The authors thank Dr. Karine Loulier for the access to the transgenic mouse line MAGIC-Markers. The authors also thank the RAM-Neuro animal core facility and the imaging facility MRI, a member of the France-BioImaging national infrastructure supported by the French National Research Agency (ANR-10-INBS-04, &#34;Investments for the future&#34;). This research was supported by the ATIP-Avenir program, Inserm, R&#233;gion Occitanie, the University of Montpellier, the French National Research Agency (ANR-21-CE17-0061), the Fondation pour la Recherche M&#233;dicale (FRM Regenerative Medicine, REP202110014140), and the Groupama Foundation.

All experiments were approved by the National Animal Experiment Board.
1. Preparations
<ol>
	<li>Prepare an anesthetic solution of ketamine-xylazine for anesthesia. Inject ketamine at 80 mg/kg and xylazine at 10 mg/kg by diluting 200 &#181;L of ketamine (100 mg/mL) and 125 &#181;L of xylazine (20 mg/mL) in 2,175 mL of sterile 0.9% NaCl.</li>
	<li>Prepare 0.02 mg/mL buprenorphine solution as an analgesic solution by adding 100 &#181;L of 0.3 mg/mL buprenorphine to 1,400 mL of sterile 0.9% NaCl.</li>
	<li>Prepare the fluorescent staining solution.
	<ol>
		<li>Use a fine scale to weigh 10 mg of fluorescein salt and dilute it in 10 mL of phosphate-buffered saline (PBS) to obtain a 0.1% fluorescein solution.</li>
		<li>Protect the solution from the light and shake it for 5 min. Use a 10 mL syringe and a syringe filter of 0.2 &#181;m to filter the solution. 
		NOTE: The fluorescent solution has to be protected from the light and can be stored at +4 &#176;C for 5 days.</li>
	</ol>
	</li>
	<li>Preparation of the mouse
	<ol>
		<li>Weight the mouse and induce anesthesia by performing an intraperitoneal injection of 10 &#181;L/g of mouse weight (MW) of the anesthetic solution. When the mouse stops moving, place it on a heated plate (37 &#176;C) and verify that it is completely anesthetized by pinching its toes.</li>
		<li>Apply a drop of artificial tear on the eye undergoing the surgery and a drop of ocular gel on the contralateral eye.</li>
		<li>Once the mouse is unresponsive to pinch, inject the analgesia (0.02 mg/mL buprenorphine) at 5 &#181;L/g of MW to provide 0.1 mg/kg buprenorphine subcutaneously in the neck.</li>
	</ol>
	</li>
</ol>
2. Abrasion of the cornea
<ol>
	<li>Follow step 1.3 to prepare the fluorescent staining solution.</li>
	<li>Follow step 1.4 to prepare the mouse.</li>
	<li>Before doing the abrasion, dip the ocular burr in 70% EtOH and then in PBS to clean it.</li>
	<li>Place the mouse on a heated plate (37 &#176;C) on the side to easily access the eye undergoing the surgery.
	<ol>
		<li>Use a cotton swab to absorb the artificial tear and brush the eyelashes away without touching the eye.</li>
		<li>Turn on the ocular burr, open the eyelid of the mouse with two fingers, and simultaneously block the vibrissae to avoid them getting stuck into the burr.</li>
	</ol>
	</li>
	<li>Localize the pupil or the center of the eye and apply the ocular burr on the surface of the eye, and do circular movements. By looking closely, the surface of the removed epithelium can be observed. Otherwise, do approximately 20 circular movements on the cornea.</li>
	<li>Put the mouse back on its belly and check if the ocular gel is still present in the contralateral eye.</li>
	<li>Apply a drop of fluorescent staining solution on the abraded eye and wait for 20 s. Absorb the drop with a tissue without touching the eye and rinse it with a drop of artificial tear. Absorb the drop of the artificial tear with a tissue and illuminate the eye with the blue cobalt lamp. 
	NOTE: The abrasion has to have a circular shape. It requires some training to obtain it.</li>
	<li>Apply a drop of ocular gel on the abraded eye and let the mouse wake up on the heated pad. When the mouse is moving on its own, put it back in its cage.</li>
	<li>Check the well-being of the animal during the following 2 days.</li>
	<li>After the use of the ocular burr, use a soft tissue to remove epithelial cells stuck into the head of the burr and dip it in 70% EtOH and then PBS. 
	&#8203;NOTE: Once the epithelial cells are removed with the soft tissue, make sure that no pieces of tissue are stuck in the head of the burr.</li>
</ol>
3. Axotomy of the corneal nerves
<ol>
	<li>Follow step 1.4 to prepare the mouse.</li>
	<li>Place the mouse under a binocular loupe on the side to access the eye undergoing the surgery. Put the bottom of a Petri dish under the head of the mouse for the eye to be horizontal. 
	NOTE: The following steps will be performed by looking through the binocular loupe.</li>
	<li>Remove the artificial tear with a cotton swab and remove the eyelashes without touching the eye.</li>
	<li>Place smooth curved pliers under the eye of the animal in order to pop it out from the orbit. Make sure to close the pliers enough for the eye not to be able to go back in the orbit once pressure is applied, but not excessively to prevent optic nerve damage.</li>
	<li>Apply the biopsy punch of 2.5 mm vertically on the eye without pressure to ensure total contact of the punch with the surface of the cornea. 
	NOTE: Once the punch is applied against the cornea, the hand of the experimenter might interfere with the field of view.</li>
	<li>Then, start applying pressure and twist the punch several times. 
	NOTE: Depending on the pressure, the number of twists may vary, but generally it is around 5. It requires training to feel the right pressure and movement. If too much pressure is applied, the eye ruptures. In this case, liquid can be observed exiting the lesion and the eye get soften. The animal has to be sacrificed.</li>
	<li>Remove the biopsy punch. A circle must be visible on the cornea where the biopsy punch was applied. 
	NOTE: The next steps do not require the binocular loupe.</li>
	<li>Check if the contralateral eye still has ocular gel and apply some on the axotomized eye.</li>
	<li>Let the mouse wake up on a heated pad (37 &#176;C) and put it back in its cage when it is moving on its own.</li>
	<li>Check the well-being of the animal during the following 2 days.</li>
</ol>
4. Cornea whole-mount processing
<ol>
	<li>Sample collection and fixation.
	<ol>
		<li>Weigh&#160;the mouse and induce anesthesia by performing an intraperitoneal injection of 10 &#181;L/g of mouse weight (MW) of the anesthetic solution.</li>
		<li>When the mouse stops moving and does not have reflexes by toe pinching, perform a cervical dislocation.</li>
		<li>Pop the eye out of the orbit using two fingers and enucleate the eye by cutting the optic nerve using curved scissors.</li>
		<li>Place the eye in a 2 mL plastic tube containing PBS.</li>
		<li>Fix the eye by replacing the PBS with 4% paraformaldehyde and placing it on a shaker (30 rpm) at room temperature (RT) for 20 min.</li>
		<li>Rinse 3 times for 10 min with PBS at RT on a shaker (30 rpm).</li>
	</ol>
	</li>
	<li>Dissection of the cornea
	<ol>
		<li>Place a drop of PBS on a piece of parafilm inside a Petri dish.</li>
		<li>Place the eye in this&#160;drop of PBS.</li>
		<li>Place the Petri dish under the binocular loupe.</li>
		<li>Dissect the cornea using microdissection scissors and forceps. Cut above the ciliary body to keep the limbus.</li>
		<li>Remove the lens, the ciliary body, and the iris with fine forceps.</li>
		<li>Place the cornea back into a 2 mL plastic tube.</li>
	</ol>
	</li>
	<li>Immunofluorescence protocol
	<ol>
		<li>Incubate the cornea in 2 mL of 2.5% fish skin gelatine, 5% goat serum, and 0.5% detergent in PBS for 1 h at RT on a shaker (30 rpm).</li>
		<li>Incubate the cornea in 150 &#181;L of a solution of 2.5% fish skin gelatine, 5% goat serum, and 0.1% detergent in PBS containing the primary antibody diluted at 1/1000 (anti &#946;III tubulin antibody) overnight at 4 &#176;C on a shaker (30 rpm).</li>
		<li>Rinse 3 times for 1 h with 0.1% detergent in PBS at RT on a shaker (30 rpm).</li>
		<li>Incubate the cornea in the same solution as for step 4.3.2, containing the secondary antibody diluted at 1/500 overnight at 4 &#176;C on a shaker (30 rpm).</li>
		<li>Rinse 3 times for 1 h with PBS at RT on a shaker (30 rpm).</li>
		<li>Incubate the cornea for 10 min with 150 &#181;L of DNA intercalator diluted at 1/5000 at RT on a shaker (30 rpm).</li>
		<li>Rinse 2 times for 5 min with PBS.</li>
	</ol>
	</li>
	<li>Mounting of the cornea
	<ol>
		<li>Place the cornea on a slide, epithelium facing the slide, and use a scalpel to create four sections without reaching the center. 
		NOTE: Do the sections opposite to each other to form a flower shape.</li>
		<li>Place the cornea endothelium facing the slide and remove the PBS around the cornea using a tissue. 
		NOTE: Do not remove the PBS under the cornea; otherwise, air bubbles may appear. If so, add PBS to the cornea and remove the air bubble. Then restart from step 4.3.2.</li>
		<li>Add model paste on the four corners of a rectangular coverslip. 
		NOTE: The model paste elevates the coverslip as the cornea is a thick tissue.</li>
		<li>Drop 50 &#181;L of a mounting medium on the cornea and lay carefully the coverslip above to avoid bubble formation.</li>
		<li>Seal the coverslip with nail polish.</li>
	</ol>
	</li>
	<li>Imaging
	<ol>
		<li>Place the slide under an epifluorescent microscope.</li>
		<li>Define the size of the mosaic and the depth of the image and start the acquisition.</li>
		<li>Apply a deblurring and deconvolution program on the acquired image. 
		&#8203;NOTE: Step 4.6.3 is an option selected on the acquisition software (Large Volume Computational Clearing [LVCC])</li>
	</ol>
	</li>
</ol>
5. Localized laser ablation of corneal nerves
<ol>
	<li>Turn on the upright multiphoton microscope and the laser combined with an optical parametric oscillator. Activate the heated stage of the microscope (37 &#176;C) and set the lasers at 850 nm and 1100 nm, corresponding to the required wavelengths for the mouse model.</li>
	<li>Use a power meter to set the power of the lasers simultaneously at around 20 mW.</li>
	<li>Follow step 1.4 to prepare the mouse and apply ocular gel on both eyes. 
	NOTE: The animal used for this protocol has to be from a transgenic mouse line (MAGIC-Marker transgenic mouse21) that expresses an endogenous fluorophore in the corneal nerve fibers.</li>
	<li>Install the animal in a head holder. Turn the animal 90&#176; for the eye of interest to face the ceiling.</li>
	<li>Strap the vibrissae of the animal to clear the eye area. Strap the animal&#39;s body to lessen the head movement induced by breathing.</li>
	<li>Position a circular coverslip on the eye above the ocular gel. The coverslip has to be horizontal. Do not hesitate to move the head of the animal for the coverslip to be positioned correctly.</li>
	<li>Add a drop of PBS above the coverslip. Place the animal on the microscope stage carefully and set the objective turret at the right distance.</li>
	<li>Use the aqueous 20x objective and epifluorescence to localize the eye and the innervation through the eyepiece.</li>
	<li>Activate the lasers and define the depth that needs to be illuminated.</li>
	<li>Acquire 1024 x 1024 (pixels) image at an acquisition speed of 70%-85% of the maximum scan speed.</li>
	<li>Once the image is acquired, take the animal off the microscope and remove the coverslip and the straps. Remove the animal from the head holder, add more ocular gel if needed, and let it wake up in a cage on a heated plate. 
	NOTE: Destruction of the innervation can be observed 1 week after the imaging. This protocol can be repeated once a week for several weeks to increase the damage caused by the lasers.</li>
	<li>When the animal is moving on its own, put the cage back in the stable and check its well-being for the following 2 days.</li>
</ol>

Corneal Sensory Fiber Damage Protocols for Axon Regeneration Studies

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C., Norden, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phototoxicity+in+live+fluorescence+microscopy,+and+how+to+avoid+it.">Phototoxicity in live fluorescence microscopy, and how to avoid it.</a> BioEssays. 39 (8), 1700003 (2017).</li><li>Volatier, T., Schumacher, B., Cursiefen, C., Notara, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=UV+protection+in+the+cornea:+Failure+and+rescue.">UV protection in the cornea: Failure and rescue.</a> Biology. 11 (2), 278 (2022).</li><li>Loulier, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiplex+cell+and+lineage+tracking+with+combinatorial+labels.">Multiplex cell and lineage tracking with combinatorial labels.</a> Neuron. 81 (3), 505-520 (2014).</li><li>Dana, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expert+consensus+on+the+identification,+diagnosis,+and+treatment+of+neurotrophic+keratopathy.">Expert consensus on the identification, diagnosis, and treatment of neurotrophic keratopathy.</a> BMC Ophthalmol. 21 (1), 327 (2021).</li><li>Matsumoto, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Autologous+serum+application+in+the+treatment+of+neurotrophic+keratopathy.">Autologous serum application in the treatment of neurotrophic keratopathy.</a> Ophthalmology. 111 (6), 1115-1120 (2004).</li><li>Bonini, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phase+II+randomized,+double-masked,+vehicle-controlled+trial+of+recombinant+human+nerve+growth+factor+for+neurotrophic+keratitis.">Phase II randomized, double-masked, vehicle-controlled trial of recombinant human nerve growth factor for neurotrophic keratitis.</a> Ophthalmology. 125 (9), 1332-1343 (2018).</li><li>Aggarwal, S., Colon, C., Kheirkhah, A., Hamrah, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficacy+of+autologous+serum+tears+for+treatment+of+neuropathic+corneal+pain.">Efficacy of autologous serum tears for treatment of neuropathic corneal pain.</a> Ocul Surf. 17 (3), 532-539 (2019).</li><li>Singh, N. 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The authors have no conflicts of interest to disclose.

Neurotrophic keratitis is considered a rare disease, affecting 5 in 10,000 individuals. However, people suffering from NK due to a physical injury such as chemical burns, or syndromes such as diabetes or multiple sclerosis are not included in those statistics3. Furthermore, this condition remains significantly underdiagnosed22, and the prevalence of the disease is underestimated. There is a strong need for new treatments and therapy that would promote axon regeneration as a...

The cornea, which is the transparent surface of the eye, is composed of three distinct layers: the epithelium, the stroma, and the endothelium. This organ has the highest density of innervation in the body and is composed mainly of sensory fibers (types A&#948; and C) originating from the ophthalmic branch of the trigeminal ganglion. Sensory fibers penetrate the periphery of the cornea in the mid-stroma in the form of big bundles that branch out to cover the surface. They then bifurcate to pierce the Bowmann&#39;s membrane and form the subbasal plexus, which is easily recognizable by the formation of a vortex in the center of the cornea. Those fibers terminate as free nerve endings at the external surface of the epithelium. They are able to transduce thermal, mechanical, and chemical stimuli and to release trophic factors that are essential for epithelium homeostasis1,2. Neurotrophic keratitis (NK) is a degenerative disease affecting the corneal sensory innervation. This rare disease stems from a decrease or a loss of corneal sensitivity that results in lower tear production and poor healing properties of the cornea3. NK progresses through three well-described stages, from stage 1 where patients suffer epithelial defects, to stage 3 where stromal melting and/or corneal perforation occur4.
Clinically, the origins of this disease can be diverse. Patients can lose corneal innervation after physical injury to the eye, surgery, or through chronic diseases, such as diabetes5,6. To date, the NK pathogenesis process remains poorly understood, and therapeutical options for this sight-threatening condition are very limited. Therefore, a better understanding of the characteristics of epithelial defects is needed to better understand the mechanisms behind the regeneration of those fibers and potentially promote them. Here, we propose several models of corneal injury that induce NK in mice.
The first model is the abrasion of the epithelial layer of the cornea with an ocular burr. This model has mainly been studied in the context of the regeneration of the epithelium in different animals, such as rodents and fish7,8,9, and to test molecules promoting corneal healing10,11. Physiologically, it takes 2-3 days for the epithelial cells to close the wound. The physiological pattern of the innervation, however, takes more than four weeks to recover from the abrasion12,13. During the surgery, the ocular burr removes the epithelial layer of the cornea that contains the subbasal plexus and the fibers&#39; free nerve endings. This procedure can be clinically compared to patients with photorefractive keratectomy (PRK) to correct eye refractive defects. The procedure consists of removing the epithelium of the cornea and then reshaping the stroma with a laser14. Patients can experience several side effects following such surgery, such as a decrease of corneal nerve density for 2 years and a reduction in sensitivity for a duration of 3 months to one year post-surgery15. Given that the surgery induces a fragility of the corneal microenvironment, this model could help investigate these side effects and develop therapeutical approaches that would promote faster reinnervation, thus reducing the side effects in question.
The second model consists of sectioning the axons at the periphery of the cornea with a biopsy punch, inducing a Wallerian degeneration of the central innervation 16. Clinically, this method could be compared to anterior lamellar keratoplasty, in which the surgeon realizes a partial trephination of the cornea to remove a part of the anterior thickness of the cornea and replace it with a donor transplant 17. Following lamellar keratoplasty, patients may suffer from a number of symptoms including dry eye, loss of corneal innervation and graft rejection18. This axotomy model performed on corneal nerves could provide insight into the mechanisms of fiber degeneration, which occurs after a graft, followed by the axons&#39; regeneration.
The third method damages the corneal nerves with a laser. By using a multiphoton microscope on the cornea of anesthetized animals, degeneration of the nerves localized in the optical field is induced as a result of reactive oxygen species (ROS) formation, which leads to DNA damage and cellular cavitation19. This method recapitulates the corneal photodamage induced by overexposure to natural UV (sunburn), which also triggers ROS formation, leading to DNA damage20. Patients who suffer from corneal sunburn experience great pain, as the deterioration of epithelial cells deprives the corneal fibers&#39; extremities of all.
The three methods described here are designed to enable the investigation of the NK pathogenesis process and axon regeneration. They are easily reproducible and precise. Moreover, they allow quick recovery and easy monitoring of the animals.

<table><tbody><tr><td>0.2 &amp;micro;m seringe filter</td><td>CLEARLINE</td><td>51733</td><td></td></tr><tr><td>0.5 mm rust ring remover</td><td>Alger Equipment Company</td><td>BU-5S</td><td></td></tr><tr><td>2 mL plastic tubes</td><td>Eppendrof&amp;nbsp;</td><td>30120094</td><td></td></tr><tr><td>Algerbrush burr, Complete instrument</td><td>Alger Equipment Company</td><td>BR2-5</td><td></td></tr><tr><td>Anti-beta III Tubulin antibody</td><td>Abcam</td><td>ab18207</td><td></td></tr><tr><td>Antigenfix</td><td>Diapath</td><td>P0016</td><td></td></tr><tr><td>Artificial tear</td><td>Larmes artificielles Martinet</td><td>N/A</td><td></td></tr><tr><td>Buprecare</td><td>Animalcare</td><td>N/A</td><td></td></tr><tr><td>Cotton swab</td><td>Any provider</td><td>N/A</td><td></td></tr><tr><td>Dissecting tools</td><td>Fine Science Tools</td><td>N/A</td><td></td></tr><tr><td>Fluorescein</td><td>Merck</td><td>103887</td><td></td></tr><tr><td>Gelatin from cold water fish skin</td><td>Sigma</td><td>G7765</td><td></td></tr><tr><td>Goat serum</td><td>Merck</td><td>S26</td><td></td></tr><tr><td>Head Holder</td><td>Narishige</td><td>SGM 4</td><td></td></tr><tr><td>Heated plate</td><td>BIOSEB LAB instruments</td><td>BIO-HE002</td><td></td></tr><tr><td>Hoechst 33342</td><td>Thermo Fisher Scientific</td><td>H3570</td><td></td></tr><tr><td>Imalgene 1000</td><td>BOEHRINGER INGELHEIM ANIMAL HEALTH France</td><td>N/A</td><td>French marketing authorization numbre: FR/V/0167433 4/1992</td></tr><tr><td>LAS X software</td><td>Leica</td><td>N/A</td><td>Large volume computational clearing (LVCC) process</td></tr><tr><td>Laser Chameleon Ultra II</td><td>Coherent</td><td>N/A</td><td></td></tr><tr><td>Laser power meter</td><td>Coherent</td><td>N/A</td><td></td></tr><tr><td>Leica Thunder Imager Tissue microscope</td><td>Leica</td><td>N/A</td><td></td></tr><tr><td>Multi-photon Zeiss LSM 7MP upright microscope</td><td>Zeiss</td><td>N/A</td><td></td></tr><tr><td>Ocry-gel</td><td>TVM lab</td><td>N/A</td><td></td></tr><tr><td>Parametric oscillator</td><td>Coherent</td><td>N/A</td><td></td></tr><tr><td>Penlights with blue cobalt filtercap</td><td>Bernell</td><td>ALPEN</td><td></td></tr><tr><td>Petri dish</td><td>Thermo Scientific</td><td>150318</td><td>Axotomy protocol</td></tr><tr><td>Petridish</td><td>Thermo Scientific</td><td>150288</td><td>Cornea whole-mount processing</td></tr><tr><td>Rompun 2%</td><td>Elanco</td><td>N/A</td><td>French marketing authorization numbre: FR/V/8146715 2/1980</td></tr><tr><td>Sterile biopsy punch 2.5 mm</td><td>LCH medical</td><td>LCH-PUK-25</td><td></td></tr><tr><td>Triton X-100</td><td>VWR</td><td>0694</td><td></td></tr><tr><td>Vectashield</td><td>EuroBioSciences</td><td>H-1000</td><td>Mounting medium</td></tr></tbody></table>

three strategies to induce neurotrophic keratitis and nerve regeneration in murine cornea

The cornea is a transparent tissue that covers the eye and is crucial for clear vision. It is the most innervated tissue in the body. This innervation provides sensation and trophic function to the eye and contributes to preserving corneal integrity. The pathological disruption of this innervation is termed neurotrophic keratitis. This can be triggered by injury to the eye, surgery, or disease. In this study, we propose three different protocols for inflicting damage on the innervation in ways that recapitulate the three types of cases generally encountered in the clinic.
The first method consists in making an abrasion of the epithelium with an ophthalmic burr. This involves the removal of the epithelial layer, the free nerve endings, and the subbasal plexus in a manner similar to the photorefractive keratectomy surgery performed in the clinic. The second method&#160;only targets the innervation by sectioning it at the periphery with a biopsy punch, maintaining the integrity of the epithelium. This method is similar to the first steps of lamellar keratoplasty and leads to a degeneration of the innervation followed by regrowth of the axons in the central cornea. The last method damages the innervation of a transgenic mouse model using a multiphoton microscope, which specifically localizes the site of cauterization of the fluorescent nerve fibers. This method inflicts the same damage as photokeratitis, an overexposure to UV light.
This study describes different options for investigating the physiopathology of corneal innervation, particularly the degeneration and regeneration of the axons. Promoting regeneration is crucial for avoiding such complications as epithelium defects or even perforation of the cornea. The proposed models can help test new pharmacological molecules or gene therapy that enhance nerve regeneration and limit disease progression.

This study proposes several protocols for inflicting damage on corneal innervation in mice. While similar protocols have been used to investigate the physiopathology of the healing of the epithelium, we chose to adapt and develop new methods of investigating corneal innervation regeneration. To observe the innervation, we used two techniques. First, we employed an immunofluorescence technique to stain the nerve fibers using a pan-neuronal antibody (BIII tubulin) and the nuclei with an intercalator. Second, we took advant...

Watch this Scientific Journal Video about Advancing Corneal Innervation Research Through Innovative Models at JoVE.com

Author Spotlight: Advancing Corneal Innervation Research Through Innovative Models 

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.

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

The cornea is a transparent tissue that covers the eye and is crucial for clear vision. It is the most innervated tissue in the body. This innervation ...

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Research

JoVE Journal

Neuroscience

1.5K Views.  University of  Montpellier, INSERM. 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.

Video: Author Spotlight: Advancing Corneal Innervation Research Through Innovative Models 

Methods for Experimental Manipulations after Optic Nerve Transection in the Mammalian CNS

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. The optic nerve can be accessed within the orbit of the eye and completely transected (axotomized), cutting the axons of the entire RGC population. Optic nerve transection is a reproducible model of apoptotic neuronal cell death in the adult CNS 1-4. This model is particularly attractive because the vitreous chamber of the eye acts as a capsule for drug delivery to the retina, permitting experimental manipulations via intraocular injections. The diffusion of chemicals through the vitreous fluid ensures that they act upon the entire RGC population. Viral vectors, plasmids or short interfering RNAs (siRNAs) can also be delivered to the vitreous chamber in order to infect or transfect retinal cells 5-12. The high tropism of Adeno-Associated Virus (AAV) vectors is beneficial to target RGCs, with an infection rate approaching 90% of cells near the injection site 6, 7, 13-15. Moreover, RGCs can be selectively transfected by applying siRNAs, plasmids, or viral vectors to the cut end of the optic nerve 16-19 or injecting vectors into their target the superior colliculus 10. This allows researchers to study apoptotic mechanisms in the injured neuronal population without confounding effects on other bystander neurons or surrounding glia. RGC apoptosis has a characteristic time-course whereby cell death is delayed 3-4 days postaxotomy, after which the cells rapidly degenerate. This provides a window for experimental manipulations directed against pathways involved in apoptosis. Manipulations that directly target RGCs from the transected optic nerve stump are performed at the time of axotomy, immediately after cutting the nerve. In contrast, when substances are delivered via an intraocular route, they can be injected prior to surgery or within the first 3 days after surgery, preceding the initiation of apoptosis in axotomized RGCs. In the present article, we demonstrate several methods for experimental manipulations after optic nerve transection.

Optic Nerve transection is a widely used model of adult CNS injury. This model is ideal for performing a number of experimental manipulations that target the retina globally or directly target the injured neuronal population of retinal ganglion cells.

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. The optic nerve can be ...

Adenoviral Gene Therapy for Diabetic Keratopathy: Effects on Wound Healing and Stem Cell Marker Expression in Human Organ-cultured Corneas and Limbal Epithelial Cells

The goal of this protocol is to describe molecular alterations in human diabetic corneas and demonstrate how they can be alleviated by adenoviral gene therapy in organ-cultured corneas. The diabetic corneal disease is a complication of diabetes with frequent abnormalities of corneal nerves and epithelial wound healing. We have also documented significantly altered expression of several putative epithelial stem cell markers in human diabetic corneas. To alleviate these changes, adenoviral gene therapy was successfully implemented using the upregulation of c-met proto-oncogene expression and/or the downregulation of proteinases matrix metalloproteinase-10 (MMP-10) and cathepsin F. This therapy accelerated wound healing in diabetic corneas even when only the limbal stem cell compartment was transduced. The best results were obtained with combined treatment. For possible patient transplantation of normalized stem cells, an example is also presented of the optimization of gene transduction in stem cell-enriched cultures using polycationic enhancers. This approach may be useful not only for the selected genes but also for the other mediators of corneal epithelial wound healing and stem cell function.

An example of adenoviral gene therapy in the human diabetic organ-cultured corneas is presented towards the normalization of delayed wound healing and markedly reduced epithelial stem cell marker expression in these corneas. It also describes the optimization of this process in stem cell-enriched limbal epithelial cultures.

The goal of this protocol is to describe molecular alterations in human diabetic corneas and demonstrate how they can be alleviated by adenoviral gene ...

Developmental Biology

Combination of Microstereolithography and Electrospinning to Produce Membranes Equipped with Niches for Corneal Regeneration

Corneal problems affect millions of people worldwide reducing their quality of life significantly. Corneal disease can be caused by illnesses such as Aniridia or Steven Johnson Syndrome as well as by external factors such as chemical burns or radiation. Current treatments are (i) the use of corneal grafts and (ii) the use of stem cell expanded in the laboratory and delivered on carriers (e.g., amniotic membrane); these treatments are relatively successful but unfortunately they can fail after 3-5 years. There is a need to design and manufacture new corneal biomaterial devices able to mimic in detail the physiological environment where stem cells reside in the cornea. Limbal stem cells are located in the limbus (circular area between cornea and sclera) in specific niches known as the Palisades of Vogt. In this work we have developed a new platform technology which combines two cutting-edge manufacturing techniques (microstereolithography and electrospinning) for the fabrication of corneal membranes that mimic to a certain extent the limbus. Our membranes contain artificial micropockets which aim to provide cells with protection as the Palisades of Vogt do in the eye.

We report a technique for the fabrication of micropockets within electrospun membranes in which to study cell behavior. Specifically, we describe a combination of microstereolithography and electrospinning for the production of PLGA (Poly(lactide-co-glycolide)) corneal biomaterial devices equipped with microfeatures.

Corneal problems affect millions of people worldwide reducing their quality of life significantly. Corneal disease can be caused by illnesses such as ...

Bioengineering

An Alkali-burn Injury Model of Corneal Neovascularization in the Mouse

Under normal conditions, the cornea is avascular, and this transparency is essential for maintaining good visual acuity. Neovascularization (NV) of the cornea, which can be caused by trauma, keratoplasty or infectious disease, breaks down the so called &lsquo;angiogenic privilege' of the cornea and forms the basis of multiple visual pathologies that may even lead to blindness. Although there are several treatment options available, the fundamental medical need presented by corneal neovascular pathologies remains unmet. In order to develop safe, effective, and targeted therapies, a reliable model of corneal NV and pharmacological intervention is required. Here, we describe an alkali-burn injury corneal neovascularization model in the mouse. This protocol provides a method for the application of a controlled alkali-burn injury to the cornea, administration of a pharmacological compound of interest, and visualization of the result. This method could prove instrumental for studying the mechanisms and opportunities for intervention in corneal NV and other neovascular disorders.

Neovascularization (NV) of the cornea can complicate multiple visual pathologies. Utilizing a controlled, alkali-burn injury model, a quantifiable level of corneal NV can be produced for mechanistic study of corneal NV and evaluation of potential therapies for neovascular disorders.

Under normal conditions, the cornea is avascular, and this transparency is essential for maintaining good visual acuity. Neovascularization (NV) of the ...

Medicine

Murine Corneal Transplantation: A Model to Study the Most Common Form of Solid Organ Transplantation

Corneal transplantation is the most common form of organ transplantation in the United States with between 45,000 and 55,000 procedures performed each year. While several animal models exist for this procedure and mice are the species that is most commonly used. The reasons for using mice are the relative cost of using this species, the existence of many genetically defined strains that allow for the study of immune responses, and the existence of an extensive array of reagents that can be used to further define responses in this species. This model has been used to define factors in the cornea that are responsible for the relative immune privilege status of this tissue that enables corneal allografts to survive acute rejection in the absence of immunosuppressive therapy. It has also been used to define those factors that are most important in rejection of such allografts. Consequently, much of what we know concerning mechanisms of both corneal allograft acceptance and rejection are due to studies using a murine model of corneal transplantation. In addition to describing a model for acute corneal allograft rejection, we also present for the first time a model of late-term corneal allograft rejection.

Mice have been used as a model for studying many forms of transplantation, including corneal transplantation. We describe in this report a murine model for both acute and late-term corneal transplantation.

Corneal transplantation is the most common form of organ transplantation in the United States with between 45,000 and 55,000 procedures performed each ...

Minimally-invasive Technique for Injection into Rat Optic Nerve

The rat optic nerve is a useful model for stem cell regeneration research. Direct injection into the rat optic nerve allows delivery into the central nervous system in a minimally-invasive surgery without bone removal. This technique describes an approach to visualization and direct injection of the optic nerve following minor fascial dissection from the orbital ridge, using a conjunctival traction suture to gently pull the eye down and out. Representative examples of an injected optic nerve show successful injection of dyed beads.

Direct injection into the rat optic nerve is useful for regenerative research. We demonstrate a minimally-invasive technique for direct injection into a rat optic nerve that does not involve opening the skull. Using this method, surgical complications are minimized and recovery is more rapid.

The rat optic nerve is a useful model for stem cell regeneration research. Direct injection into the rat optic nerve allows delivery into the central ...

Ex Vivo Corneal Organ Culture Model for Wound Healing Studies

The cornea has been used extensively as a model system to study wound healing. The ability to generate and utilize primary mammalian cells in two dimensional (2D) and three dimensional (3D) culture has generated a wealth of information not only about corneal biology but also about wound healing, myofibroblast biology, and scarring in general. The goal of the protocol is an assay system for quantifying myofibroblast development, which characterizes scarring. We demonstrate a corneal organ culture ex vivo model using pig eyes. In this anterior keratectomy wound, corneas still in the globe are wounded with a circular blade called a trephine. A plug of approximately 1/3 of the anterior cornea is removed including the epithelium, the basement membrane, and the anterior part of the stroma. After wounding, corneas are cut from the globe, mounted on a collagen/agar base, and cultured for two weeks in supplemented-serum free medium with stabilized vitamin C to augment cell proliferation and extracellular matrix secretion by resident fibroblasts. Activation of myofibroblasts in the anterior stroma is evident in the healed cornea. This model can be used to assay wound closure, the development of myofibroblasts and fibrotic markers, and for toxicology studies. In addition, the effects of small molecule inhibitors as well as lipid-mediated siRNA transfection for gene knockdown can be tested in this system.

A protocol for an ex vivo corneal organ culture model useful for wound healing studies is described. This model system can be used to assess the effects of agents to promote regenerative healing or drug toxicity in an organized 3D multicellular environment.

The cornea has been used extensively as a model system to study wound healing. The ability to generate and utilize primary mammalian cells in two ...

Sheath-Preserving Optic Nerve Transection in Rats to Assess Axon Regeneration and Interventions Targeting the Retinal Ganglion Cell Axon

Retinal ganglion cell (RGC) axons converge at the optic nerve head to convey visual information from the retina to the brain. Pathologies such as glaucoma, trauma, and ischemic optic neuropathies injure RGC axons, disrupt transmission of visual stimuli, and cause vision loss. Animal models simulating RGC axon injury include optic nerve crush and transection paradigms. Each of these models has inherent advantages and disadvantages. An optic nerve crush is generally less severe than a transection and can be used to assay axon regeneration across the lesion site. However, differences in crush force and duration can affect tissue responses, resulting in variable reproducibility and lesion completeness. With optic nerve transection, there is a severe and reproducible injury that completely lesions all axons. However, transecting the optic nerve dramatically alters the blood brain barrier by violating the optic nerve sheath, exposing the optic nerve to the peripheral environment. Moreover, regeneration beyond a transection site cannot be assessed without reapposing the cut nerve ends. Furthermore, distinct degenerative changes and cellular pathways are activated by either a crush or transection injury.
The method described here incorporates the advantages of both optic nerve crush and transection models while mitigating the disadvantages. Hydrostatic pressure delivered into the optic nerve by microinjection completely transects the optic nerve while maintaining the integrity of the optic nerve sheath. The transected optic nerve ends are reapposed to allow for axon regeneration assays. A potential limitation of this method is the inability to visualize the complete transection, a potential source of variability. However, visual confirmation that the visible portion of the optic nerve has been transected is indicative of a complete optic nerve transection with 90-95% success. This method could be applied to assess axon regeneration promoting strategies in a transection model or investigate interventions that target the axonal compartments.

This protocol describes an optic nerve transection that preserves the optic nerve sheath in rats. Hydrostatic pressure from microinjections into the optic nerve generate a complete transection, allowing for sutureless reapposition of the transected optic nerve ends and direct targeting of the axonal compartment in a transection model.

Retinal ganglion cell (RGC) axons converge at the optic nerve head to convey visual information from the retina to the brain. Pathologies such as ...

A Modified Biopsy Punch Method for Precise Alkali Burns in Mice

Corneal and Limbal Alkali Injury Induction Using a Punch-Trephine Technique in a Mouse Model

The cornea is critical for vision, and corneal healing after trauma is fundamental in maintaining its transparency and function. Through the study of corneal injury models, researchers aim to enhance their understanding of how the cornea heals and develop strategies to prevent and manage corneal opacities. Chemical injury is one of the most popular injury models that has extensively been studied on mice. Most previous investigators have used a flat paper soaked in sodium hydroxide to induce corneal injury. However, inducing corneal and limbal injury using flat filter paper is unreliable, since the mouse cornea is highly curved. Here, we present a new instrument, a modified biopsy punch, that enables the researchers to create a well-circumscribed, localized, and evenly distributed alkali injury to the murine cornea and limbus. This punch-trephine method enables researchers to induce an accurate and reproducible chemical burn to the entire murine cornea and limbus while leaving other structures, such as the eyelids, unaffected by the chemical. Moreover, this study introduces an enucleation technique that preserves the medial caruncle as a landmark for identifying the nasal side of the globe. The bulbar and palpebral conjunctiva, and lacrimal gland are also kept intact using this technique. Ophthalmologic examinations were performed via slit lamp biomicroscope and fluorescein staining on days 0, 1, 2, 6, 8, and 14 post-injury. Clinical, histological, and immunohistochemical findings confirmed limbal stem cell deficiency and ocular surface regeneration failure in all experimental mice. The presented alkali corneal injury model is ideal for studying limbal stem cell deficiency, corneal inflammation, and fibrosis. This method is also suitable for investigating pre-clinical and clinical efficacies of topical ophthalmologic medications on the murine corneal surface.

Author Spotlight: Advancing Research in Corneal Opacity Treatment and Regeneration 

This protocol describes a method to induce an accurate and reproducible corneal and limbal alkali injury in a mouse model. The protocol is advantageous as it allows for an evenly distributed injury to the highly curved mouse cornea and limbus.

The cornea is critical for vision, and corneal healing after trauma is fundamental in maintaining its transparency and function. Through the study of ...

Assessing the Health of the Corneal Nerves in Ocular Surface Diseases

Corneal Sensitivity Testing Procedure for Ophthalmologic and Optometric Patients

The cornea is the most densely innervated structure in the human body, making it one of the most sensitive tissues. Changes in corneal nerve sensitivity can be observed in several ocular surface diseases. Nerve sensitivity may be increased, as is often seen in patients with a neuropathic component to ocular pain, or decreased, as is seen in patients with neurotrophic keratitis. Corneal sensitivity testing involves assessing a patient's reaction to brief corneal stimulation, yields insight into the health of the corneal nerves, and provides diagnostic value for evaluating the health of the nerves and the interplay with the ocular surface. Currently, there is limited published guidance on how to conduct corneal sensitivity testing in a clinical setting. This article presents a protocol for testing corneal sensitivity using easy-to-use, low-cost materials that are readily accessible to eye care providers (either a cotton swab, a piece of dental floss, or a finely tapered tissue). This protocol allows for qualitative assessment of corneal sensitivity in which responses to corneal stimulation are rated from 0 (no response) to 3 (hypersensitive response). This test can be performed quickly (in approximately 30 s). Given its diagnostic value and accessibility, corneal sensitivity testing should be included as part of the standard eye examination for any patient undergoing an ocular surface examination.

 Author Spotlight: Establishing a Practical and Cost-Effective Protocol for Corneal Sensitivity Testing in Clinical Settings

Corneal sensitivity testing provides insight into the health of the corneal nerves and helps diagnose ocular surface diseases. We present a concise protocol to qualitatively assess corneal sensitivity that can be readily used by eye care providers across clinical settings.

The cornea is the most densely innervated structure in the human body, making it one of the most sensitive tissues. Changes in corneal nerve sensitivity ...