Eye Removal in Living Zebrafish Larvae to Examine Innervation-dependent Growth and Development of the Visual System

The methods in this paper were conducted in accordance with guidelines and approval of the Institutional Animal Care and Use Committees of Reed College and University College London. See the Table of Materials for details about zebrafish strains used in this study.

1. Prepare materials and tools

1. Make solutions.
   1. Make embryo medium (E3¹⁴) by diluting a 60x stock (300 mM NaCl, 10.2 mM KCl, 20 mM CaCl₂-dihydrate, and 20 mM MgCl₂-hexahydrate in deionized water, autoclaved, and stored at room temperature). Add 160 mL of 60x E3 to 10 L of deionized water. Store at room temperature.
   2. Make 1% low melting point (LMP) agarose in E3. Dissolve 1 g of LMP agarose into 100 mL of E3 by boiling in the microwave for 1-2 min on high power. Aliquot molten agarose into 1.7 mL microfuge tubes, and store at 4 °C. Boil in a water bath and hold in a 40 °C heat block when ready to use for embedding.
   3. Make Marc's Modified Ringers (MMR¹⁵) isotonic solution by diluting a 10x stock (1 M NaCl, 20 mM KCl, 10 mM MgSO₄, 20 mM CaCl₂-dihydrate, 50 mM HEPES, pH adjusted to 7.5 with 10 M NaOH, then autoclaved, and stored at room temperature). Add 10 mL of 10x MMR to 90 mL of deionized water.
2. Pour Sylgard plates according to the manufacturer's instructions, allowing them to set and dry for several days¹⁶.
3. Electrolytically sharpen tungsten needles (protocol modified from ¹⁷).
   1. First, prepare a 10% (w/v) KOH solution by adding 200 mL of deionized water to a wide-mouthed glass jar with a screw-top lid. Slowly stir in 20 g of KOH pellets.
      NOTE: This caustic solution is highly exothermic. Wear gloves and eye protection, and stir the solution in the hood.
   2. Cut a 2-3 cm length of tungsten wire and insert it into the needle holder.
   3. Connect the cathode and anode wires to the power supply. Attach the alligator clip at the end of the cathode wire to a partially straightened paper clip and insert the paper clip into the KOH solution, attaching it to the side of the jar.
   4. Next, attach the alligator clip of the anode wire to the neck of the needle holder. Turn the power on (to ~20 V) and dip the tungsten wire into the KOH solution, pulling the wire out of the solution at an angle to electrolytically sharpen the wire into a fine tip.
      NOTE: Bubbles will come from the paper clip as the reaction proceeds.
   5. Check the tip of the needle under the dissecting microscope to ensure that it is sharp enough.
   6. Rinse the needle with deionized water to remove all KOH residue. Make several needles ahead of time and keep them until the larval surgeries and dissections.
4. Make chambered slides for imaging zebrafish specimens with an upright confocal microscope.
   1. Obtain glass microscope slides and two-part epoxy resin (see the Table of Materials). Mix the epoxy according to the instructions on the package, and paint thick rings or rectangles on the glass slides. Allow the slides to cure and dry in the hood for at least 24 h before use.

2. Embryo collection and rearing

1. Pair adult male and female fish of the desired genotypes in breeding boxes in the late afternoon/early evening. The next morning, after they have spawned, collect newly fertilized eggs with a mesh strainer and transfer them to E3 in 100 mm Petri dishes.
2. Incubate the embryos at ~27.5 °C with a 14 h light/10 h dark cycle until the day of the surgery. Change E3 daily or as needed.
3. Feed the larvae using a dropper full of harvested rotifers once a day, beginning either at 5 days post fertilization (dpf) or one day after surgery. After the larvae have several hours to eat, change the E3 to remove any remaining rotifers and waste products.
   ​NOTE: Do not feed larvae on the day of surgery.

3. Prepare larvae for surgery

1. On the day of surgery, use a wide-bore, glass Pasteur pipette to transfer 10-15 larvae to a 35 mm Petri dish filled with fresh E3.
2. Anesthetize the larvae by adding 3-5 drops of 0.4% w/v Tricaine solution (0.4 g Tricaine dissolved in 210 mM Tris, pH 9; final solution adjusted to a pH of 7.4 if necessary¹⁸) to the dish for a final concentration of ~0.0015% w/v Tricaine. Look for lack of response to touch to determine if the larvae are adequately anesthetized. If they are still responsive to touch after ~3 min, add 2-3 more drops of 0.4% w/v Tricaine solution and reassess.
   NOTE: At this stage of development, it is possible to monitor blood flow and heart rate in addition to motility under the dissecting microscope.
3. Immobilize the anesthetized zebrafish larvae by embedding them in 1% LMP agarose dissolved in E3.
   1. First, place the lid of a 35 mm Petri dish face up under the dissecting microscope. Next, take one larva up into a narrow-bore, glass Pasteur pipette with only a small amount of E3.
   2. Then, aspirate ~200 µL of melted, warm (~40 °C) 1% LMP agarose into the pipette with the larva. Finally, squirt the larva and agarose onto the upside-down Petri dish lid.
   3. Use a dull tungsten needle to quickly but gently maneuver the larva so that it is lateral, with one eye facing up. Wait for the agarose to set (~5 min).
      ​NOTE: If the agarose hardens while the larva is not lateral, free it from the agarose (as described in step 5) and return it to the dish with E3 and tricaine.

4. Eye-removal

1. Once the agarose is set, and the larva is positioned laterally, use the tip of a very fine, electrolytically sharpened tungsten needle to pierce the skin around the eye, following the edge of the eye orbit.
2. Next, slide the edge of the needle (not the tip) under the eye from the temporal-ventral side of the eye. Use controlled pressure to release the eye from the socket.
3. Use very fine surgical forceps to remove the eye by pinching the optic nerve and pushing the eye from medial to lateral. Alternatively, simply keep pressing the eye dorsally and anteriorly with the side of the needle, eventually slicing through the optic nerve and releasing the eye.
   ​NOTE: If tissues other than the eye are accidentally stabbed during the surgery, quickly and humanely kill the larva by rapid decapitation.

5. Postoperative care until experimental endpoint

1. After successful eye removal, cover the agarose with MMR solution.
2. Liberate each larva from the agarose by gently brushing a tungsten needle around their head and then around their body while stabilizing the Petri dish lid with forceps.
3. Transfer the one-eyed larvae to a 35 mm Petri dish that contains MMR supplemented with a 1:100 dilution of a penicillin/streptomycin cocktail. Incubate the embryos at ~27.5 °C with a 14 h light/10 h dark cycle.
   ​NOTE: This isotonic solution of MMR supplemented with antibiotics initially aids larval healing and survival. Each dish can hold up to 15 recovering larvae.
4. The next day, return the larvae to E3. Starting the afternoon following surgery, feed larvae (~6 dpf and older) using a dropper full of harvested rotifers once a day. After the larvae have had several hours to eat, change the E3 to remove any remaining rotifers and waste products.

6. Fixing larvae

1. Once the larvae have reached the desired developmental stage for analysis, anesthetize them with a lethal dose of tricaine (~0.4% final concentration) until their hearts have stopped.
2. Pipette up to 30 larvae per 1.5 mL microfuge tube. Remove excess E3 and then add 0.5-1 mL of fixative solution (4% paraformaldehyde (PFA) and 4% sucrose in phosphate-buffered saline (PBS)) to fix the tissue. Incubate the larvae overnight at 4 °C.
3. Wash the larvae with PBS the next day by removing the fixative and rinsing the larvae 3-4 times with PBS. Store the larvae at 4 °C for up to 7 days before dissection.

7. Dissecting larvae to reveal brains (adapted from ¹⁵)

1. Suspend the fixed larvae in PBS droplets on a Sylgard plate under a dissecting microscope. Secure them laterally by placing two tungsten pins (typically old tungsten needles that are less than 2 cm) through the notochord, with one pin just posterior to the pigmented area covering the aorta-gonad-mesonephros (AGM) region and another in line with the end of the yolk extension.
   NOTE: Position the tungsten pins in as few attempts as possible because the tissue will become more friable with each puncture.
2. Use a very sharp tungsten needle and needle-sharp forceps to expose the brain by removing, in this order, the eye(s), ear, jaw, digestive tract, and dorsal cranial skin.
   1. First, remove the eye using a technique similar to the surgical eye-removal in step 4. Then, unpin the larva and flip it to the opposite side, so the other eye is accessible and repeat. Alternatively, poke the needle through the jaw to the other side and remove the eye without unpinning.
      NOTE: Eyes may be collected at this point if analysis of this tissue is desired (similar to ¹⁹).
   2. Next, use the tungsten needle to scratch from the temporal to the ventral side of the ear. In the same action/motion, bring the needle posterior to the jaw and gently pull anteriorly until the ear and jaw are removed.
   3. Use forceps to pull the ventral organs and any remaining yolk out.
   4. Finally, make a shallow incision in the dorsal cranial skin near the junction between the hindbrain and spinal cord. Lift the skin with the forceps and pull it anteriorly and around the telencephalon. If this proves too difficult, start at the initial incision in the hindbrain and pull the skin first laterally and then ventrally and anteriorly, taking great care not to scratch the lateral edges of the tectum.
      NOTE: Because the midbrain is the object of study, the hindbrain can be cut; however, a deep incision may result in decapitation.
   5. Remove any remaining tissue with forceps. Unpin the larva and transfer to PBS in a 1.5 mL microfuge tube.
      ​NOTE: Leave the tail attached to the brain for better visibility when performing wholemount protocols.

8. Brain dehydration and storage

1. Remove PBS from the brains and add 1 mL of PBS containing 0.1% TritonX-100 (PBST). Incubate for 5-10 min at room temperature.
2. Remove the PBST and replace it with a 50:50 methanol:PBST solution. Incubate for 5-10 min at room temperature.
3. Wash the brains with 100% methanol (MeOH) twice for 5 min each at room temperature.
4. Store the brains in 100% MeOH at -20 °C for at least 16 h before performing immunohistochemistry or other wholemount procedures.
   ​NOTE: Brains can be stored in MeOH for up to 2 years at -20 °C.

9. Immunohistochemistry

NOTE: Established protocols for many useful wholemount techniques in zebrafish can be found on ZFIN²⁰. This manuscript provides examples comparing one-eyed and two-eyed larvae that were immunostained with antibodies that recognize either the red fluorescent protein (RFP), which is expressed in optic nerve axons in the Tg[atoh7:RFP] line (Figure 2), or phosphorylated histone H3 (PH3), which highlights mitotic cells (Figure 3). A standard immunohistochemistry protocol for wholemount embryos and larvae is summarized below.

1. First, rehydrate the larval brains with a graded series of MeOH:PBST washes at room temperature by incubating the specimens in 50:50 MeOH:PBST for 5 min, then in 30:70 MeOH:PBST for 5 min, and ending with 3 washes of PBST.
2. To permeabilize the brains, incubate them in PBST supplemented with proteinase K (PK) at a final concentration of 20 µg/ml PK for 30 min at 37 °C. Store aliquots of 10 mg/mL PK at -20 °C and thaw them before use.
3. Remove the PK solution and wash away any remaining enzyme by rinsing the brains with PBST 3 times over 15 min.
4. Refix the permeabilized brains by incubating them in 4% PFA for 20 min at 25 °C (or room temperature). Then, remove PFA and wash away any remaining fixative by rinsing the brains 3 times over 15 min with PBST.
5. Replace the final PBST rinse with freshly made immunoblocking (IB) buffer (10% normal goat serum and 1% DMSO in PBST) and incubate at room temperature for at least 1 h.
6. Dilute primary antibodies into IB buffer at the appropriate concentration (e.g., 1:500 for RFP and 1:300 for PH3 antibodies).
7. Remove IB buffer from brains and replace it with the primary antibody dilution. Incubate overnight at 4 °C on an orbital shaker with gentle rocking. Ensure tubes are securely resting on their sides.
8. The next morning, remove the primary antibody solution with a micropipette, rinse a couple of times in PBST, and then wash the brains in PBST at room temperature for 4 x 30 min.
   NOTE: Primary antibody dilutions can be reused once within ~7 days if stored at 4 °C.
9. Rinse the brains in IB buffer while diluting secondary antibodies into IB buffer at the appropriate concentration (e.g., 1:500 for Alexa-fluor 568 anti-rabbit).
10. Remove the IB buffer and replace it with the secondary antibody dilution. Incubate overnight at 4 °C on an orbital shaker with gentle rocking of the tubes resting on their sides.
11. The next morning, remove the secondary antibody solution, rinse with PBST, and then wash brains in PBST at room temperature for 4 x 30 min.
12. Counterstain with 4',6-diamidino-2-phenylindole (DAPI) or ToPro3 (1:5,000 in PBST at 4 °C overnight).
13. The next morning, transfer the larval brains into PBS and proceed to the mounting and imaging steps listed below.

10. Mounting and imaging

1. Secure a chambered slide into the lid of a 100 mm Petri dish with vacuum grease.
2. Transfer the immunostained and processed larvae to a well plate or depression slide to view them with a dissecting microscope.
3. Mount the larvae on the chambered slide in columns of 1% LMP agarose.
   1. Using a glass Pasteur pipette, put one larva on the chambered slide, depositing as little PBS as possible. With the same pipette, cover the larva with molten 1% LMP agarose (~40 °C).
   2. With a blunt tungsten needle or forceps, draw the agarose into a column and then position the larva as symmetrically as possible, with the dorsal surface visible.
   3. Repeat this procedure for the remaining larvae.
   4. Once the agarose has gelled, cover it with a few drops of PBS and then place it on the stage of the confocal microscope.
4. Align the objective lens with the sample, focus the microscope using transmitted light, and illuminate the sample with the appropriate lasers.
   NOTE: In this example, 405 nm and 568 nm wavelengths were used to excite DAPI and RFP, respectively.
5. Adjust the gain, offset, and laser power by moving the slider bars to an appropriate level relative to the signal and the used detector. Check the histograms for each channel and click the State of saturation indicator button above the image to gauge the level of exposure, ensuring that none of the pixels are saturated and that the signal-to-noise ratio is high. For an example of optimum confocal imaging parameters for cellular and subcellular resolution within zebrafish, see ²¹.
6. Set the upper and lower limits of the stack of z-planes by using the scrollwheel on the mouse to find the top and bottom of the sample. Click the top and bottom limits for image acquisition and then trigger the z-stack capture by clicking the Run now button.
   NOTE: Depending on how symmetrically the brain is mounted, the total z-depth for a 9 dpf larval zebrafish brain will be ~150-200 µm.
7. Process and colorize the images with colorblind accessibility in mind (e.g., use blue and orange or magenta and green for two-color images). Set lookup tables in the software used to capture the images or image processing software such as Photoshop from Adobe.
8. In Photoshop, color raw photomicrographs saved as 8 bit Tagged Image File Format (TIFF) by (i) converting the Image Mode to RGB and (ii) accessing the Curves option under the Image | Adjustments menus and then (iii) sliding the appropriate colored light outputs to 0 or 255 levels to achieve the desired color. For example, to achieve a green signal, select the Red channel and slide its output to 0; then, select the Blue channel and slide its output to 0 too. Similarly, to achieve a magenta signal, select the Green channel and slide its output to 0; leave the red and blue channels as they are.
   NOTE: Similar steps can be performed in the free Gnu Image Manipulation Program (GIMP).
9. Manually quantify the numbers of cells displaying a specific marker using the cell counter plugin in ImageJ²², available under Analyze in the Plugins menu. Examples of how to use ImageJ for cell counting can be found in many tutorials, including ²³.
10. Generate graphical representations of data in statistical software such as RStudio (see supplemental data for .Rmd file).