
10. Transfer the brains into 1.5 mL tubes with 1 mL of 4 % paraformaldehyde in 1x PBS and fix them overnight at 4 °C.

3. Preparation of frozen sections

1. Wash the fixed brains three times in 1x PBS for 5 min each.
2. To cryoprotect the brains, transfer them into 1.5 mL tubes with 1 mL of 30 % (w/v) sucrose in 1x PBS and incubate them overnight at 4 °C. Prepare the embedding compound (2:1 mixture of OCT compound and 30% sucrose, see Table of Materials) by combining 30 mL of OCT and 15 mL of 30 % sucrose. Store this at 4 °C.
   NOTE: To remove the bubbles from the mixed compound, centrifuge the 50 mL tubes at 10,000 x g for 2 min at room temperature or place the tubes upright overnight.
3. Incubate an aluminum block overnight at -80 °C in order to cool the cryomold (see Table of Materials) immediately after the dissected brain is embedded in the cryomold with the embedding compound.
   NOTE: Liquid nitrogen can also be used to cool the aluminum block. In that case, the aluminum block should be placed in a Styrofoam box filled with enough liquid nitrogen to cover the aluminum block just before embedding. After confirming the liquid nitrogen sublimation, place the cryomold on the precooled aluminum block. It is better to confirm how many cryomolds can be placed on the aluminum block at once. It is advisable to use liquid nitrogen or an additional precooled block to ensure enough space to cool all samples quickly.
4. Put the precooled aluminum block in a Styrofoam box. Use a Styrofoam box with a lid to prevent the aluminum block from warming.
5. For coronal sections, embed the brain in a cryomold and adjust the orientation using forceps under the microscope (Figure 3A).
6. Place the cryomold on the precooled aluminum block in the Styrofoam box and allow the OCT compound to freeze (Figure 3B). When the OCT compound starts to freeze, it becomes white.
7. Apply a small circle of OCT compound on a specimen disc, and mount the cryoblock to attach the cryoblock to the specimen disc.
8. Place the specimen disc in the cryostat and freeze the OCT compound completely.
9. Place the specimen disc on the specimen head in the cryostat.
10. Set the orientation of the cryoblock and trim the OCT to remove any extra regions.
    NOTE: Adjust the orientation of the cutting plane while cutting the telencephalon. The blade is equipped with a cryostat. It is possible to adjust the orientation of the cutting plane by changing the angle of the specimen head.
11. Cut 14 µm thick serial sections through the whole optic tectum using a cryostat. Store the slides at -25 °C.
    ​NOTE: Long-term storage should be at -80 °C.

4. Fluorescent immunostaining

1. Dry glass slides in a slide rack for 30 min at room temperature (RT). Wash slides in 1x PBS for 30 min at RT. If immunostaining with anti-PCNA or BrdU antibodies is to be performed, proceed to step 4.2 or step 4.3.
2. Prepare 500 mL of 10 mM sodium citrate in a 500 mL beaker and warm the citrate buffer to 85 °C on a hot plate magnetic stirrer, then place the slides in a slide staining rack and incubate the rack in the citrate buffer at 85 °C for 30 min. To avoid boiling, keep the temperature around 85 °C. After antigen retrieval, wash the slides three times in 0.1 % Triton in 1x PBS (PBSTr), with each wash taking 5 min at RT.
   NOTE: This step is an optional step for proliferating cell nuclear antigen (PCNA) retrieval.
3. Prepare a 2 N HCl solution by diluting 12 N HCl in distilled water. Using a liquid blocker (see Table of Materials), draw a line as a hydrophobic barrier to keep the 2 N HCl solution around the sections.
   1. Place the slides on immunostaining trays and apply 200-300 µL of the 2N HCl solution. Incubate the trays at 37 °C for 30 min. After antigen retrieval, wash three times in 0.1 % PBSTr with each wash taking 5 min at RT.
      NOTE: This step is an optional step for BrdU antigen retrieval.
4. Absorb and remove the remaining solution using paper towels. Then, use a liquid blocker to draw a hydrophobic barrier to keep the antibody solution around the sections as necessary.
5. Place the slides on trays and apply 200-300 µL of blocking solution, 3% (v/v) horse serum in PBSTr) to every slide, and incubate the slides for 1 h at RT.
6. Wash the sections with PBSTr for 5 min at RT and repeat three times.
7. Prepare the primary antibodies (see Table of Materials) using the blocking solution (200-300 µL of antibody solution for each slide). Remove the remaining PBSTr with paper towels and place the slides on the immunostaining trays. Apply the primary antibody solution to every slide and incubate trays overnight at 4 °C.
8. Wash the sections with PBSTr for 5 min at RT and repeat three times.
9. Prepare fluorescent secondary antibody solution (see Table of Materials) using the blocking buffer (1:500). Remove the remaining PBSTr with paper towels and place the slides on the immunostaining trays. Apply the secondary antibody solution to every slide and shade trays with aluminum foil. Incubate for 1-2 h at RT.
10. Wash the sections with PBSTr for 5 min at RT three times without exposing them to light.
11. Prepare Hoechst solution (1:500 dilution in 1x PBS, see Table of Materials) for nuclear staining. Remove the remaining PBSTr with paper towels and place the slides on the immunostaining trays. Apply the Hoechst solution to every slide and cover the trays with aluminum foil. Incubate for 30 min at RT.
12. Wash the sections with 1x PBS for 10 min at RT without exposing them to light.
13. Absorb and remove the remaining 1x PBS using paper towels and mount the sections on slides using a water-soluble mounting medium (see Table of Materials) for fluorescent immunostaining. Slowly set a 24 x 45 mm coverslip onto the sections. Store at 4 °C, without exposing the slides to light.
14. Observe and image the sections under fluorescent microscopy or confocal microscopy.