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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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>
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			<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 &#38; 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...

Watch this Scientific Journal Video about Corneal Epithelial Abrasion with Ocular Burr As a Model for Cornea Wound Healing at JoVE.com

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 ...

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20.2K 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

A Mouse Model of the Cornea Pocket Assay for Angiogenesis Study

A normal cornea is clear of vascular tissues. However, blood vessels can be induced to grow and survive in the cornea when potent angiogenic factors are administered 1. This uniqueness has made the cornea pocket assay one of the most used models for angiogenesis studies. The cornea composes multiple layers of cells. It is therefore possible to embed a pellet containing the angiogenic factor of interest in the cornea to investigate its angiogenic effect 2,3. Here, we provide a step by step demonstration of how to (I) produce the angiogenic factor-containing pellet (II) embed the pellet into the cornea (III) analyze the angiogenesis induced by the angiogenic factor of interest. Since the basic fibroblast growth factor (bFGF) is known as one of the most potent angiogenic factors 4, it is used here to induce angiogenesis in the cornea.

The cornea is unique in that it lacks vascular tissues. However, robust blood vessel growth and survival can be induced in the cornea by potent angiogenic factors. Therefore, the cornea can provide with us a valuable tool for angiogenic studies. This protocol demonstrates how to perform the mouse model of cornea pocket assay and how to assess the angiogenesis induced by angiogenic factors using this model.

A normal cornea is clear of vascular tissues. However, blood vessels can be induced to grow and survive in the cornea when potent angiogenic factors are ...

Murine Model of Wound Healing

Wound healing and repair are the most complex biological processes that occur in human life. After injury, multiple biological pathways become activated. Impaired wound healing, which occurs in diabetic patients for example, can lead to severe unfavorable outcomes such as amputation. There is, therefore, an increasing impetus to develop novel agents that promote wound repair. The testing of these has been limited to large animal models such as swine, which are often impractical. Mice represent the ideal preclinical model, as they are economical and amenable to genetic manipulation, which allows for mechanistic investigation. However, wound healing in a mouse is fundamentally different to that of humans as it primarily occurs via contraction. Our murine model overcomes this by incorporating a splint around the wound. By splinting the wound, the repair process is then dependent on epithelialization, cellular proliferation and angiogenesis, which closely mirror the biological processes of human wound healing. Whilst requiring consistency and care, this murine model does not involve complicated surgical techniques and allows for the robust testing of promising agents that may, for example, promote angiogenesis or inhibit inflammation. Furthermore, each mouse acts as its own control as two wounds are prepared, enabling the application of both the test compound and the vehicle control on the same animal. In conclusion, we demonstrate a practical, easy-to-learn, and robust model of wound healing, which is comparable to that of humans.

A murine model of cutaneous wound healing that can be used to assess therapeutic compounds in physiological and pathophysiological settings.

Wound healing and repair are the most complex biological processes that occur in human life. After injury, multiple biological pathways become activated. ...

Multimodal Quantitative Phase Imaging with Digital Holographic Microscopy Accurately Assesses Intestinal Inflammation and Epithelial Wound Healing

The incidence of inflammatory bowel disease, i.e., Crohn&#39;s disease and Ulcerative colitis, has significantly increased over the last decade. The etiology of IBD remains unknown and current therapeutic strategies are based on the unspecific suppression of the immune system. The development of treatments that specifically target intestinal inflammation and epithelial wound healing could significantly improve management of IBD, however this requires accurate detection of inflammatory changes. Currently, potential drug candidates are usually evaluated using animal models in vivo or with cell culture based techniques in vitro. Histological examination usually requires the cells or tissues of interest to be stained, which may alter the sample characteristics and furthermore, the interpretation of findings can vary by investigator expertise. Digital holographic microscopy (DHM), based on the detection of optical path length delay, allows stain-free quantitative phase contrast imaging. This allows the results to be directly correlated with absolute biophysical parameters. We demonstrate how measurement of changes in tissue density with DHM, based on refractive index measurement, can quantify inflammatory alterations, without staining, in different layers of colonic tissue specimens from mice and humans with colitis. Additionally, we demonstrate continuous multimodal label-free monitoring of epithelial wound healing in vitro, possible using DHM through the simple automated determination of the wounded area and simultaneous determination of morphological parameters such as dry mass and layer thickness of migrating cells. In conclusion, DHM represents a valuable, novel and quantitative tool for the assessment of intestinal inflammation with absolute values for parameters possible, simplified quantification of epithelial wound healing in vitro and therefore has high potential for translational diagnostic use.

Accurate assessment of anti-inflammatory effects is of utmost importance for the evaluation of potential new drugs for the treatment of inflammatory bowel disease. Digital holographic microscopy provides assessment of inflammation in murine and human colonic tissue samples as well as automated multimodal evaluation of epithelial wound healing in vitro.

The incidence of inflammatory bowel disease, i.e., Crohn&#39;s disease and Ulcerative colitis, has significantly increased over the last decade. The ...

Creation and Transplantation of an Adipose-derived Stem Cell (ASC) Sheet in a Diabetic Wound-healing Model

Artificial skin has achieved considerable therapeutic results in clinical practice. However, artificial skin treatments for wounds in diabetic patients with impeded blood flow or with large wounds might be prolonged. Cell-based therapies have appeared as a new technique for the treatment of diabetic ulcers, and cell-sheet engineering has improved the efficacy of cell transplantation. A number of reports have suggested that adipose-derived stem cells (ASCs), a type of mesenchymal stromal cell (MSC), exhibit therapeutic potential due to their relative abundance in adipose tissue and their accessibility for collection when compared to MSCs from other tissues. Therefore, ASCs appear to be a good source of stem cells for therapeutic use. In this study, ASC sheets from the epididymal adipose fat of normal Lewis rats were successfully created using temperature-responsive culture dishes and normal culture medium containing ascorbic acid. The ASC sheets were transplanted into Zucker diabetic fatty (ZDF) rats, a rat model of type 2 diabetes and obesity, that exhibit diminished wound healing. A wound was created on the posterior cranial surface, ASC sheets were transplanted into the wound, and a bilayer artificial skin was used to cover the sheets. ZDF rats that received ASC sheets had better wound healing than ZDF rats without the transplantation of ASC sheets. This approach was limited because ASC sheets are sensitive to dry conditions, requiring the maintenance of a moist wound environment. Therefore, artificial skin was used to cover the ASC sheet to prevent drying. The allogenic transplantation of ASC sheets in combination with artificial skin might also be applicable to other intractable ulcers or burns, such as those observed with peripheral arterial disease and collagen disease, and might be administered to patients who are undernourished or are using steroids. Thus, this treatment might be the first step towards improving the therapeutic options for diabetic wound healing.

Creation and Transplantation of an Adipose-derived Stem Cell ASC Sheet in a Diabetic Wound-healing Model

Adipose-derived stem cells (ASCs) are easily isolated and harvested from the fat of normal rats. ASC sheets can be created using cell-sheet engineering and can be transplanted into Zucker diabetic fatty rats exhibiting full-thickness skin defects with exposed bone and then covered with a bilayer of artificial skin.

Artificial skin has achieved considerable therapeutic results in clinical practice. However, artificial skin treatments for wounds in diabetic patients ...

Development of a Direct Pulp-capping Model for the Evaluation of Pulpal Wound Healing and Reparative Dentin Formation in Mice

Dental pulp is a vital organ of a tooth fully protected by enamel and dentin. When the pulp is exposed due to cariogenic or iatrogenic injuries, it is often capped with biocompatible materials in order to expedite pulpal wound healing. The ultimate goal is to regenerate reparative dentin, a physical barrier that functions as a "biological seal" and protects the underlying pulp tissue. Although this direct pulp-capping procedure has long been used in dentistry, the underlying molecular mechanism of pulpal wound healing and reparative dentin formation is still poorly understood. To induce reparative dentin, pulp capping has been performed experimentally in large animals, but less so in mice, presumably due to their small sizes and the ensuing technical difficulties. Here, we present a detailed, step-by-step method of performing a pulp-capping procedure in mice, including the preparation of a Class-I-like cavity, the placement of pulp-capping materials, and the restoration procedure using dental composite. Our pulp-capping mouse model will be instrumental in investigating the fundamental molecular mechanisms of pulpal wound healing in the context of reparative dentin in vivo by enabling the use of transgenic or knockout mice that are widely available in the research community.

We describe a step-by-step method of performing direct pulp capping on mice teeth for the evaluation of pulpal wound healing and reparative dentin formation in vivo.

Dental pulp is a vital organ of a tooth fully protected by enamel and dentin. When the pulp is exposed due to cariogenic or iatrogenic injuries, it is ...

Microscopy Based Methods for the Assessment of Epithelial Cell Migration During In Vitro Wound Healing

Cell migration is a mandatory aspect for wound healing. Creating artificial wounds on research animal models often results in costly and complicated experimental procedures, while potentially lacking in precision. In vitro culture of epithelial cell lines provides a suitable platform for researching the cell migratory behavior in wound healing and the impact of treatments on these cells. The physiology of epithelial cells is often studied in non-confluent conditions; however, this approach may not resemble natural wound healing conditions. Disrupting the epithelium integrity by mechanical means generates a realistic model, but may impede the application of molecular techniques. Consequently, microscopy based techniques are optimal for studying epithelial cell migration in vitro. Here we detail two specific methods, the artificial wound scratch assay and the artificial migration front assay, that can obtain quantitative and qualitative data, respectively, on the migratory performance of epithelial cells.

Microscopy Based Methods for the Assessment of Epithelial Cell Migration During  In Vitro Wound Healing

This manuscript describes how incision-like lesions made on cultured epithelial cell monolayers conveniently model wound healing in vitro, allowing for imaging by confocal or laser scanning microscopy, and which can provide high-quality quantitative and qualitative data for studying both cell behavior and the mechanisms involved in migration.

Cell migration is a mandatory aspect for wound healing. Creating artificial wounds on research animal models often results in costly and complicated ...

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 ...

Targeting Gray Rami Communicantes in Selective Chemical Lumbar Sympathectomy

Chemical lumbar sympathectomy (CLS) is a commonly used, minimally invasive procedure for the treatment of conditions including ischemic diseases of the lower extremities, hyperhidrosis, etc. It is commonly practiced to position the puncture needle tip in front of the anterior fascia of the psoas major muscle and inject the inactivating agent around the sympathetic trunk, which is defined as conventional CLS. Although relatively rare, ureteropelvic damage is the most frequently reported complication of conventional CLS and can cause serious harm to patients. We found that injecting the inactivating agent behind the anterior fascia, which only targets gray rami communicantes, helped achieve therapeutic efficacy in&#160;vasodilation, sweat reduction, and pain relief comparable to conventional CLS, and serious complications were largely reduced. We define this procedure as selective CLS. Here, we present a protocol of selective CLS. The precise needle tract and accurate evaluation of the spreading of the contrast agent are critical to ensure that the drug is injected behind the anterior fascia of the psoas major muscle. The needle tip is at approximately one-third the dividing line of the vertebral body in the lateral view of a lumbar X-ray. The contrast is mainly confined around the needle tip and spreads outward and downward along the psoas muscle fibers. In this way, the anterior fascia provides a natural barrier for the ureteropelvic area, and the psoas major muscle provides a natural barrier for the lumbar nerve root. There are several highlights of this article, including 1) a detailed description of the selective CLS procedures, 2) an explanation of the anatomical basis for the implementation of selective CLS, and 3) an explanation of the differences between selective and conventional CLS.

We present a protocol for selective chemical lumbar sympathectomy (CLS), which inactivates only gray rami communicantes and not the sympathetic trunk. Selective CLS can help achieve therapeutic efficacy in vasodilation, sweat reduction, and pain relief that are comparable to conventional CLS, and serious complications, particularly ureteropelvic damages, can be reduced.

Chemical lumbar sympathectomy (CLS) is a commonly used, minimally invasive procedure for the treatment of conditions including ischemic diseases of the ...

A Porcine Corneal Endothelial Organ Culture Model Using Split Corneal Buttons

Experimental research on corneal endothelial cells is associated with several difficulties. Human donor corneas are scarce and rarely available for experimental investigations as they are normally needed for transplantation. Endothelial cell cultures often do not translate well to in vivo situations. Due to the biostructural characteristics of non-human corneas, stromal swelling during cultivation induces substantial corneal endothelial cell loss, which makes it difficult to perform cultivation for an extended period of time. Deswelling agents such as dextran are used to counteract this response. However, they also cause significant endothelial cell loss. Therefore, an ex vivo organ culture model not requiring deswelling agents was established. Pig eyes from a local slaughterhouse were used to prepare split corneal buttons. After partial corneal trephination, the outer layers of the cornea (epithelium, bowman layer, parts of the stroma) were removed. This significantly reduces corneal endothelial cell loss induced by massive stromal swelling and Descemet&#39;s membrane folding throughout longer cultivation periods and improves general preservation of the endothelial cell layer. Subsequent complete corneal trephination was followed by the removal of the split corneal button from the remaining eye bulb and cultivation. Endothelial cell density was assessed at follow-up times of up to 15 days after preparation (i.e., days 1, 8, 15) using light microscopy. The preparation technique used allows a better preservation of the endothelial cell layer enabled by less stromal tissue swelling, which results in slow and linear decline rates in split corneal buttons comparable to human donor corneas. As this standardized organo-typically cultivated research model for the first time allows a stable cultivation for at least two weeks, it is a valuable alternative to human donor corneas for future investigations of various external factors with regards to their effects on the corneal endothelium.

Here, a step-by-step protocol for the preparation and cultivation of porcine split corneal buttons is presented. As this organo-typically cultivated organ culture model shows cell death rates within 15 days, comparable to human donor corneas, it represents the first model allowing long-term cultivation of non-human corneas without adding toxic dextran.

Experimental research on corneal endothelial cells is associated with several difficulties. Human donor corneas are scarce and rarely available for ...

Development of a Noninvasive, Laser-Assisted Experimental Model of Corneal Endothelial Cell Loss

Nd:YAG lasers have been used to perform noninvasive intraocular surgery, such as capsulotomy for several decades now. The incisive effect relies on the optical breakdown at the laser focus. Acoustic shock waves and cavitation bubbles are generated, causing tissue rupture. Bubble sizes and pressure amplitudes vary with pulse energy and position of the focal point. In this study, enucleated porcine eyes were positioned in front of a commercially available Nd:YAG laser. Variable pulse energies as well as different positions of the focal spots posterior to the cornea were tested. Resulting lesions were evaluated by two-photon microscopy and histology to determine the best parameters for an exclusive detachment of corneal endothelial cells (CEC) with minimum collateral damage. The advantages of this method are the precise ablation of CEC, reduced collateral damage, and above all, the non-contact treatment.

Here, we present a protocol to detach corneal endothelial cells (CEC) from Descemet&#8217;s membrane (DM) using a neodymium:YAG (Nd:YAG) laser as an ex vivo disease model for bullous keratopathy (BK).

Nd:YAG lasers have been used to perform noninvasive intraocular surgery, such as capsulotomy for several decades now. The incisive effect relies on the ...