Three Strategies to Induce Neurotrophic Keratitis and Nerve Regeneration in Murine Cornea

Here, we propose three different methods of damaging the sensory fibres innervating the cornea. These methods facilitate the study of axon regeneration in mice. These three methods, which are adaptable to other animal models, are ideal for the study of corneal innervation physiology and regeneration.

The cornea, which is the transparent tissue covering the eye, requires a balanced microenvironment to maintain its integrity. The dense innervation of the cornea participates to this microenvironment. If there is a loss, which could be partial or total, of this innervation, then a specific condition arises, which is called neurotrophic keratitis.

To find new cures and efficient cures for this disease, we need to have specific models. The expansion of the use of transgenic animal bearing fluorescent reporters and microscopic advances have opened new possibilities in visualizing axonary generation. In vivo imaging has become more accessible to any research lab willing to follow the nerve reneration process.

The various causes inducing neurotrophic keratitis underlying the difficulties of having only one model to test nerve regeneration therapies. These three models are reproducing various situations seen in clinics, and sharing those methods will allow other labs to uncover new aspects of corneal innervation from which innovative treatments will be designed. To begin corneal abrasion, place an anesthetized mouse on a table.

Apply a drop of artificial tear on the eye undergoing the surgery. Apply a drop of ocular gel on the contralateral eye. Next, position the mouse on its side on a heated plate.

Gently use a cotton swab to absorb the artificial tear and brush aside the mouse's eyelashes without making contact with the eye. Turn on the ocular burr and carefully open the mouse's eyelid with two fingers. Simultaneously shield the vibrissae to prevent them from entangling in the burr.

Locate the pupil or the eye center and gently apply the ocular burr to the eye's surface in circular motions until the epithelium is removed. Alternatively, perform about 20 circular movements on the cornea. Now apply a drop of fluorescent staining solution on the abraded eye.

After 20 seconds, dab the eye with a tissue to absorb the solution, then rinse it with a drop of artificial tear. Finally, absorb the artificial tear with a tissue and illuminate the eye using a blue cobalt lamp. Prior to the surgery, the basal epithelium was intact and the nerve fibers in the subbasal plexus formed a vortex.

The burr tip removed the outer corneal layer, leaving the stroma and endothelium intact, with a visible gap at the basal epithelium and subbasal plexus. Regeneration of the epithelium occurred over a week post-surgery, however, the vortex remained absent from the corneal center. To begin corneal nerve axotomy, place an anesthetized mouse on a table.

Apply a drop of artificial tear on the eye undergoing the surgery. Apply a drop of ocular gel on the contralateral eye. Position the mouse under a binocular loupe to assess the eye that is being operated on.

Place the bottom of a Petri dish under the mouse's head to keep the eye horizontal. With a cotton swab, carefully dab the artificial tear from the eye and remove the eyelashes without touching the eye. Place a pair of smooth curved pillars under the animal's eye and gently pop it out of the orbit.

Gently place a biopsy punch vertically against the cornea's surface, apply pressure, and twist the punch several times. Then remove the biopsy punch, and a circular mark should be visible on the cornea. A thin peripheral ring was seen at the point where the biopsy punch was applied, which damaged the basal epithelium.

The peripheral nerves appeared to be sectioned, triggering the wallerian axon degeneration. Heavy degeneration was observed, leading to total loss of innervation in the cornea center. This resulted in the disintegration of the epithelial layer, creating an ulcer.

To begin, place an anesthetized mouse on a table. Apply a drop of ocular gel on both eyes. Place the animal in a head holder to secure its position.

Rotate the animal by 90 degrees so that the eye being operated on faces the ceiling. Secure the animal's vibrissae to clear the area around the eye. Secure the animal's body to minimize head movements caused by breathing.

Place a circular cover slip horizontally over the eye on top of the ocular gel. Adjust the animal's head as necessary to correctly position the cover slip. Add a drop of PBS on top of the cover slip.

Then carefully place the animal onto a microscope stage. Adjust the objective turret of the microscope to the appropriate distance for observation. Activate the lasers to induce corneal nerve ablation.

Once nerve ablation is complete, remove the animal from the head holder and place it in a cage on a heated plate. The vortex was visibly absent one week after laser ablation. A group of immune cells were observed to have migrated to the illumination site.

The stromal fiber bundles appeared to have thinned out.