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

This video demonstrates a method to capture tissue repair dynamics on a stereo microscope using time-lapse imaging. This is accomplished by first raising the zebrafish embryos to larval stages when the tail fin has sufficiently developed to a size in which amputations can be readily performed. This is typically achieved post hatching.

The second step is to amputate the tail fin using a syringe needle. The third step is to mount the larvae into an imaging chamber and capture a time-lapse movie on a stereo microscope. The final step is to quantify tail fin regeneration utilizing imaging analysis software such as Bit Plains S or the open source software image J.Hi.My lab studies wound repair and tissue regeneration using zebrafish as a model.

Today we will demonstrate a method that captures the dynamics of these processes with real time using brightfield stereo microscopy. Today we'll demonstrate a method which captures the early processes of tissue regeneration using the zebrafish larval tail fin. The presented imaging method provides an opportunity for observing and analyzing whole tissue scale behaviors in a simple microscope setting, which is easily adaptable to any stereo microscope equipped with time-lapse capabilities.

So let's get started. Raising of zebrafish To larval stages. Wound repair in the tail fin can be best observed after hatching stages as the tail fin has sufficiently developed by this time, and amputations can be easily performed without damaging the spinal cord or other tissues.

In close proximity, collect the eggs and incubate them in a 28 degree Celsius incubator overnight. Next morning, remove the dead embryos and rinse with embryo medium containing 0.03%instant ocean aquarium salt solution. If required, add embryo medium with 0.003%pheno thyroid to the dish to prevent pigmentation of the larvae.

For this demonstration, we let the embryos develop until two days post fertilization in the incubator when the larvae have hatched from their kons preparation of the imaging chamber. In this step, we provide two methods for constructing the imaging chamber. Method one consists of an imaging chamber made from drinking water grade PVC or Teflon tubing, which can be obtained in any hardware store.

Utilize tubing with a 25 millimeter outer and a 20 millimeter inner diameter as this has the appropriate size for cover slip attachment. Cut the tubing to make 10 millimeter thick rings as even as possible. Use sandpaper to smoothen the edges and rinse in water and ethanol.

Let the chamber air dry. Method two consists of either utilizing a pre-manufactured matte tech glass bottom, glass top imaging chamber, or making an imaging chamber from a small Petri dish. To make your own imaging chamber, utilize a 35 or 50 millimeter Petri dish and drill a 10 millimeter hole into the lid.

Next applies silicone grease to the outside of the cover slip and attach a 25 by 25 millimeter square or round cover slip to the outside with a clean pipette tip to ensure that the mounting agros stays firmly attached. During the imaging procedure, attach a fine plastic mesh to the inside of the cover slip. This can be accomplished by cutting the mesh into the size of the inner ring diameter and then cutting a small rectangle about three times the size of the larvae into the middle of the mesh.

Mounting and imaging of the pre-injury larvae. This step is suitable for later quantification of fin regeneration. First, prepare a 0.5%low melt agro solution in embryo medium for immobilization of the specimen transferring anesthetized larva into the aro solution that has been maintained at 42 degrees Celsius.

In a heating block, transfer the larvae with a drop of agros into a small Petri dish and position the larvae on its side. Using a capped micro loader tip, allow the aros to solidify and add trica solution. Proceed to the stereo microscope that will be used later.

For time-lapse imaging, we use a zes discovery V 12 stereo microscope and Axio vision software for demonstration. First, adjust the microscope settings by selecting the appropriate objective and magnification for imaging. Then select the camera detection mode on the microscope in the alvis software.

Select the live mode to view the larva on the screen. Open the properties window to automatically detect the brightness manually adjust the contrast at the microscope transillumination base. Move the larvae out of the field of view and select the shading correction feature.

To minimize background noise, position the larvae back in the field of view and take a snapshot. Save the image to prepare for wounding. Remove the larvae from the aros by first scraping the aeros off the head.

This allows the larvae to be able to slip out of the aeros by gently pulling the head away from the remaining aeros amputation assay. Prepare a 1.5%aeros plate using instant ocean salt solution. Under a stereo microscope, place the larvae sideward onto the solidified aros and amputate the distal part of the tail fin with a 23 gauge syringe needle.

Mounting the larvae for time-lapse imaging. We will present two time-lapse imaging methods. In the first method, we will show how to image the larvae in the chamber ring following mounting of the larvae.

Carefully scrape off the solidified agros surrounding the distal tail fin with a capped micro loader pipette tip. This allows for proper wound healing or tissue regeneration Without restraining the tissue, try not to injure the fin repeatedly. During this procedure, remove unwanted debris that interferes with the imaging procedure from the chamber by replacing the old solution.

With fresh Trica solution, apply silicon grease to the top of the chamber and fill the chamber ring with fresh trica solution. Apply a glass slide to the top to seal the chamber method two. In this method, we will show you how to image the larvae in a glass top Petri dish.

Mount the larvae onto the lid, cover slip and fill the lid. With trica solution. Scrape off the agros from the tail fin.

Next, apply silicon grease to the top rim of the bottom chamber and fill the chamber with trica solution. Carefully decant the trica solution from the lid and turn the lid over to immerse larva into the trica solution of the bottom chamber, which can be done at a slight angle. To avoid air pockets, the chamber will be sealed with the silicon grease time-lapse imaging to ensure that a proper temperature of 28 degrees Celsius is maintained.

Assemble a heated incubation chamber made of cardboard bubble wrap for insulation and a wired dome as used for chicken egg incubation as a heat source. Place the incubation chamber around the microscope and adjust the temperature to 28 degrees Celsius. For about 10 to 20 minutes or until the temperature has stabilized, open the front of the heated incubation chamber and place the imaging chamber onto the microscope.

Stand with the cover slip facing upward toward the objective. Position the larval fin in a way that two thirds of the field of view remains unoccupied to ensure the capture of the growth and regeneration of the fin over the course of the imaging procedure. Without having to reposition the larvae in the 60 multi-dimensional acquisition window of the software, select the Z stack and time-lapse option in the Zack tab and slice mode.

Select the slice thickness and then select the start stop mode. Define the upper and Lower positions of the stack in the time-lapse table. Select the interval and the duration of the movie, and then press the start button to begin the recording.

Within the first hour, check the and Zack boundaries of the larvae. If necessary, reposition the larvae over time as it may have shifted due to the growth or differences in the agros composition. Save the file at the end of the time-lapse recording and proceed with the post-processing and quantification step data analysis.

In this section, we discuss several ways to analyze the data generated method one imas. First, we will discuss how to determine fin growth with imas software. Open the time-lapse movie in the Amaris software and save files in the MRS file format.

To enhance Software performance, select the orthogonal view to display The movie as a projection. To measure the fin length, select the add new measurement points option in the upper left toolbar. Under configure list of visible statistics value, select the statistics values to be displayed in the line mode.

Under the settings tab, select pairs in labels properties, select name and distance to display the distance between points A and B next to the measurement line, switch to the edit mode and hold down the shift button to select the first point at the distal fin. Then use the same configuration to select the second point at the distal end of the no accord. The distance will be displayed in the image and can be exported under the statistics tab.

By selecting the disc button export all at the lower right. Alternatively to the measurement points option, the slice viewer can be utilized to measure distances in the slice view mode. Scroll to a desired position and click on the first and second positions with the left mouse button, the distance will be displayed.

This option, however, does not allow for data export. Now we will briefly discuss how to determine the fin area using the open source software. Image J.Open the TimeLapse movie in image J using a plugin that recognizes the Zeiss Axio vision file format.

Alternatively load in an uncompressed TIFF sequence. This will ensure that the file information is saved under analyze select set scale to define the distance and pixels and known distance, if not preserved in the file. Also, set the unit length to micrometers.

In the toolbar. Select the free hand selection tool and outline the wound by holding the left mouse button down. While drawing along the wound edges.

Click the measure option under analyze to display the area of the wound representative results. Amputation of the fin leads to a partial regeneration over the course of 36 hours following the amputation. The area measurements of the fin show that fin amputation triggers a decrease in fin tissue until about 14 hours post amputation, presumably due to cell death, which is followed by thin regeneration length.

Comparison of the amputated and regenerated fin shows a 2.48 fold increase in fin length within 36 hours. Amputation initially triggers a purse string effect characterized by contractions via actin myosin cables that are present in the fin fold cells extrude from the amputation wound and are subsequently cleared. The body length increases initially, but the fin does not regenerate.

During the early phases. At about 14 hours post amputation, the fin starts to grow out from proximal to distal positions. In summary, our results demonstrate that amputation triggers wound contraction, which may be essential to promote cell extrusion.

Once the process is completed, the fin begins to regenerate. The developmental fin growth appears to occur. Independent of this process, We have presented a method which allows for the observation of the dynamic processes of tissue repair using living zebrafish larvae.

This procedure requires certain aspects which we have tested in order to optimize the outcome. A clear advantage of the presented imaging method is that it is readily adaptable to any stereo microscope equipped with a CCD camera and TimeLapse software. It also offers a low-cost alternative to the more expensive confocal imaging system.

While this method does not used fluorescence or cell detection, it can be expanded for such applications by utilizing an automated system for shatter control and post imaging deconvolution software to further observe the processes of wound repair and regeneration with subcellular resolution. This method can provide students with a better understanding of the basic biological processes, underlying tissue repair and regeneration. The optical clarity and the ease with which embryonic and larval zebrafish can be used and the adaptability of the system to any stereo microscope makes it suitable for teaching basic vertebrae biology in a classroom setting.