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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.2 &#181;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&#160;</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>
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		<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 ...

three-strategies-to-induce-neurotrophic-keratitis-nerve-regeneration

Research

JoVE Journal

Neuroscience

1.4K 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 

An Optic Nerve Crush Injury Murine Model to Study Retinal Ganglion Cell Survival

Injury to the optic nerve can lead to axonal degeneration, followed by a gradual death of retinal ganglion cells (RGCs), which results in irreversible vision loss. Examples of such diseases in human include traumatic optic neuropathy and optic nerve degeneration in glaucoma. It is characterized by typical changes in the optic nerve head, progressive optic nerve degeneration, and loss of retinal ganglion cells, if uncontrolled, leading to vision loss and blindness.
The optic nerve crush (ONC) injury mouse model is an important experimental disease model for traumatic optic neuropathy, glaucoma, etc. In this model, the crush injury to the optic nerve leads to gradual retinal ganglion cells apoptosis. This disease model can be used to study the general processes and mechanisms of neuronal death and survival, which is essential for the development of therapeutic measures. In addition, pharmacological and molecular approaches can be used in this model to identify and test potential therapeutic reagents to treat different types of optic neuropathy.
Here, we provide a step by step demonstration of (I) Baseline retrograde labeling of retinal ganglion cells (RGCs) at day 1, (II) Optic nerve crush injury at day 4, (III) Harvest the retinae and analyze RGC survival at day 11, and (IV) Representative result.

This protocol shows how to retrogradely label retinal ganglion cells, and how to subsequently make an optic nerve crush injury in order to analyze retinal ganglion cell survival and apoptosis. It is an experimental disease model for different types of optic neuropathy, including glaucoma.

Injury to the optic nerve can lead to axonal degeneration, followed by a gradual death of retinal ganglion cells (RGCs), which results in irreversible ...

Reproducible Mouse Sciatic Nerve Crush and Subsequent Assessment of Regeneration by Whole Mount Muscle Analysis

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative response mounted by PNS neurons thereby possibly illuminating the failures of CNS regeneration1. Sciatic nerve crush (axonotmesis) is one of the most common models of peripheral nerve injury in rodents2. Crushing interrupts all axons but Schwann cell basal laminae are preserved so that regeneration is optimal3,4. This allows the investigator to study precisely the ability of a growing axon to interact with both the Schwann cell and basal laminae4. Rats have generally been the preferred animal models for experimental nerve crush. They are widely available and their lesioned sciatic nerve provides a reasonable approximation of human nerve lesions5,4. Though smaller in size than rat nerve, the mouse nerve has many similar qualities. Most importantly though, mouse models are increasingly valuable because of the wide availability of transgenic lines now allows for a detailed dissection of the individual molecules critical for nerve regeneration6, 7. Prior investigators have used multiple methods to produce a nerve crush or injury including simple angled forceps, chilled forceps, hemostatic forceps, vascular clamps, and investigator-designed clamps8,9,10,11,12. Investigators have also used various methods of marking the injury site including suture, carbon particles and fluorescent beads13,14,1. We describe our method to obtain a reproducibly complete sciatic nerve crush with accurate and persistent marking of the crush-site using a fine hemostatic forceps and subsequent carbon crush-site marking. As part of our description of the sciatic nerve crush procedure we have also included a relatively simple method of muscle whole mount we use to subsequently quantify regeneration.

In this report we describe a method to crush mouse sciatic nerve. This method uses readily available hemostatic forceps and easily and reproducibly produces complete sciatic nerve crush. In addition, we describe a method to prepare muscle whole mounts suitable for analysis of nerve regeneration after sciatic nerve crush.

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative ...

Transplantation of Olfactory Ensheathing Cells to Evaluate Functional Recovery after Peripheral Nerve Injury

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth and regrowth of the primary olfactory neurons. Indeed, the primary olfactory system is characterized by its ability to give rise to new neurons even in adult animals. This particular ability is partly due to the presence of OECs which create a favorable microenvironment for neurogenesis. This property of OECs has been used for cellular transplantation such as in spinal cord injury models. Although the peripheral nervous system has a greater capacity to regenerate after nerve injury than the central nervous system, complete sections induce misrouting during axonal regrowth in particular after facial of laryngeal nerve transection. Specifically, full sectioning of the recurrent laryngeal nerve (RLN) induces aberrant axonal regrowth resulting in synkinesis of the vocal cords. In this specific model, we showed that OECs transplantation efficiently increases axonal regrowth.
OECs are constituted of several subpopulations present in both the olfactory mucosa (OM-OECs) and the olfactory bulbs (OB-OECs). We present here a model of cellular transplantation based on the use of these different subpopulations of OECs in a RLN injury model. Using this paradigm, primary cultures of OB-OECs and OM-OECs were transplanted in Matrigel after section and anastomosis of the RLN. Two months after surgery, we evaluated transplanted animals by complementary analyses based on videolaryngoscopy, electromyography (EMG), and histological studies. First, videolaryngoscopy allowed us to evaluate laryngeal functions, in particular muscular cocontractions phenomena. Then, EMG analyses demonstrated richness and synchronization of muscular activities. Finally, histological studies based on toluidine blue staining allowed the quantification of the number and profile of myelinated fibers.
All together, we describe here how to isolate, culture, identify and transplant OECs from OM and OB after RLN section-anastomosis and how to evaluate and analyze the efficiency of these transplanted cells on axonal regrowth and laryngeal functions.

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth of the primary olfactory neurons. This specific property can be used for cellular transplantation. We present here a model of cellular transplantation based on the use of OECs in a laryngeal nerve injury model.

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth and regrowth of the primary olfactory neurons. Indeed, the primary olfactory ...

Dorsal Root Ganglia Neurons and Differentiated Adipose-derived Stem Cells: An In Vitro Co-culture Model to Study Peripheral Nerve Regeneration

Dorsal root ganglia (DRG) neurons, located in the intervertebral foramina of the spinal column, can be used to create an in vitro system facilitating the study of nerve regeneration and myelination. The glial cells of the peripheral nervous system, Schwann cells (SC), are key facilitators of these processes; it is therefore crucial that the interactions of these cellular components are studied together. Direct contact between DRG neurons and glial cells provides additional stimuli sensed by specific membrane receptors, further improving the neuronal response. SC release growth factors and proteins in the culture medium, which enhance neuron survival and stimulate neurite sprouting and extension. However, SC require long proliferation time to be used for tissue engineering applications and the sacrifice of an healthy nerve for their sourcing. Adipose-derived stem cells (ASC) differentiated into SC phenotype are a valid alternative to SC for the set-up of a co-culture model with DRG neurons to study nerve regeneration. The present work presents a detailed and reproducible step-by-step protocol to harvest both DRG neurons and ASC from adult rats; to differentiate ASC towards a SC phenotype; and combines the two cell types in a direct co-culture system to investigate the interplay between neurons and SC in the peripheral nervous system. This tool has great potential in the optimization of tissue-engineered constructs for peripheral nerve repair.

Dorsal Root Ganglia Neurons and Differentiated Adipose-derived Stem Cells: An In Vitro Co-culture Model to Study Peripheral Nerve Regeneration

Dorsal root ganglia (DRG) are structures containing the sensory neurons of the peripheral nervous system. When dissociated, they can be co-cultured with SC-like adipose-derived stem cells (ASC), providing a valuable model to study in vitro nerve regeneration and myelination, mimicking the in vivo environment at the injury site.

Dorsal root ganglia (DRG) neurons, located in the intervertebral foramina of the spinal column, can be used to create an in vitro system facilitating the ...

Three-dimensional Imaging and Analysis of Mitochondria within Human Intraepidermal Nerve Fibers

The goal of this protocol is to study mitochondria within intraepidermal nerve fibers. Therefore, 3D imaging and analysis techniques were developed to isolate nerve-specific mitochondria and evaluate disease-induced alterations of mitochondria in the distal tip of sensory nerves. The protocol combines fluorescence immunohistochemistry, confocal microscopy and 3D image analysis techniques to visualize and quantify nerve-specific mitochondria. Detailed parameters are defined throughout the procedures in order to provide a concrete example of how to use these techniques to isolate nerve-specific mitochondria. Antibodies were used to label nerve and mitochondrial signals within tissue sections of skin punch biopsies, which was followed by indirect immunofluorescence to visualize nerves and mitochondria with a green and red fluorescent signal respectively. Z-series images were acquired with confocal microscopy and 3D analysis software was used to process and analyze the signals. It is not necessary to follow the exact parameters described within, but it is important to be consistent with the ones chosen throughout the staining, acquisition and analysis steps. The strength of this protocol is that it is applicable to a wide variety of circumstances where one fluorescent signal is used to isolate other signals that would otherwise be impossible to study alone.

This protocol uses three-dimensional (3D) imaging and analysis techniques to visualize and quantify nerve-specific mitochondria. The techniques are applicable to other situations where one fluorescent signal is used to isolate a subset of data from another fluorescent signal.

The goal of this protocol is to study mitochondria within intraepidermal nerve fibers. Therefore, 3D imaging and analysis techniques were developed to ...

Ethanol-Induced Cervical Sympathetic Ganglion Block Applications for Promoting Canine Inferior Alveolar Nerve Regeneration Using an Artificial Nerve

Polyglycolic acid collagen (PGA-C) tubes are bio-absorbable nerve tubes filled with collagen of multi-chamber structure, which consist of thin collagen films. Favorable clinical outcomes have been achieved when using these tubes for the treatment of damaged inferior alveolar nerve (IAN). A critical factor for the successful nerve regeneration using PGA-C tubes is blood supply to the surrounding tissue. Cervical sympathetic ganglion block (CSGB) creates a sympathetic blockade in the head and neck region thus increasing blood flow in the area. To ensure an adequate effect, the blockade must be administered with local anesthetics one to two times a day for several consecutive weeks; this poses a challenge when creating animal models for investigating this technique. To address this limitation, we developed an ethanol-induced CSGB in a canine model of long-term increase in blood flow in the orofacial region. We examined whether IAN regeneration via PGA-C tube implantation can be enhanced by this model. Fourteen Beagles were each implanted with a PGA-C tube across a 10-mm gap in the left IAN. The IAN is located within the mandibular canal surrounded by bone, therefore we chose piezoelectric surgery, consisting of ultrasonic waves, for bone processing, in order to minimize the risk of nerve and vessel injury. A good surgical outcome was obtained with this approach. A week after surgery, seven of these dogs were subjected to left CSGB by injection of ethanol. Ethanol-induced CSGB resulted in improved nerve regeneration, suggesting that the increased blood flow effectively promotes nerve regeneration in IAN defects. This canine model can contribute to further research on the long-term effects of CSGB.

We evaluated the effect of cervical sympathetic ganglion block on nerve repair using artificial nerve conduits. Male beagle dogs were each implanted with an artificial nerve across a 10-mm gap in the left inferior alveolar nerve; left cervical sympathetic ganglion was blocked by injecting 99.5% ethanol via lateral thoracotomy.

Polyglycolic acid collagen (PGA-C) tubes are bio-absorbable nerve tubes filled with collagen of multi-chamber structure, which consist of thin collagen ...

Facial Nerve Surgery in the Rat Model to Study Axonal Inhibition and Regeneration

This protocol describes consistent and reproducible methods to study axonal regeneration and inhibition in a rat facial nerve injury model. The facial nerve can be manipulated along its entire length, from its intracranial segment to its extratemporal course. There are three primary types of nerve injury used for the experimental study of regenerative properties: nerve crush, transection, and nerve gap. The range of possible interventions is vast, including surgical manipulation of the nerve, delivery of neuroactive reagents or cells, and either central or end-organ manipulations. Advantages of this model for studying nerve regeneration include simplicity, reproducibility, interspecies consistency, reliable survival rates of the rat, and an increased anatomic size relative to murine models. Its limitations involve a more limited genetic manipulation versus the mouse model and the superlative regenerative capability of the rat, such that the facial nerve scientist must carefully assess time points for recovery and whether to translate results to higher animals and human studies. The rat model for facial nerve injury allows for functional, electrophysiological, and histomorphometric parameters for the interpretation and comparison of nerve regeneration. It thereby boasts tremendous potential toward furthering the understanding and treatment of the devastating consequences of facial nerve injury in human patients.

This protocol describes a reproducible approach to facial nerve surgery in the rat model, including descriptions of various inducible patterns of injury.

This protocol describes consistent and reproducible methods to study axonal regeneration and inhibition in a rat facial nerve injury model. The facial ...

A Revised Surgical Approach to Induce Endolymphatic Hydrops in the Guinea Pig

Endolymphatic hydrops is an enlargement of scala media that is most often associated with Meniere&#39;s disease, though the pathophysiologic mechanism(s) remain unclear. In order to adequately study the attributes of endolymphatic hydrops, such as the origins of low-frequency hearing loss, a reliable model is needed. The guinea pig is a&#160;good model because it hears in the low-frequency regions that are putatively affected by endolymphatic hydrops. Previous research has demonstrated that endolymphatic hydrops can be induced surgically via intradural or extradural approaches that involve drilling on the endolymphatic duct and sac. However, whether it was possible to create an endolymphatic hydrops model using an extradural approach that avoided dangerous drilling on the endolymphatic duct and sac was unknown. The objective of this study was to demonstrate a revised extradural approach to induce experimental endolymphatic hydrops at 30 days post-operatively by obliterating the endolymphatic sac and injuring the endolymphatic duct with a fine pick. The sample size consisted of seven guinea pigs. Functional measurements of hearing were made and temporal bones were subsequently harvested for histologic analysis. The approach had a success rate of 86% in achieving endolymphatic hydrops. The risk of cerebrospinal fluid leak was minimal. No perioperative deaths or injuries to the posterior semicircular canal occurred in the sample. The presented method demonstrates a safe and reliable way to induce endolymphatic hydrops at a relatively quick time point of 30 days. The clinical implications are that the presented method provides a reliable model to further explore the origins of low-frequency hearing loss that can be associated endolymphatic hydrops.

This article demonstrates an extradural approach to obliterate the guinea pig endolymphatic sac and injure the endolymphatic duct with a fine pick in order to induce experimental endolymphatic hydrops.

Endolymphatic hydrops is an enlargement of scala media that is most often associated with Meniere&#39;s disease, though the pathophysiologic mechanism(s) ...

Assessing Early Stage Open-Angle Glaucoma in Patients by Isolated-Check Visual Evoked Potential

Recently, the isolated-check visual evoked potential (icVEP) technique was designed and has been reported to detect glaucomatous damage earlier and faster. It creates low spatial frequency/high temporal frequency bright stimuli and records cortical activity initiated primarily by afferents in the magnocellular ON pathway. This pathway contains neurons with larger volumes and axonal diameters, and it is preferentially damaged in early glaucoma, which can result in visual field loss. The study presented here uses standard operative procedures (SOP) of icVEP to obtain reliable results. It can detect visual function loss using a signal-to-noise ratio (SNR) corresponding to the defects of retinal nerve fiber layer (RNFL) in early stage open-angle glaucoma (OAG). A setting of 10 Hz and condition of 15% positive-contrast (bright) are selected to differentiate OAG patients and control subjects, with each check containing eight runs. Each run persists for 2 s (for 20 total cycles). A flowchart is constructed, which consists of pupil size and intraocular pressure over a 30 min rest period before each examination. Additionally, the testing order of eyes is performed to obtain reliable electroencephalographic signals. VEPs are recorded and analyzed automatically by software, and SNRs are derived based on a multivariate statistic. An SNR of &#8804; 1 is considered abnormal. A receiver-operating-characteristic (ROC) curve is applied to analyze the accuracy of group classification. Then, the SOP is applied in a cross-sectional study, showing that icVEP can detect glaucomatous visual function abnormality in the central visual field in the form of SNR. This value also correlates with the thickness thinning of RNFL and produces high classification accuracy for early stage OAG. Thus, it serves as a useful and objective diagnostic technology for the early detection of glaucoma.

The isolated-check visual evoked potential (icVEP) method is implemented here to assess the magnocellular ON pathway that is initially damaged in glaucoma. The study shows standard operative procedures using icVEP to obtain reliable results. It is proved to serve as a useful objective diagnosing technology for the early detection of glaucoma.

Recently, the isolated-check visual evoked potential (icVEP) technique was designed and has been reported to detect glaucomatous damage earlier and ...

Purification of Fibroblasts and Schwann Cells from Sensory and Motor Nerves in Vitro

The principal cells in the peripheral nervous system are the Schwann cells (SCs) and the fibroblasts. Both these cells distinctly express the sensory and motor phenotypes involved in different patterns of neurotrophic factor gene expression and other biological processes, affecting nerve regeneration. The present study has established a protocol to obtain highly purified rat sensory and motor SCs and fibroblasts more rapidly. The ventral root (motor nerve) and the dorsal root (sensory nerve) of neonatal rats (7-days-old) were dissociated and the cells were cultured for 4-5 days, followed by isolation of sensory and motor fibroblasts and SCs by combining differential digestion and differential adherence methods sequentially. The results of immunocytochemistry and flow cytometry analyses showed that the purity of the sensory and motor SCs and fibroblasts were &#62;90%. This protocol can be used to obtain a large number of sensory and motor fibroblasts/SCs more rapidly, contributing to the exploration of sensory and motor nerve regeneration.

Here, we present a method to purify fibroblasts and Schwann cells from sensory and motor nerves in vitro.

The principal cells in the peripheral nervous system are the Schwann cells (SCs) and the fibroblasts. Both these cells distinctly express the sensory and ...