Name: corneal epithelial abrasion with ocular burr as a model for cornea wound healing
Uploaded: 2018-07-10T14:30:00.000+00:00
Duration: 7 min 28 s
Description: Watch this Scientific Journal Video about Corneal Epithelial Abrasion with Ocular Burr As a Model for Cornea Wound Healing at JoVE.com

We would like to thank Kaisa Ikkala for her invaluable technical assistance and insightful help when actualizing this method as well as later on during implementation to our central research questions. We would also like to thank the Laboratory Animal Center and Anna Meller for her help with planning the guidelines of the veterinary work.

All experiments are approved by the national animal experiment board.
1. Preparations
<ol>
	<li>Prepare all solutions and keep in room temperature, unless otherwise indicated. Follow waste disposal regulations to dispose used materials and solutions.</li>
	<li>Use NMRI and ICR outbred stocks, between ages 4-12 weeks and either gender. If using the C57BL/6 strain, follow the ketamine-medetomidine preparation method in step 1.3.2. For other steps, follow the instructions described in 1.3.3.-1.3.5.</li>
	<li>Prepare the veterinary medicine fresh and in time before starting the operation. Carprofen will be used later, thus it is not necessary to prepare it at this point.
	<ol>
		<li>To prepare a solution of ketamine-medetomidine for anesthesia, mix 0.375 mL (stock concentration 50 mg/mL) of ketamine (Ketaminol Vet) with 0.25 mL (stock concentration 1 mg/mL) of medetomidine (Cepetor Vet) and add 1.25 mL of sterile 0.9% NaCl. Administer 0.15 mL/20 g of mouse weight (7.5 mg/kg, ketamine and 1 mg/kg, medetomidine).</li>
		<li>To prepare a more dilute solution of ketamine-medetomidine for anesthesia on C57BL/6 mice, mix 0.375 mL (stock concentration 50 mg/mL) of ketamine with 0.25 mL (stock concentration 1 mg/mL) of medetomidine and add 2.5 mL of sterile 0.9% NaCl. Administer 0.15 mL/20 g of mouse weight (3.75 mg/kg, ketamine and 0.5 mg/kg, medetomidine). 
		NOTE: This is an alternative step, only for adult C57BL/6 mice.</li>
		<li>To prepare buprenorfin for analgesia, add 0.1 mL (stock concentration 0.3 mg/mL) of buprenorfin to 1.9 mL of sterile 0.9% NaCl. Administer 0.2 mL/20 g mouse weight (0.15 mg/kg).</li>
		<li>To prepare atipamezole for discontinuing anesthesia, add 0.1 mL (stock concentration 5 mg/mL) of atipamezole to 4.9 mL of 0.9% NaCl. Administer 0.15 mL/20 g mouse weight (0.75 mg/kg).</li>
		<li>For analgesia during post-anesthesia use carprofen. Add 0.1 mL (stock concentration 50 mg/mL) of carprofen to 4.9 mL of 0.9% NaCl. Administer 0.15 mL/20 g of mouse weight (7.5 mg/kg).</li>
	</ol>
	</li>
	<li>Prepare fluorescent staining solution
	<ol>
		<li>For visualizing the abrasion under the Cobalt blue light (Figure 1), use a fluorescent solution. Prepare a 0.1% fluorescein solution by first measuring 10 mg of fluorescein salt with a fine scale and then add it to 10 mL in phosphate-buffered saline solution (PBS).</li>
		<li>Protect the fluorescein solution from light and shake for 5 minutes on a shaker. 
		NOTE: The fluorescein solution is light-sensitive; keep the solution covered from light. This solution can be stored in +4&#160;&#176;C for 2-3 days. For use, it is helpful to place the fluorescein solution in a dropper bottle that is used for eye drops. Use a sterile-filter when moving the solution to the dropper bottle.</li>
	</ol>
	</li>
	<li>Clean the ocular burr tip before and after use; first with PBS and then with 70% EtOH. If making an abrasion to several mice during the same operation, do the cleaning steps between each mouse.</li>
	<li>Put the heated plate on (+37&#160;&#176;C) and place a paper towel on it.</li>
</ol>
2. Corneal Abrasion
CAUTION: Use protective wear (gloves, lab coat) when handling mice. Weigh the mouse to estimate the volume of medication to administer.
<ol>
	<li>Use the one-handed scruffing method to handle the mouse23. Administer ketamine-medetomidine mixture intraperitoneally (i.p.) to the left lower abdomen. Place the mouse to an individual cage and wait for it to fall asleep. This typically takes less than 5 minutes.&#160;Then, administer first buprenorfin and then atipamezole via i.p. injections. Use the same handling technique. 
	NOTE: Once anesthesia is given, the protocol should not be paused at any moment.</li>
	<li>Before continuing, check that the heated plate is warm. Place the anesthetized mouse on the heated plate. The level of anesthesia is good if the tail reflex is absent, but the toe reflex is present. Check these reflexes by pinching the tail and any toe with fingers or forceps. Do not use excessive force. Anesthesia will last 20-25 minutes.</li>
	<li>To make an abrasion, use the cleaned ocular burr. Rotate the base of the burr to turn on the vibration that is essential to make the abrasion. Feel the vibration in hand, when the burr is functioning properly.
	<ol>
		<li>Open one eye at a time by holding the eyelids separately with fingers. Then firmly touch the burr to the cornea and move the instrument back and forth as well as sideways on the ocular surface. The burr vibration will perform the scratch; do not press, shake or tear the cornea. In the central cornea, 20 abrading movements on the surface are sufficient to induce the wound. Do not lift the burr and try to stay in the region of the cornea to be abraded. Acquiring a standard-sized abrasion will require several rounds of practice.</li>
	</ol>
	 
	NOTE: Corneal asepsis can be achieved before abrasion by using betadine ophthalmic solution.</li>
</ol>
3. Imaging the Abrasion
<ol>
	<li>Illuminate the abraded region using Cobalt blue pen light (Figure 1) and fluorescein solution. The ocular surface appears colorless in ambient light. After fluorescein application, the abraded region fluoresces in green when illuminating the eye with the pen light.
	<ol>
		<li>If needed, open the abraded eye similarly as in 2.3 and administer one drop of fluorescein solution from the dropper bottle. Wash the eye once with PBS or 0.9% NaCl to reduce background of the fluorescein. Use a dropper bottle or a pipette for washing. Aspirate the liquid away with a soft wipe and clean the eyelashes, if needed. The excess liquid usually collects to the nasal border of the eye, close to the opening of the tear canal. 
		CAUTION: Avoid touching the cornea while cleaining as minor abrasions might be induced.</li>
	</ol>
	</li>
	<li>Take pictures of the corneal abrasion.
	<ol>
		<li>To obtain similar images during each imaging session, use steady attachment tools for the Cobalt blue pen light and the SLR camera (Figure 2). This protocol provides a setup that is easy to repeat (Figure 2).
		<ol>
			<li>To attach the pen light and the camera, use a table lamp with a flexible arm and clamp and an adjustable camera arm with a clamp, respectively. Cable-tie the pen light to the table lamp and position just above the mouse eye. Positioning the light differently might alter the visualization of the abrasion. A suggestion for positioning these tools is described in Figure 2.</li>
		</ol>
		</li>
		<li>Use the SLR camera to take pictures of the eye. Place the animal in a desired distance and position and focus the camera (1:12) in room light. After that, close all other lights except the Cobalt blue pen light. 
		NOTE: Tape the pen light handle down or use a rubber band to keep the light on all the time during imaging. This might help if the image becomes blurry. It is easiest to perform the imaging step with a colleague.</li>
	</ol>
	</li>
</ol>
4. Waking up after Corneal Abrasion
<ol>
	<li>Administer first buprenorfin and then atipamezole via i.p. injections. Use the same handling technique as in 2.1.</li>
	<li>Place a drop of antibacterial, fucidin acid eye ointment (Isathal) to both abraded and non-abraded eyes to moisturize and keep the ocular surface clean from bacteria after anesthesia.</li>
	<li>As the mouse will wake up in 5-20 minutes on the heated plate, observe the mouse during the post-anesthetic wake-up period. When it becomes mobile, place it in an individual cage. 
	NOTE: If waking up takes longer, place the mouse in the cage with a heated pad or place the entire cage on the heated plate and monitor the mouse regularly. While the anesthesia wears off, the mouse typically shakes and reaches upwards with the anterior body. This behavior should return to normal in 3-4 hours and then you can place the mouse back to a shared cage.</li>
	<li>Administer carprofen i.p. for analgesia and eye ointment on two consecutive days after the abrasion, similarly as in steps 2.2. and 4.2.</li>
</ol>
5. Imaging during Wound Healing
<ol>
	<li>In this protocol, use time points 18 h and 72 h after abrasion to observe the wound closure.
	<ol>
		<li>For consecutive imaging, anesthetize the mice as explained in 2.2.</li>
		<li>Take pictures as explained in 3.</li>
		<li>Wake up the animals with atipamezole i.p. and monitor the wake-up period as explained in 4.3. 
		NOTE: If not continuing the follow-up, euthanize the mouse right after 5.1.2. Euthanasia is described in 6.1.</li>
	</ol>
	</li>
</ol>
6. Cornea Collection and Paraffin Embedding
<ol>
	<li>After the last time point, sacrifice the animal. Place the mouse in a chamber that can be filled with CO2. Fill the chamber air with CO2. When the animal is immobile, dislocate the cervical vertebrae by pulling firmly from the base of the tail and pressing the neck region down at the same time. 
	CAUTION: Always follow the guidelines of the local animal welfare body.</li>
	<li>Collect the whole eyeball by cutting an opening in the skin to the side of the eye with dissection scissors. Place the scissors under the eyeball and cut the ocular nerve to pop the eyeball out of the orbit. Use forceps if the eye does not come out easily.
	<ol>
		<li>Make a hole on the back of the eye, on the retinal side, with a 26G needle to allow free penetration of solutions inside the eye. 
		NOTE: Do not let the eye become dry, instead, place it in PBS until continuing with the protocol.</li>
	</ol>
	</li>
	<li>Collect the eyeball in a 2 mL tube with PBS. Do not pause the experiment at this point.</li>
	<li>Remove PBS and fix eyeballs in 4% paraformaldehyde (PFA) in +4&#160;&#176;C for 4-5 hours.</li>
	<li>Move the fixed eyeball to a tissue cassette for tissue processing. Use a tissue processing machine and the following program:
	<ol>
		<li>Rinse with PBS.</li>
		<li>Incubate 2 x 45 min in 70% ethanol at room temperature.</li>
		<li>Incubate 2 x 1 h in 94% ethanol at room temperature.</li>
		<li>Incubate 3 x 45 min in absolute ethanol at room temperature.</li>
		<li>Incubate 3 x 1 h in xylene, at room temperature and last xylene at 37&#160;&#176;C.</li>
		<li>Incubate 3 x 1 h in paraffin wax at 60&#160;&#176;C.</li>
	</ol>
	</li>
	<li>After tissue processing, embed the eyeball into liquid paraffin (+ 60&#160;&#176;C) using a tissue embedding block. Place the eye inside the block so that it is looking sideways and the peripheral border is facing upwards. Let the block solidify on a cool plate for 5-10 minutes.</li>
</ol>
7. Paraffin Sections of the Cornea
<ol>
	<li>Prepare the microtome for sectioning according to the institution&#39;s or manufacturer&#39;s instructions. 
	CAUTION: Be careful when handling the sharp microtome blade.</li>
	<li>Place one water bath with room temperature ultrapure water beside the microtome and another with ultrapure water heated to +50&#160;&#176;C.</li>
	<li>Make 5 &#181;m thick sections of the eyeball. 
	NOTE: The lens breaks easily during sectioning. To avoid this, wipe the cutting surface with a moist tissue each time a new row of sections is started. Use tap water to moisten the tissue.</li>
	<li>Place the sections beside each other in the room temperature water bath and then move them to the hot water bath with the help of a glass slide. Stretch the sections in the hot water bath until all wrinkles relax and the sections become matte.</li>
	<li>Dry the glass slides with sections overnight in +37&#160;&#176;C.</li>
	<li>Attach the sections to the slides by keeping the slide for 1 minute on a heated plate in +60&#160;&#176;C. After this, the sections can be directly processed for staining, immunohistological methods or stored in +4&#160;&#176;C.</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+mouse+model+for+the+study+of+recurrent+corneal+epithelial+erosions:+alpha9beta1+integrin+implicated+in+progression+of+the+disease.">A mouse model for the study of recurrent corneal epithelial erosions: alpha9beta1 integrin implicated in progression of the disease.</a> Investigative Ophthalmology &amp; Visual Science. 45 (6), 1775-1788 (2004).</li></ol>

The authors have nothing to disclose.

Wounding methods are popular tools to study different aspects of corneal homeostasis and pathologies. The abrasion model offers a well-controlled method to address relevant problems in ophthalmology. However, certain critical points in the protocol are worth emphasizing. Notably, the details outlined regarding the veterinary medicine, wound healing timeline, and outcome are optimized for use with outbred NMRI and ICR stocks, but may vary among strains of mice26. With this protocol, the experimenta...

Epithelial layers of numerous organs are exposed to injuries. However, they also contain the ability to compensate for tissue loss through wound healing. The cornea offers an excellent model to study wound healing. It forms the external surface of the eye and provides a protective layer for the sensitive ocular machinery. Thus, cornea functions as a physical barrier to pathogens and water loss. It is composed of three layers; epithelium, stroma and endothelium. The epithelium of the cornea makes up the outermost layer of the cornea. Epithelial cells maintain the barrier function of the cornea by adhering strictly to each other through tight junctions1,2,3. An acellular corneal basement membrane, the Bowman&#39;s membrane, separates the epithelium from the extensive stroma, which contains refractory keratocytes. Under the stroma, endothelial cells channel nutrients, water, and oxygen to the upper layer.
Corneal abrasions are very common in the clinic4. Injuries to the cornea are diverse, but are largely caused by small particles such as dust or sand, scratches, or other foreign objects. The protocol described here aims at reproducing a clinically relevant type of corneal epithelial abrasion. In doing so, this protocol provides a controllable and seminal method for clinicians and corneal scientists to implement in their own studies. We have performed an in vivo injury repair assay on the murine cornea by abrading the tissue with a dulled ocular burr, the Algerbrush II. Here, we target the abrasion only to the central corneal epithelium and leave the other parts of the organ without damage. Thus, the protocol is ideal to study corneal epithelial cell dynamics or the basement membrane during re-epithelialization, cell migration, proliferation and differentiation in vivo5. Recently, this model was used to analyze progenitor cell dynamics in the murine cornea as well as to unveil the capacity of the differentiated corneal epithelial cells in re-establishing the corneal stem cell niche after injury6,7. Following abrasion, the cornea returns to its normal transparency and tensile strength. Interestingly, an in vitro study indicated that re-epithelialization occurs without increased cell proliferation8. This protocol describes the timeline of uninterrupted healing in the murine cornea. The method is thus applicable to test the effect of drugs on healing patterns and speed.
The cornea has been extensively used for wound healing studies. However, many studies have relied on other models of injury. A well-established model of corneal injury is the alkaline burn that is performed by applying sodium hydroxide (NaOH) with or without filter paper on the corneal surface9. Alkaline exposure results in a large and diffuse injury that affects not only the corneal epithelium, but also the conjunctiva and stroma9,10. Strong alkaline solutions have been shown to induce corneal ulcers, opacification, and neovascularization9. Inflammatory cells invade the stroma typically within 6 h and remain there until 24 h11. Thus, alkaline injury is an advisable method in studies related to stromal activation. Another type of chemical injury can be inflicted by applying dimethyl sulfoxide (DMSO) on the cornea9,10. Other commonly used injury models include incisional wounds that penetrate through the stroma and keratectomy wounds, which are limited to the upper portion of the stroma14,15. These methods are also useful to answer questions regarding stromal wound healing. Different injury models have their own advantages and disadvantages. Abrasion, or debridement, of the corneal epithelium was first developed using dulled scalpels or blades on ex vivo corneas16. This method has later been used in vivo on mouse, rat, and rabbit17,18,19,20,21,22. Using the ocular burr (Figure 1), we remove only a selected region of the epithelium, leaving the rest of the epithelium unaffected. This way, it is possible to precisely target the epithelial removal to different parts of the cornea. In addition, the abrasion size can be assessed with fluorescein staining. Furthermore, here we follow abrasion closure during the healing period.
This method poses several advantages, i) including precise location of abrasion site, which is not possible with chemical injury, ii) the abrasion is quick to perform, and iii) it is non-invasive. Herein, we describe the method using the outbred NMRI mouse as a model, however this could be applied to the vast array of mouse genetic models, as well as to the rat and rabbit, which are common models used to study human corneal disruption.

<table><tbody><tr><td>NMRI mouse</td><td>Envigo</td><td>275</td><td></td></tr><tr><td>0.9% NaCl</td><td></td><td></td><td>use sterile</td></tr><tr><td>Medetomidine</td><td>Vetmedic</td><td>Vnr087896</td><td>Market name: Cepetor Vet</td></tr><tr><td>Ketamine</td><td>Intervet</td><td>Vnr511485</td><td>Market name: Ketaminol Vet</td></tr><tr><td>Buprenorfin</td><td>Invidior</td><td>3015248</td><td>Market name: Temgesic</td></tr><tr><td>Atipamezol</td><td>Orion Pharma</td><td>Vnr471953</td><td>Market name: Antisedan Vet</td></tr><tr><td>Carprofen</td><td>Norbrook</td><td>Vnr027579</td><td>Market name: Norocarp Vet</td></tr><tr><td>1% fucidin acid eye ointment</td><td>Dechra</td><td>Vnr080899</td><td>Market name: Isathal</td></tr><tr><td>Fluorescein salt</td><td>Sigma-Aldrich</td><td>F6377</td><td></td></tr><tr><td>Phosphate-buffered saline solution</td><td></td><td></td><td>PBS</td></tr><tr><td>Algerbrush ii ocular burr (0.5 mm tip)</td><td>Algerbrush</td><td>6.39768E+11</td><td></td></tr><tr><td>Cobalt Blue pen light</td><td>SP Services</td><td>DE/003</td><td></td></tr><tr><td>Hot plate</td><td>Kunz Instruments</td><td>2007-0217</td><td></td></tr><tr><td>Digital SLR camera</td><td>Nikon</td><td>D80</td><td></td></tr><tr><td>Adjustable camera arm and clamp</td><td>Neewer</td><td>10086132</td><td>Height 28 cm</td></tr><tr><td>Table lamp with a flexible arm and a clamp</td><td>Prisma</td><td></td><td></td></tr><tr><td>Soft wipe</td><td>KimtechScience</td><td>7552</td><td></td></tr><tr><td>CO2 chamber</td><td></td><td></td><td></td></tr><tr><td>Dissection toolset</td><td>Fine Science Tools</td><td></td><td></td></tr><tr><td>Syringes</td><td>Beckton Dickinson</td><td>303172</td><td></td></tr><tr><td>26G needles</td><td>Beckton Dickinson</td><td>303800</td><td></td></tr><tr><td>2 mL Eppendorf tube</td><td>Sarstedt</td><td>689</td><td></td></tr><tr><td>Tissue casette</td><td>Sakura Finetech</td><td>4118F</td><td></td></tr><tr><td>Tissue processing machine ASP200S</td><td>Leica</td><td></td><td></td></tr><tr><td>Xylene</td><td>VWR</td><td>UN1307</td><td></td></tr><tr><td>Paraffin wax</td><td>Millipore</td><td>K95523361</td><td></td></tr><tr><td>Tissue embedding mold 32 x 25 x 6 mm</td><td>Sakura Finetech</td><td>4123</td><td></td></tr><tr><td>Microtome</td><td>Microm</td><td>HM355</td><td></td></tr><tr><td>Water bath for sectioning</td><td>Orthex</td><td>60591</td><td></td></tr><tr><td>Water bath for sectioning</td><td>Leica</td><td>HI1210</td><td></td></tr><tr><td>Microtome blade</td><td>Feather</td><td>S35</td><td></td></tr><tr><td>Glass slide</td><td>Th.Geyer GmbH &amp;amp; Co.</td><td>7,695,019</td><td></td></tr><tr><td>Ultrapure water</td><td>Millipore</td><td>MPGP04001</td><td>MilliQ</td></tr><tr><td>Paraformaldehyde</td><td>Sigma-Aldrich</td><td>158127</td><td>PFA</td></tr></tbody></table>

corneal epithelial abrasion with ocular burr as a model for cornea wound healing

The murine cornea provides an excellent model to study wound healing. The cornea is the outermost layer of the eye, and thus is the first defense to injury. In fact, the most common type of eye injury found in clinic is a corneal abrasion. Here, we utilize an ocular burr to induce an abrasion resulting in removal of the corneal epithelium in vivo on anesthetized mice. This method allows for targeted and reproducible epithelial disruption, leaving other areas intact. In addition, we describe the visualization of the abraded epithelium with fluorescein staining and provide concrete advice on how to visualize the abraded cornea. Then, we follow the timeline of wound healing 0, 18, and 72 h after abrasion, until the wound is re-epithelialized. The epithelial abrasion model of corneal injury is ideal for studies on epithelial cell proliferation, migration and re-epithelialization of the corneal layers. However, this method is not optimal to study stromal activation during wound healing, because the ocular burr does not penetrate to the stromal cell layers. This method is also suitable for clinical applications, for example, pre-clinical test of drug effectiveness.

This protocol describes a model to inflict an abrasion injury to the mouse cornea and suggests how to follow and visualize the healing process after abrasion. Recently, we employed this method to study the role of corneal epithelial progenitor cells during wound healing6. The use of established tools is the key to a successful abrasion experiment. We, and others, have used the Algerbrush II ocular burr (Figure 1) to perform the abrasio...

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Corneal Epithelial Abrasion with Ocular Burr As a Model for Cornea Wound Healing

This protocol describes a method to inflict an abrasion to the ocular surface of the mouse, and to follow the wound healing process thereafter. The protocol takes advantage of an ocular burr to partially remove the surface epithelium of the eye in anaesthetized mice.

The murine cornea provides an excellent model to study wound healing. The cornea is the outermost layer of the eye, and thus is the first defense to ...

corneal-epithelial-abrasion-with-ocular-burr-as-model-for-cornea

Research

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Medicine

20.3K Views.  University of Helsinki. This protocol describes a method to inflict an abrasion to the ocular surface of the mouse, and to follow the wound healing process thereafter. The protocol takes advantage of an ocular burr to partially remove the surface epithelium of the eye in anaesthetized mice.

Video: Corneal Epithelial Abrasion with Ocular Burr As a Model for Cornea Wound Healing

Establishment of a Clinically Relevant Ex Vivo Mock Cataract Surgery Model for Investigating Epithelial Wound Repair in a Native Microenvironment

The major impediment to understanding how an epithelial tissue executes wound repair is the limited availability of models in which it is possible to follow and manipulate the wound response ex vivo in an environment that closely mimics that of epithelial tissue injury in vivo. This issue was addressed by creating a clinically relevant epithelial ex vivo injury-repair model based on cataract surgery. In this culture model, the response of the lens epithelium to wounding can be followed live in the cells&#8217; native microenvironment, and the molecular mediators of wound repair easily manipulated during the repair process. To prepare the cultures, lenses are removed from the eye and a small incision is made in the anterior of the lens from which the inner mass of lens fiber cells is removed. This procedure creates a circular wound on the posterior lens capsule, the thick basement membrane that surrounds the lens. This wound area where the fiber cells were attached is located just adjacent to a continuous monolayer of lens epithelial cells that remains linked to the lens capsule during the surgical procedure. The wounded epithelium, the cell type from which fiber cells are derived during development, responds to the injury of fiber cell removal by moving collectively across the wound area, led by a population of vimentin-rich repair cells whose mesenchymal progenitors are endogenous to the lens1. These properties are typical of a normal epithelial wound healing response. In this model, as in vivo, wound repair is dependent on signals supplied by the endogenous environment that is uniquely maintained in this ex vivo culture system, providing an ideal opportunity for discovery of the mechanisms that regulate repair of an epithelium following wounding.

Establishment of a Clinically Relevant Ex Vivo Mock Cataract Surgery Model for Investigating Epithelial Wound Repair in a Native Microenvironment

Described here is the establishment of a clinically relevant ex vivo mock cataract surgery model that can be used to investigate mechanisms of the injury response of epithelial tissues within their native microenvironment.

The major impediment to understanding how an epithelial tissue executes wound repair is the limited availability of models in which it is possible to ...

Developmental Biology

Investigating Scarless Tissue Regeneration in Embryonic Wounded Chick Corneas

Chick embryonic corneal wounds display a remarkable capacity to fully and rapidly regenerate, whereas adult wounded corneas experience a loss of transparency due to fibrotic scarring. The tissue integrity of injured embryonic corneas is intrinsically restored with no detectable scar formation. Given its accessibility and ease of manipulation, the chick embryo is an ideal model for studying scarless corneal wound repair. This protocol demonstrates the different steps involved in wounding the cornea of an embryonic chick in ovo. First, eggs are windowed at early embryonic ages to access the eye. Second, a series of in ovo physical manipulations to the extraembryonic membranes are conducted to ensure access to the eye is maintained through later stages of development, corresponding to when the three cellular layers of the cornea are formed. Third, linear cornea wounds that penetrate the outer epithelial layer and the anterior stroma are made using a microsurgical knife. The regeneration process or fully restored corneas can be analyzed for regenerative potential using various cellular and molecular techniques following the wounding procedure. Studies to date using this model have revealed that wounded embryonic corneas display activation of keratocyte differentiation, undergo coordinated remodeling of ECM proteins to their native three-dimensional macrostructure, and become adequately re-innervated by corneal sensory nerves. In the future, the potential impact of endogenous or exogenous factors on the regenerative process could be analyzed in healing corneas by using developmental biology techniques, such as tissue grafting, electroporation, retroviral infection, or bead implantation. The current strategy identifies the embryonic chick as a crucial experimental paradigm for elucidating the molecular and cellular factors coordinating scarless corneal wound healing.

The present protocol demonstrates the different steps involved in wounding the cornea of an embryonic chick in ovo. The regenerating or fully restored corneas can be analyzed for regenerative potential using various cellular and molecular techniques following the wounding procedure.&#160;

Chick embryonic corneal wounds display a remarkable capacity to fully and rapidly regenerate, whereas adult wounded corneas experience a loss of ...

A Simple Mechanical Procedure to Create Limbal Stem Cell Deficiency in Mouse

Limbal stem cell deficiency (LSCD) is a state of malfunction or loss of limbal epithelial stem cells, after which the corneal epithelium is replaced with conjunctiva. Patients suffer from recurrent corneal defects, pain, inflammation, and loss of vision.
Previously, a murine model of LSCD was described and compared to two other models. The goal was to produce a consistent mouse model of LSCD that both mimics the phenotype in humans and lasts long enough to make it possible to study the disease pathophysiology and to evaluate new treatments. Here, the technique is described in more detail.
A motorized tool with a rotating burr has been designed to remove the rust rings from the corneal surface or to smooth the pterygium bed in patients. It is a suitable device to create the desired LSCD model. It is a readily available, easy-to-use tool with a fine tip that makes it appropriate for working on small eyes, as in mice. Its application prevents unnecessary trauma to the eye and it does not result in unwanted injuries, as often is the case with chemical injury models. As opposed to a blunt scraper, it removes the epithelium with the basement membrane. In this protocol, the limbal area was abraded two times, and then the whole corneal epithelium was shaved from limbus to limbus. To avoid stroma injury, care was taken not to brush the corneal surface once the epithelium was already removed.

The following article provides an easy, reproducible technique to effectively create a sustainable mouse model of limbal stem cell deficiency (LSCD). This animal model is useful in testing and comparing the efficacy of treatments for limbal stem cell diseases.

Limbal stem cell deficiency (LSCD) is a state of malfunction or loss of limbal epithelial stem cells, after which the corneal epithelium is replaced with ...

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 ...

An Epithelial Abrasion Model for Studying Corneal Wound Healing

The cornea is critical for vision, accounting for about two-thirds of the refractive power of the eye. Crucial to the role of the cornea in vision is its transparency. However, due to its external position, the cornea is highly susceptible to a wide variety of injuries that can lead to the loss of corneal transparency and eventual blindness. Efficient corneal wound healing in response to these injuries is pivotal for maintaining corneal homeostasis and preservation of corneal transparency and refractive capabilities. In events of compromised corneal wound healing, the cornea becomes vulnerable to infections, ulcerations, and scarring. Given the fundamental importance of corneal wound healing to the preservation of corneal transparency and vision, a better understanding of the normal corneal wound healing process is a prerequisite to understanding impaired corneal wound healing associated with infection and disease. Toward this goal, murine models of corneal wounding have proven useful in furthering our understanding of the corneal wound healing mechanisms operating under normal physiological conditions. Here, a protocol for creating a central corneal epithelial abrasion in mouse using a trephine and a blunt golf club spud is described. In this model, a 2 mm diameter circular trephine, centered over the cornea, is used to demarcate the wound area. The golf club spud is used with care to debride the epithelium and create a circular wound without damaging the corneal epithelial basement membrane. The resulting inflammatory response proceeds as a well-characterized cascade of cellular and molecular events that are critical for efficient wound healing. This simple corneal wound healing model is highly reproducible and well-published and is now being used to evaluate compromised corneal wound healing in the context of disease.

Here, a protocol for creating a central corneal epithelial abrasion wound in the mouse using a trephine and a blunt golf club spud is described. This corneal wound healing model is highly reproducible and is now being used to evaluate compromised corneal wound healing in the context of diseases.

The cornea is critical for vision, accounting for about two-thirds of the refractive power of the eye. Crucial to the role of the cornea in vision is its ...

Ex Vivo and In Vivo Animal Models for Mechanical and Chemical Injuries of Corneal Epithelium

Corneal injury to the ocular surface, including chemical burn and trauma, may cause severe scarring, symblepharon, corneal limbal stem cells deficiency, and result in a large, persistent corneal epithelial defect. Epithelial defect with the following corneal opacity and peripheral neovascularization result in irreversible visual impairment and hinder future management, especially keratoplasty. Since the animal model can be used as an effective drug development platform, models of corneal injury to the mouse and alkali burn to rabbit corneal epithelium are developed here. New Zealand white rabbit is used in the alkali burn model. Different concentrations of sodium hydroxide can be applied onto the central circular area of the cornea for 30 s under intramuscular and topical anesthesia. After copious isotonic normal saline irrigation, residual loose corneal epithelium was removed with corneal burr deep down to the Bowman's layer within this circular area. Wound healing was documented by fluorescein staining under Cobalt blue light. C57BL/6 mice were used in the traumatic model of murine corneal epithelium. The murine central cornea was marked using a skin punch, 2 mm in diameter, and then debrided by a corneal rust ring remover with a 0.5 mm burr under a stereomicroscope. These models can be prospectively used to validate the therapeutic effect of eye drops or mixed agents such as stem cells, which potentially facilitate corneal epithelial regeneration. By observing corneal opacity, peripheral neovascularization, and conjunctival congestion with stereomicroscope and imaging software, therapeutic effects in these animal models can be monitored.

Ex Vivo and In Vivo Animal Models for Mechanical and Chemical Injuries of Corneal Epithelium

Here, animal models based on mouse and rabbit are developed for mechanical and chemical injury of corneal epithelium to screen new therapeutics and the underlying mechanism.

Corneal injury to the ocular surface, including chemical burn and trauma, may cause severe scarring, symblepharon, corneal limbal stem cells deficiency, ...

Corneal Sensory Fiber Damage Protocols for Axon Regeneration Studies

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.

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.

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 ...

Neuroscience

Mongolian Gerbils as an Animal Model of Wound Healing

Corneal wound healing studies have been conducted for a long time and have helped to reduce suffering and develop treatments that contribute to improving patients' eye health. Historically, corneal healing has been studied in rodents such as mice and rats, but these models might not completely mimic human disorders. However, information on other rodents such as Mongolian gerbils (Meriones unguiculatus) is scant in corneal research.
Here, we describe a technique to develop a novel animal model for studying corneal healing after photorefractive keratectomy. Due to the limited literature available on the cornea of M. unguiculatus, we also describe a histological analysis of the normal cornea. These research techniques can also be employed in the study of eye diseases because of the similarity between the corneas of Mongolian gerbils and humans in terms of genetics, anatomy, and physiology.

This article describes a new animal model developed to study the anatomy and histology of the cornea and its healing processes. This new animal model uses the Mongolian gerbil, which has a cornea with many similarities to the human cornea.

Corneal wound healing studies have been conducted for a long time and have helped to reduce suffering and develop treatments that contribute to improving ...

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 ...

Zebrafish Corneal Wound Healing: From Abrasion to Wound Closure Imaging Analysis

As the transparent surface of the eye, the cornea is instrumental for clear sight. Due to its location, this tissue is prone to environmental insults. Indeed, the eye injuries most frequently encountered clinically are those to the cornea. While corneal wound healing has been extensively studied in small mammals (i.e., mice, rats, and rabbits), corneal physiology studies have neglected other species, including zebrafish, despite zebrafish being a classic research model.
This report describes a method of performing a corneal abrasion on zebrafish. The wound is performed in vivo on anesthetized fish using an ocular burr. This method allows for a reproducible epithelial wound, leaving the rest of the eye intact. After abrasion, wound closure is monitored over the course of 3 h, after which the wound is reepithelialized. By using scanning electron microscopy, followed by image processing, the epithelial cell shape, and apical protrusions can be investigated to study the various steps during corneal epithelial wound closure.
The characteristics of the zebrafish model permit study of the epithelial tissue physiology and the collective behavior of the epithelial cells when the tissue is challenged. Furthermore, the use of a model deprived of the influence of the tear film can produce&#160;new answers regarding corneal response to stress. Finally, this model also allows the delineation of the cellular and molecular events involved in any epithelial tissue subjected to a physical wound. This method can be applied to the evaluation of drug effectiveness in preclinical testing.

This protocol focuses on damaging the ocular surface of zebrafish through abrasion to assess the subsequent wound closure at the cellular level. This approach exploits an ocular burr to partly remove the corneal epithelium and uses scanning electron microscopy to track changes in cell morphology during wound closure.

As the transparent surface of the eye, the cornea is instrumental for clear sight. Due to its location, this tissue is prone to environmental insults. ...

Corneal Epithelial Abrasion with Ocular Burr As a Model for Cornea Wound Healing

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Chapters in this video

Establishment of a Clinically Relevant Ex Vivo Mock Cataract Surgery Model for Investigating Epithelial Wound Repair in a Native Microenvironment

Investigating Scarless Tissue Regeneration in Embryonic Wounded Chick Corneas

A Simple Mechanical Procedure to Create Limbal Stem Cell Deficiency in Mouse

Ex Vivo Corneal Organ Culture Model for Wound Healing Studies

An Epithelial Abrasion Model for Studying Corneal Wound Healing

Ex Vivo and In Vivo Animal Models for Mechanical and Chemical Injuries of Corneal Epithelium

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

Mongolian Gerbils as an Animal Model of Wound Healing

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

Zebrafish Corneal Wound Healing: From Abrasion to Wound Closure Imaging Analysis