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

Corneal transparency is instrumental to clear sight. In pathological context, this transparency can be challenge. Studying pathogenesis mechanism requires simple models having a high similarity to human organ.

In these aspects, zebrafish is an excellent model. And this is the first detailed description of corneal abrasion in zebrafish. This technique is a simple and fast way to induce surface entry to the eye.

The wound closure can be followed easily after abrasion. Also, chemicals, or specific factors, can be added to the tank water to study their impact on wound closure. As the wound is created manually, the method requires some practice to gain consistent results.

Before the experiment, prepare a 0.02%working solution of tricaine. Thaw 2 mL of 0.4%stock solution. Add to 40 mL of system water.

And place the solution in a small container. Have the ophthalmic burr ready by checking if the burr tip is clean. And if needed, remove cell debris with a moist cotton swab.

Make an incision to the side of a soft sponge. And moisten the sponge with system water. Place the sponge on the stage of a dissecting microscope.

Ensure enough working space for using the burr. And enough illumination from the side and above to see the eye surface correctly. Gently place the anesthetized fish with a spoon into the incision on the sponge with the head protruding from the sponge surface.

Carefully approach the eye surface with the burr tip. And start moving the tip on the eye surface with a circular motion. When the abrasion is done, carefully place the fish in the fresh system water containing analgesic for recovery.

Clean the burr immediately after use with a moist cotton swab. Pick up the fish at the desired time point and place it in 0.02%tricaine solution. Keep the animal in the solution until the opercular movement has ceased entirely and the fish does not react to touching.

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.

Put the tissue into a sample tube containing 0.1 M sodium phosphate. And rinse it with a clean buffer so that no blood remains in the solution. Fix the tissue in 2.5%glutaraldehyde in 0.1 M sodium phosphate for 24 hours at 4 C.Remove the fixing solution and rinse the sample several times with 0.1 M sodium phosphate.

Dissect the sample by placing it onto a drop of 0.1 M sodium phosphate on a dissecting plate. Cut the head sample into two with fine dissecting scissors. Alternatively, collect the eyes by carefully placing the tips of fine tweezers into the eye sockets from the side of the eye.

Then, pull out the eye from the socket and remove extra tissue. Transfer the dissected sample into a tube containing 0.1 M sodium phosphate. Ensure there is no extra tissue in the sample tube, as it may adhere to the top of the eye during sample processing.

Place the mount with the tab on a holder. And the holder to the base of a dissecting microscope. Gently place the tissue sample on the mount with fine tweezers, cornea facing up.

Coat the specimen with platinum using the appropriate device. And store the samples at room temperature until imaging. Acquire images of the desired magnification.

And use 2000 to 2500x images for analysis. Adjust the brightness and contrast to clearly see all the cell borders and microridges. And avoid overexposed areas in the image.

Open the TIFF image with Fiji ImageJ 1.53. And use a scale bar to set the scale. Create a line equal to the scale bar with the Line tool.

Select Analyze. Then, Set scale. And type in the known distance.

Then, open the ROI manager using the Analyze, Tools menu. For cell size and roundness select Analyze, Set Measurements, Shape descriptors. And 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. To proceed with microridge analysis, ensure the image is in 8-bit format by clicking on the Image, Type menu.

Then, select the cell with the Polygon tool. And clear the background by clicking on Edit, then Clear Outside. 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. Convolve the image by clicking on Process, Filters, Convolve. And turn it into binary by clicking Process, Binary, Make Binary.

Then, skeletonize the black and white image by selecting Process, Binary, Skeletonize. Use the Analyzed Skeleton function to measure the microridge parameters and save the values. The wound area is closed at three hours post wound.

But the site where the wound borders are joined remains visible. The results showed that on abraded corneal epithelium microridges can be observed in some elongated cells next to the wound site. In some peripheral regions, the microridges are lost from the center of the cell.

A comparison between the two of peripheral regions, showed elongated cells with shorter average microridges, indicating cell rearrangements as a reaction to wounding. These results suggest that cell shape changes correlate with microridge modification and emphasize the heterogeneity within the corneal epithelium in wound response. Remember that the operation requires training to perform a homogeneous and reproducible wound.

In addition to electron microscopy, the tissue can be used for histology, immunological stainings and in situ hybridization. These will provide further insight in the cell and molecular changes during wound healing. Using zebrafish broadens our knowledge on corneal biology.

Moreover, this model is excellent for pharmacological studies, and can be used to understand more about eye evolution.