Name: zebrafish corneal wound healing: from abrasion to wound closure imaging analysis
Uploaded: 2022-03-01T15:00:00.000+00:00
Duration: 8 min 19 s
Description: Watch this Scientific Journal Video about Zebrafish Corneal Wound Healing: From Abrasion to Wound Closure Imaging Analysis at JoVE.com

The authors thank Pertti Panula for the access to the Zebrafish unit and Henri Koivula for the guidance and help with the zebrafish experiments. This research was supported by the Academy of Finland, the Jane and Aatos Erkko Foundation, the Finnish Cultural Foundation, and the ATIP-Avenir Program. Imaging was performed at the Electron Microscopy unit and the Light Microscopy Unit, Institute of Biotechnology, supported by HiLIFE and Biocenter Finland.

All experiments were approved by the national animal experiment board.
1. Preparations
<ol>
	<li>Prepare the tricaine stock solution used for anesthesia26 in advance (0.4% stock solution used in this protocol). Use gloves and keep the materials in a fume hood whenever possible.
	<ol>
		<li>For 50 mL of a 0.4% solution, weigh 200 mg of tricaine powder into a 50 mL tube. Dissolve the powder in approximately 45 mL of double-distilled water.</li>
		<li>Adjust the pH of the tricaine stock solution to 7 with 1 M Tris (pH 8.8, ~1.25 mL). Add the Tris solution to the tricaine stock in aliquots, mix the stock thoroughly after each aliquot, and check the pH after each addition of Tris.</li>
	</ol>
	</li>
	<li>Before the experiment, prepare a 0.02% working solution of tricaine.
	<ol>
		<li>Thaw 2 mL of 0.4% stock solution and add to 40 mL of system water (final concentration 0.02%). Place the solution in a small container.</li>
	</ol>
	</li>
	<li>Before the experiment, prepare the recovery water containing analgesic. Use gloves and keep the materials in fume hood whenever possible.
	<ol>
		<li>For one liter of recovery water, weigh 2.5 mg of lidocaine hydrochloride powder and dissolve it in fresh system water. Check the pH and adjust to 7 if necessary.</li>
	</ol>
	</li>
	<li>Before the experiment, prepare the fixing solution (2.5% glutaraldehyde in 0.1 M sodium phosphate (Na-PO4) solution at pH 7.4). Use gloves and keep the materials in a fume hood.
	<ol>
		<li>For 10 mL of the fixing solution, pipette 5 mL of 0.2 M Na-PO4 into a tube. Add 0.5 mL of 50% glutaraldehyde, and add double-distilled water to obtain the final volume of 10 mL. Protect the solution from light, and keep it on ice or in the fridge prior to use. 
		NOTE: If samples must be collected for several hours after wounding, prepare the fixing solution just prior to use.</li>
	</ol>
	</li>
	<li>Prepare the equipment for wounding (Figure 1).
	<ol>
		<li>Fill the recovery tanks or smaller containers with system water.</li>
		<li>Have the ophthalmic burr ready. Check that the burr tip is clean. If needed, remove cell debris with a moist cotton swab.</li>
		<li>Make an incision to the side of a soft sponge, and moisten the sponge with system water. Place the sponge on the base/stage of a dissecting microscope. Ensure enough working space for using the burr and enough illumination from the sides and/or above to see the eye surface properly.</li>
	</ol>
	</li>
</ol>
2. Anesthesia
<ol>
	<li>Transfer a fish from the tank to the 0.02% tricaine solution with a net as gently as possible.</li>
	<li>Monitor the anesthesia, checking for lack of response to light mechanical stimulus. 
	NOTE: For consistent anesthesia, a 2 min exposure to tricaine is used prior to abrasion with adult wild-type AB fish. With fish of other genetic background, a different duration may be needed.</li>
</ol>
3. Abrasion
<ol>
	<li>Gently place the anesthetized fish with a spoon into the incision on the sponge, head protruding from the sponge surface.</li>
	<li>Turn on the burr, and focus the microscope view onto the eye surface.</li>
	<li>Carefully approach the eye surface with the burr tip. When touching the eye surface, start moving the burr tip on the eye surface with circular motion. Avoid sudden movement, as it might lead to the eye tilting in the socket and the burr tip to slip.</li>
	<li>When the abrasion is done, carefully place the fish in system water containing the analgesic for recovery.</li>
	<li>Clean the burr right after use with a moist cotton swab.</li>
</ol>
4. Collecting samples
<ol>
	<li>At the desired time point, pick the fish up with a net and place it in 0.02% tricaine solution. Keep the animal in the solution until the opercular movement has ceased completely, and the fish does not react to touching.</li>
	<li>Place the fish on a Petri dish with a spoon, and hold it with tweezers. Decapitate the fish with dissecting scissors. Avoid making any scratches on the eye surface when handling the sample.</li>
	<li>Put the tissue into a sample tube containing 0.1 M Na-PO4.
	<ol>
		<li>Rinse the tissue by replacing the 0.1 M Na-PO4 with clean buffer so that no blood remains in the solution.</li>
	</ol>
	</li>
</ol>
5. Sample processing for electron microscopy
<ol>
	<li>Fix the tissue in 2.5% glutaraldehyde/0.1 M Na-PO4 (pH 7.4) for ~24 h at +4 &#176;C. Keep the sample on a rotating/shaking sample holder to ensure proper fixation.</li>
	<li>Remove the fixing solution and rinse the sample several times with 0.1 M Na-PO4.</li>
	<li>Dissect the sample at this point.
	<ol>
		<li>Place the sample onto a drop of 0.1 M Na-PO4 on a dissecting plate. If both eyes from the same fish must be imaged, cut the head sample into two with fine dissecting scissors.</li>
		<li>Alternatively, collect the eyes only by carefully placing the tips of fine tweezers into the eye socket from the side of the eye, taking extra care not to scratch the eye surface. Then, pull the eye out from the socket.</li>
		<li>Transfer the dissected sample into a tube containing 0.1 M Na-PO4. Ensure there is no extra tissue in the sample tube, as it may adhere to the top of the eye during sample processing.</li>
	</ol>
	</li>
	<li>Store the sample in 0.1 M Na-PO4 (maximum one week) at +4 &#176;C.</li>
	<li>Process the samples for electron microscopy imaging.
	<ol>
		<li>Postfix the samples in 2% osmium tetroxide in 0.1 M Na-PO4 buffer for 1 h at room temperature (RT).</li>
		<li>Wash the samples 3 times for 5 minutes each wash in 0.1 M Na-PO4 at RT.</li>
		<li>Dehydrate the samples successively in 30%, 50%, and 70% ethanol for 1 h in each solution at RT.</li>
		<li>Immerse the samples in 96% ethanol for 2-3 h at RT.</li>
		<li>Next, incubate the samples two times in 100% ethanol, first for 1 h and then in fresh 100% ethanol overnight at +4 &#176;C.</li>
		<li>Subject the samples to 30 cycles in an automated critical point dryer.</li>
	</ol>
	</li>
	<li>Embed and platinum-coat the samples.
	<ol>
		<li>Place an adhesive tab onto a mount. If the sample must be marked on top of the mount, leave a piece of tab cover paper on the tab and write the sample ID on the paper.</li>
		<li>Place the mount with the tab on the base of a dissecting microscope.</li>
		<li>Gently place the tissue sample on the mount with fine tweezers, cornea facing up.</li>
		<li>Coat the specimen with platinum using the appropriate device. After coating, store the samples at room temperature until imaging.</li>
	</ol>
	</li>
</ol>
6. Imaging (Figure 2)
<ol>
	<li>Operate the devices as advised in the user&#39;s manual and by imaging experts.</li>
	<li>Acquire images of the desired magnification, and use 2,000-2,500x images for analysis.</li>
	<li>Adjust the brightness and contrast so that there are no overexposed areas in the image, and cell borders and microridges are seen as clearly as possible. 
	NOTE: The position and angle of the tissue affect the brightness and contrast settings. They may need to be adjusted from sample to sample and between different regions of the tissue.</li>
</ol>
7. Measuring cell shape, size, and microridge pattern
<ol>
	<li>Open the TIFF image in Fiji ImageJ 1.5327. Set the scale using the scale bar of the image: create a line equal in size to the scale bar with the Line tool. Select Analyze | Set scale, and type in the known distance. Open the ROI manager from the Analyze | Tools menu.</li>
	<li>For cell size and roundness, select Analyze | Set measurements | Shape descriptors. Use the Magnifying glass tool to see the cells under magnification. Select cells with the Polygon tool, and add each selection to the ROI manager. Finally, measure the selected cells, and save the measurement.</li>
	<li>Microridge analysis (Figure 3 and Figure 4)
	<ol>
		<li>Ensure that the image is in 8-bit format from the Image | Type menu.</li>
		<li>Select a cell with the Polygon tool, and clear the background from Edit | Clear outside.</li>
		<li>Smoothen the image one to three times by selecting Process | Smooth, and adjust the brightness and contrast from Image | Adjust | Brightness/Contrast so that the microridges stand out as clearly as possible.</li>
		<li>Convolve the image from Process | Filters | Convolve, turn into binary from Process | Binary | Make binary, and skeletonize the black-and-white image by selecting Process | Binary | Skeletonize.</li>
		<li>Use the Analyze skeleton function in the Analyze | Skeleton menu to measure the microridge parameters and save the values. 
		NOTE: In SEM, individual images may differ in brightness and contrast. Thus, the steps in the analysis may need adjustments from image to image.</li>
	</ol>
	</li>
</ol>

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The authors have no conflicts of interest to disclose.

Corneal physical injuries are the most common cause of ophthalmology patient visits to the hospital. Therefore, it is important to establish relevant models for the study of different aspects of corneal pathophysiology. So far, the mouse is the most commonly used model for the study of corneal wound healing. However, adding eyedrops on murine wounded eyes to validate the impact of specific drugs on corneal wound healing can be difficult. In this respect, the zebrafish model is particularly useful for the pharmacological ...

As most of the epithelia are in contact with the external environment, they are prone to physical injury, making them well suited for the study of wound healing processes. Among the well-studied tissues, the cornea is an extremely useful model in the investigation of the cellular and molecular aspects of wound healing. As a transparent external surface, it provides physical protection to the eye and is the first element to focus the light onto the retina. While the structure and cell composition of the retina differ between species1, these elements of the cornea are generally similar in all camera-type eyes, regardless of species.
The cornea is composed of three main layers2. The first and outermost layer is the epithelium, which is constantly renewed to ensure its transparency. The second layer is the stroma, which contains scattered cells, called keratocytes, within a thick layer of strictly organized collagen fibers. The third and innermost layer is the endothelium, which allows nutrient and liquid diffusion from the anterior chamber to the outer layers. The epithelial and stromal cells interact via growth factors and cytokines3. This interaction is highlighted by the rapid apoptosis and subsequent proliferation of keratocytes after epithelial injury4,5. In case of a deeper wound, such as a puncture, keratocytes take an active part in the healing process6.
Being in contact with the external environment, corneal physical injuries are common. Many of them are caused by small foreign objects7, such as sand or dust. The reflex of eye rubbing can lead to extensive epithelial abrasions and corneal remodelling8. According to wound size and depth, these physical injuries are painful and take several days to heal9. The optimal wound healing characteristics of a model facilitate the understanding of the cellular and molecular aspects of wound closure. Furthermore, such models have also proved useful for testing new molecules with the potential to accelerate corneal healing, as previously demonstrated10,11.
The protocol described here aims to use zebrafish as a relevant model to study corneal physical injury. This model is highly convenient for pharmacological screening studies as it allows molecules to be added directly to the tank water and, therefore, to come into contact with a healing cornea. The details provided here will help scientists perform their studies on the zebrafish model. The in vivo injury is performed with a dulled ocular burr. The impact on epithelial cells adjoining or at a distance from it can be analyzed by specifically removing the central corneal epithelium. In recent years, numerous reports focused on such a method on rodent cornea12,13,14,15,16,17; however, to date, only a single report has applied this method to zebrafish18.
Because of its simplicity, the physical wound is useful in delineating the role of epithelial cells in wound closure. Another well-established model of corneal injury is the chemical burn, especially the alkali burn19,20,21. However, such an approach indirectly damages the entire eye surface, including the peripheral cornea and corneal stroma19. Indeed, alkali burns potentially induce corneal ulcers, perforations, epithelial opacification, and swift neovascularization22, and the uncontrollable outcome of alkali burns disqualifies that approach for general wound healing studies. Numerous other methods are also used to investigate corneal wound healing according to the particular focus of the study in question (e.g., complete epithelial debridement23, the combination of chemical and mechanical injury for partial-thickness wound24, excimer laser ablation for wounds extending to the stroma25). The use of an ocular burr restricts the focal point to the epithelial response to the wound and provides a highly reproducible wound.
As with each method of wound infliction, the use of an ocular burr has advantages and disadvantages. The main disadvantage is that the response being mostly epithelial, it does not perfectly reflect the abrasions seen in the clinical setting. However, this method has numerous advantages, including the ease with which it can be set up and performed, its precision, its reproducibility, and the fact that it is noninvasive, making it a method well tolerated by animals.

<table><tbody><tr><td>0.1M Na-PO&lt;sub&gt;4 &lt;/sub&gt;(sodium phosphate buffer), pH 7.4</td><td>in-house</td><td></td><td>Solution is prepared from 1M sodium phosphate buffer (1M Na&lt;sub&gt;2 &lt;/sub&gt;HPO&lt;sub&gt;4 &lt;/sub&gt; adjusted to pH 7.4 with 1M NaH&lt;sub&gt;2 &lt;/sub&gt;PO&lt;sub&gt;4 &lt;/sub&gt;).</td></tr><tr><td>0.2M Na-PO&lt;sub&gt;4 &lt;/sub&gt;(sodium phosphate buffer), pH 7.4</td><td>in-house</td><td></td><td>Solution is prepared from 1M sodium phosphate buffer (1M Na&lt;sub&gt;2 &lt;/sub&gt;HPO&lt;sub&gt;4 &lt;/sub&gt; adjusted to pH 7.4 with 1M NaH&lt;sub&gt;2 &lt;/sub&gt;PO&lt;sub&gt;4 &lt;/sub&gt;).</td></tr><tr><td>0.5mm burr tips</td><td>Alger Equipment Company</td><td>BU-5S</td><td></td></tr><tr><td>1M Tris, pH 8.8</td><td>in-house</td><td></td><td></td></tr><tr><td>adhesive tabs</td><td>Agar Scientific</td><td>G3347N</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>Cotton swaps</td><td>Heinz Herenz Hamburg</td><td>1030128</td><td></td></tr><tr><td>Dissecting plate</td><td>in-house</td><td></td><td></td></tr><tr><td>Dissecting tools</td><td>Fine Science Tools</td><td></td><td></td></tr><tr><td>double-distilled water</td><td>in-house</td><td></td><td></td></tr><tr><td>Eppedorf tubes, 2ml</td><td>any provider</td><td></td><td></td></tr><tr><td>Ethyl 3-aminobenzoate methanesulfonate salt</td><td>Sigma</td><td>A5040</td><td>Caution: causes irritation.</td></tr><tr><td>Glutaraldehyde, 50% aqueous solution, grade I</td><td>Sigma</td><td>G7651</td><td>Caution: toxic.</td></tr><tr><td>Lidocaine hydrochloride</td><td>Sigma</td><td>L5647</td><td>Caution: toxic.</td></tr><tr><td>mounts</td><td>Agar Scientific</td><td>G301P</td><td></td></tr><tr><td>Petri dish</td><td>Thermo Scientific</td><td>101VR20</td><td></td></tr><tr><td>pH indicator strips</td><td>Macherey-Nagel</td><td>92110</td><td></td></tr><tr><td>Plastic spoons</td><td>any provider</td><td></td><td></td></tr><tr><td>Plastic tubes, 15 ml</td><td>Greiner Bio-One</td><td>188271</td><td></td></tr><tr><td>Plastic tubes, 50 ml</td><td>Greiner Bio-One</td><td>227261</td><td></td></tr><tr><td>Scanning electron microscope</td><td>FEI</td><td>Quanta 250 FEG</td><td></td></tr><tr><td>Soft sponge</td><td>any provider</td><td></td><td></td></tr><tr><td>Sputter coater</td><td>Quorum Technologies</td><td>GQ150TS</td><td></td></tr><tr><td>Stereomicroscope</td><td>Leica</td><td></td><td></td></tr></tbody></table>

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 study describes a method using an ophthalmic burr in zebrafish corneal wound healing experiments. The method is modified from previous studies on mice, where the burr was shown to remove the epithelial cell layers efficiently13. The challenges in zebrafish corneal wounding include the relatively small size of the eye, and in the case of time-consuming experiments, the need to maintain a constant water flow through the gills (as described by Xu and colleagues28). The ma...

Watch this Scientific Journal Video about Zebrafish Corneal Wound Healing: From Abrasion to Wound Closure Imaging Analysis at JoVE.com

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

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. ...

zebrafish-corneal-wound-healing-from-abrasion-to-wound-closure

Research

JoVE Journal

Biology

2.8K Views.  University of Helsinki. 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.

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

Zebrafish Keratocyte Explants to Study Collective Cell Migration and Reepithelialization in Cutaneous Wound Healing

Due to their unique motile properties, fish keratocytes dissociated from explant cultures have long been used to study the mechanisms of single cell migration. However, when explants are established, these cells also move collectively, maintaining many of the features which make individual keratocytes an attractive model to study migration: rapid rates of motility, extensive actin-rich lamellae with a perpendicular actin cable, and relatively constant speed and direction of migration. In early explants, the rapid interconversion of cells migrating individually with those migrating collectively allows the study of the role of cell-cell adhesions in determining the mode of migration, and emphasizes the molecular links between the two modes of migration. Cells in later explants lose their ability to migrate rapidly and collectively as an epithelial to mesenchymal transition occurs and genes associated with wound healing and inflammation are differentially expressed. Thus, keratocyte explants can serve as an in vitro model for the reepithelialization that occurs during cutaneous wound healing and can represent a unique system to study mechanisms of collective cell migration in the context of a defined program of gene expression changes. A variety of mutant and transgenic zebrafish lines are available, which allows explants to be established from fish with different genetic backgrounds. This allows the role of different proteins within these processes to be uniquely addressed. The protocols outlined here describe an easy and effective method for establishing these explant cultures for use in a variety of assays related to collective cell migration.

Zebrafish keratocytes migrate in cell sheets from explants and provide an in vitro model for the study of the mechanisms of collective cell migration in the context of epithelial wound healing. These protocols detail an effective way to establish primary explant cultures for use in collective cell migration assays.

Due to their unique motile properties, fish keratocytes dissociated from explant cultures have long been used to study the mechanisms of single cell ...

Developmental Biology

Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two layers of skin that enclose undifferentiated mesenchyme, and because the larval tail fin regenerates rapidly within 2-3 days. Using this system, we demonstrate a method for capturing the repair dynamics of the amputated tail fin with time-lapse video brightfield stereomicroscopy. We demonstrate that fin amputation triggers a contraction of the amputation wound and extrusion of cells around the wound margin, leading to their subsequent clearance. Fin regeneration proceeds from proximal to distal direction after a short delay. In addition, developmental growth of the larva can be observed during all stages. The presented method provides an opportunity for observing and analyzing whole tissue-scale behaviors such as fin development and growth in a simple microscope setting, which is easily adaptable to any stereomicroscope with time-lapse capabilities.

We present a protocol for capturing the dynamics of zebrafish larval tail fin regeneration on a whole-tissue scale using brightfield-based stereomicroscopy. This technique enables capturing the regeneration dynamics with single cell resolution. This methodology can be adapted to any stereomicroscope equipped with a CCD camera and time-lapse software.

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two ...

Lens Transplantation in Zebrafish and its Application in the Analysis of Eye Mutants

The lens plays an important role in the development of the optic cup[1,2]. Using the zebrafish as a model organism, questions regarding lens development can be addressed. The zebrafish is useful for genetic studies due to several advantageous characteristics, including small size, high fecundity, short lifecycle, and ease of care. Lens development occurs rapidly in zebrafish. By 72 hpf, the zebrafish lens is functionally mature [3]. Abundant genetic and molecular resources are available to support research in zebrafish. In addition, the similarity of the zebrafish eye to those of other vertebrates provides basis for its use as an excellent animal model of human defects[4-7]. Several zebrafish mutants exhibit lens abnormalities, including high levels of cell death, which in some cases leads to a complete degeneration of lens tissues [8]. To determine whether lens abnormalities are due to intrinsic causes or to defective interactions with the surrounding tissues, transplantation of a mutant lens into a wild-type eye is performed. Using fire-polished metal needles, mutant or wild-type lenses are carefully dissected from the donor animal, and transferred into the host. To distinguish wild-type and mutant tissues, a transgenic line is used as the donor. This line expresses membrane-bound GFP in all tissues, including the lens. This transplantation technique is an essential tool in the studies of zebrafish lens mutants.

Lens development involves interactions with other tissues. Several zebrafish eye mutants are characterized by an abnormally small lens size. Here we demonstrate a lens transplantation experiment to determine whether this phenotype is due to intrinsic causes or defective interactions with tissues that surround the lens.

The lens plays an important role in the development of the optic cup[1,2]. Using the zebrafish as a model organism, questions regarding lens development ...

Examining Muscle Regeneration in Zebrafish Models of Muscle Disease

Skeletal muscle has a remarkable ability to regenerate following injury, which is driven by obligate tissue resident muscle stem cells. Following injury, the muscle stem cell is activated and undergoes cell proliferation to generate a pool of myoblasts, which subsequently differentiate to form new muscle fibers. In many muscle wasting conditions, including muscular dystrophy and ageing, this process is impaired resulting in the inability of muscle to regenerate. The process of muscle regeneration in zebrafish is highly conserved with mammalian systems providing an excellent system to study muscle stem cell function and regeneration, in muscle wasting conditions such as muscular dystrophy. Here, we present a method to examine muscle regeneration in zebrafish models of muscle disease. The first step involves the use of a genotyping platform that allows the determination of the genotype of the larvae prior to eliciting an injury. Having determined the genotype, the muscle is injured using a needle stab, following which polarizing light microscopy is used to determine the extent of muscle regeneration. We therefore provide a high throughput pipeline which allows the examination of muscle regeneration in zebrafish models of muscle disease.

Skeletal muscle regeneration is driven by tissue resident muscle stem cells, which are impaired in many muscle diseases such as muscular dystrophy, and this results in the inability of muscle to regenerate. Here, we describe a protocol that allows the examination of muscle regeneration in zebrafish models of muscle disease.

Skeletal muscle has a remarkable ability to regenerate following injury, which is driven by obligate tissue resident muscle stem cells. Following injury, ...

Long-term Live Imaging Device for Improved Experimental Manipulation of Zebrafish Larvae

The zebrafish larva is an important model organism for both developmental biology and wound healing. Further, the zebrafish larva is a valuable system for live high-resolution microscopic imaging of dynamic biological phenomena in space and time with cellular resolution. However, the traditional method of agarose encapsulation for live imaging can impede larval development and tissue regrowth. Therefore, this manuscript describes the zWEDGI (zebrafish Wounding and Entrapment Device for Growth and Imaging), which was designed and fabricated as a functionally compartmentalized device to orient larvae for high-resolution microscopy while permitting caudal fin transection within the device and subsequent unrestrained tail development and re-growth. This device allows for wounding and long-term imaging while maintaining viability. Given that the zWEDGI mold is 3D printed, the customizability of its geometries make it easily modified for diverse zebrafish imaging applications. Furthermore, the zWEDGI offers numerous benefits, such as access to the larva during experimentation for wounding or for the application of reagents, paralleled orientation of multiple larvae for streamlined imaging, and reusability of the device.

This manuscript describes the zWEDGI (zebrafish Wounding and Entrapment Device for Growth and Imaging), which is a compartmentalized device designed to orient and restrain zebrafish larvae. The design permits tail transection and long-term collection of high-resolution fluorescent microscopy images of wound healing and regeneration.

The zebrafish larva is an important model organism for both developmental biology and wound healing. Further, the zebrafish larva is a valuable system for ...

Bioengineering

Stab Wound Injury Model of the Adult Optic Tectum Using Zebrafish and Medaka for the Comparative Analysis of Regenerative Capacity

While zebrafish have a superior capacity to regenerate their central nervous system (CNS), medaka has a lower CNS regenerative capacity. A brain injury model was developed in the adult optic tectum of zebrafish and medaka and comparative histological and molecular analyses were performed to elucidate the molecular mechanisms regulating the high regenerative capacity of this tissue across these fish species. Here a stab wound injury model is presented for the adult optic tectum using a needle and histological analyses for proliferation and differentiation of the neural stem cells (NSCs). A needle was manually inserted into the central region of the optic tectum, and then the fish were intracardially perfused, and their brains were dissected. These tissues were then cryosectioned and evaluated using immunostaining against the appropriate NSC proliferation and differentiation markers. This tectum injury model provides robust and reproducible results in both zebrafish and medaka, allowing for comparing NSC responses after injury. This method is available for small teleosts, including zebrafish, medaka, and African killifish, and enables us to compare their regenerative capacity and investigate unique molecular mechanisms.

A mechanical brain injury model in the adult zebrafish is described to investigate the molecular mechanisms regulating their high regenerative capacity. The method explains to create a stab wound injury in the optic tectum of multiple species of small fish to evaluate the regenerative responses using fluorescent immunostaining.

While zebrafish have a superior capacity to regenerate their central nervous system (CNS), medaka has a lower CNS regenerative capacity. A brain injury ...

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 ...

Live Imaging of Chemokine Receptors in Zebrafish Neutrophils During Wound Responses

Leukocyte guidance by chemical gradients is essential for immune responses. Neutrophils are the first cells to be recruited to sites of tissue damage where they execute crucial antimicrobial functions. Their trafficking to these loci is orchestrated by several inflammatory chemoattractants, including chemokines. At the molecular level, chemoattractant signaling is regulated by the intracellular trafficking of the corresponding receptors. However, it remains unclear how subcellular changes in chemokine receptors affect leukocyte migration dynamics at the cell and tissue level. Here we describe a methodology for live imaging and quantitative analysis of chemokine receptor dynamics in neutrophils during inflammatory responses to tissue damage. These tools have revealed that differential chemokine receptor trafficking in zebrafish neutrophils coordinates neutrophil clustering and dispersal at sites of tissue damage. This has implications for our understanding of how inflammatory responses are self-resolved. The described tools could be used to understand neutrophil migration patterns in a variety of physiological and pathological settings and the methodology could be expanded to other signaling receptors.

Here we describe protocols to perform live imaging and quantitative analysis of chemoattractant receptor dynamics in zebrafish neutrophils

Leukocyte guidance by chemical gradients is essential for immune responses. Neutrophils are the first cells to be recruited to sites of tissue damage ...

Immunology and Infection

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 ...

Medicine

Assessment of Zebrafish Lens Nucleus Localization and Sutural Integrity

The zebrafish is uniquely suited to genetic manipulation and in vivo imaging, making it an increasingly popular model for reverse genetic studies and for generation of transgenics for in vivo imaging. These unique capabilities make the zebrafish an ideal platform to study ocular lens development and physiology. Our recent findings that an Aquaporin-0, Aqp0a, is required for stability of the anterior lens suture, as well as for the shift of the lens nucleus to the lens center with age led us to develop tools especially suited to analyzing the properties of zebrafish lenses. Here we outline detailed methods for lens dissection that can be applied to both larval and adult lenses, to prepare them for histological analysis, immunohistochemistry and imaging. We focus on analysis of lens suture integrity and cortical cell morphology and compare data generated from dissected lenses with data obtained from in vivo imaging of lens morphology made possible by a novel transgenic zebrafish line with a genetically encoded fluorescent marker. Analysis of dissected lenses perpendicular to their optical axis allows quantification of the relative position of the lens nucleus along the anterior-posterior axis. Movement of the lens nucleus from an initial anterior position to the center is required for normal lens optics in adult zebrafish. Thus, a quantitative measure of lens nuclear position directly correlates with its optical properties.

These protocols were developed to analyze cortical lens morphology, structural integrity of the zebrafish lens sutures in fixed and live lenses and to measure the position of the zebrafish lens nucleus along the anterior-posterior axis.

The zebrafish is uniquely suited to genetic manipulation and in vivo imaging, making it an increasingly popular model for reverse genetic studies and for ...