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

So my lab is interested in understanding how the eye and brain grow together. This technique allows us to study how retinal input influences brain growth and development. Surgical removal of one eye from the living larval zebrafish, followed by observation of the optic tectum allows us to compare innervated and denervated tectal lobes within the same animal and across animals.

Combining this technique with modern molecular approaches can give new insight into mechanisms underlying neural development, regeneration, and degeneration. To begin, connect the cathode and anode wires to the power supply. Attach the alligator clip at the end of the cathode wire to a partially straighten paperclip and insert the paperclip into the potassium hydroxide solution, attaching it to the side of the jar.

Attach the alligator clip of the anode wire to the neck of the needle holder. Turn the power to approximately 20 volts and dip the tungsten wire into the potassium hydroxide solution, pulling the wire out of the solution at an angle to electrolytically sharpen the wire into a fine tip. Check the tip of the needle under the dissecting microscope to ensure that it is sharp enough.

The curve needle tip results when the sharp needle is bent by gently touching it to the Petri dish. On the day of surgery, use a wide-bore glass pasture pipette to transfer 10 to 15 larvae to a 35 millimeter Petri dish filled with fresh E3.Add three to five drops of anesthetic to the larva. To determine if the larvae are adequately anesthetized, look for lack of touch response.

Add two to three more drops of the anesthetic if the larvae are still responsive to touch after three minutes and reassess. To immobilize the larva for surgery, take one larva up into a narrow-bore glass pasture pipette with only a small amount of E3.Next, take up approximately 200 microliters of melted 1%low melting point or LMP agarose into the pipette containing the larva and mix to ensure that the larva is suspended in the LMP agarose. Place the lid of a 35 millimeter Petri dish faced up under the dissecting microscope, then squirt the larva and the agarose onto the upside down Petri dish lid.

Spread the agarose so that the larva is in the center of the drop. Using a dull tungsten needle, quickly but gently maneuver the larva so that it is lateral with one eye facing upwards. Wait several minutes for the agarose to set.

Following the edge of the eye orbit, use the tip of a fine electrolytically sharpened tungsten needle to pierce the skin around the eye. Then slide the edge of the needle under the eye from the temporal-ventral side of the eye. Use controlled pressure to release the eye from the socket.

Keep pressing the eye dorsally and anteriorly with the side of the needle, eventually slicing through the optic nerve and releasing the eye. Alternatively, you can use very fine surgical forceps to remove the eye by pinching the optic nerve and pushing the eye from medial to lateral. After successful eye removal, cover the agarose with MMR solution, 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.

After the surgeries, place the larva in MMR solution supplemented with antibiotics until the following day, then return the larva to E3 and rear them until the endpoint of the experiment. After larva are terminally anesthetized, fix them with 4%paraformaldehyde. Suspend the fixed larvae in PBS droplets on a sealguard plate under a dissecting microscope.

Secure them laterally by placing two tungsten pins through the notochord with one pin posterior to the pigmented area covering the AGM region and another in line with the end of the yolk extension. Remove the eye with a sharp tungsten needle and fine forceps. Sometimes it is possible to poke the needle through the jaw to the opposite side and remove the other eye.

Use the tungsten needle to scratch from the temporal to the ventral side of the ear. In the same action in motion, bring the needle posterior to the jaw and gently pull anteriorly until the ear and jaw are removed. Remove the other eye and ear by poking the needle through the jaw.

Use forceps to pull out the ventral organs and remaining yolk. Finally, make a shallow incision in the dorsal cranial skin near the junction between the hind brain and spinal cord. Lift the skin with the forceps and pull it anteriorly and around the telencephalon.

Remove any remaining tissue with forceps. Unpin the larva and transfer to PBS into 1.5 milliliter microcentrifuge tube. Secure a chambered slide into the lid of a 100 millimeter Petri dish with vacuum grease.

Transfer the immunostained and processed larva to a well plate or depression slide to view them with a dissecting microscope. Mount the larva on the chambered slide in 1%LMP agarose columns. Using a glass past your pipette, put one larva on the chambered slide, depositing as little PBS as possible.

Cover the larva with melted and warm 1%LMP agarose using the same pipette. Pipette the LMP agarose into a column and then position the larva as symmetrically as possible with the dorsal surface visible. After eye removal, progressive degeneration of retinal axons was observed in the optic tectum neuropil.

By two days postsurgery, RFP labeled axons exhibited hallmarks of rapid Wallerian degeneration, such as blabbing and fragmentation. RFP positive puncta were noted both within and outside the neuropil on the right side of the tectum as indicated by the arrows. By four days post-surgery, fragmented axons and RFP labeled axonal debris were considerably reduced in the right tectal lobe, indicating relatively rapid clearance of the dying and degenerating axons.

The brain was exposed by dissecting the eyes, jaw, ears, and skullcap of the skin and connective tissues. Ideally, the brain remains intact during this procedure. However, parts of the fore brain, particularly the olfactory bulb, were likely damaged or completely removed when the skull cap of the skin or jaw was removed.

Moreover, sometimes the lateral ledge of the tectum gets sliced when piercing or pinching the skin to pull it off the brain. No two samples are identical. So you need to be able to make subtle adaptations through the eye removal and brain dissection as needed.

Patients and sharp tools are essential for success. This technique can be followed up with cellular and molecular approaches, like in vivo live cell imaging or RNA sequencing to gain new insights into the molecular and cellular mechanisms of optic tectum growth and development.