Evaluation of Zebrafish Kidney Function Using a Fluorescent Clearance Assay

The overall goal of this procedure is to generate a quantitative kidney function assay in zebra fish. This is accomplished by first anesthetizing zebra fish embryos at 72 HPF. Next, the embryos are transferred to an injection mold and orientated so that their hearts are to the left of the field of view.

Then the pericardial cavity is injected with a rumine dextra fluorescent dye and fluorescent images are required of the cardiac region using an epi fluorescence dissecting microscope. Ultimately, kidney clearance of a fluorescent dye is assessed using epi fluorescence microscopy. The main advantage of this technique is that it produces a quantitative readout of zebra fish kidney function without requiring blood or urinary tests, which aren’t possible in zebra fish.

This method provides a powerful tool for evaluating kidney function in zebrafish disease models. After pulling needles and preparing a mold according to the text protocol, cast the mold by first pouring a 2%aro solution made in fish water into a 90 millimeter petri dish. Place the mold on the Petri dish, permitting the stacked slide edges to submerge at an angle under the agro surface.

Allow it to set for 30 minutes. Remove the slide cast and use fresh fish water containing trica anesthetic at a one to 25 ratio. To cover the slide imprinted aros to perform zebra fish injections of fluorescent dye.

Use a standard microinjection setup consisting of an air compressor connected to a pressure regulator system that feeds into a straight pipette holder for use with 1.0 outer diameter capillaries. The pipette holder should be housed within an MN 33 compact. Three axis control micro manipulator secured to a steel base plate by a magnetic stand user dissecting microscope to visualize, manipulate, and inject embryos while maintaining zebra fish using standard conditions as outlined in the text protocol.

Use wild type or transgenic zebra fish lines as appropriate to the experiment. Maintaining a stocking density of 15 males to 15 females per eight liter tank to ensure maximum eggs are collected without disturbing the fish. Submerge breeding tanks into wild type stock tanks the evening before collection.

On the day of collection, allow 30 to 40 minutes for the fish to spawn. Then use a mesh strainer and fresh aquarium water to collect and rinse the eggs. Deposit the cleaned eggs in a 90 millimeter Petri dish containing embryo, medium, and incubate at 28.5 degrees Celsius to inhibit myogenesis.

At eight hours. Post fertilization or HPF. Transfer the viable embryos in minimal liquid to fresh Petri dishes containing one to 100 PTU to embryo medium and further incubate a 28.5 degrees Celsius.

Use fine tipped forceps to break the end of the needle. Then move the RD to the tip of the needle by maximizing pulse duration to the minute setting. When the solution reaches the tip, switch back to milliseconds.

Place a micrometer on the microscope stage and with the pressure regulator and refinements to the needle tip, adjust the size of the expel droplet to 100 microns in diameter or a volume of 0.5 nanoliters prior to injection. Set up 2 35 millimeter Petri dishes. Each containing five milliliters of embryo medium label one trica and the other recovery.

Add 200 microliters of stock trica to the trica labeled dish. Once ready to inject, select an individual embryo at 72 HPF. Transfer minimal liquid to the trica dish and monitor the activity of the embryo.

Once the embryo is anesthetized, transfer it to the injection mold and orientate it in the trough so that the left side is facing up. Positioning the heart to the left side of the field of view. To inject the RD into the heart, use the needle to pierce the pericardium and inject one nanoliter of RD into the pericardial cavity.

After withdrawing the needle, transfer the injected embryo in minimal liquid to the recovery dish and monitor for one minute. Then transfer the individual embryo to a 24 well plate containing one milliliter of fresh embryo medium containing PTU and label appropriately. After injecting at least 10 embryos per experimental group incubate at 28.5 degrees Celsius for three hours.

At three hours post injection or HPI anesthetized embryos as previously described. And transfer to a 35 millimeter Petri dish containing 3%methyl cellulose. Gently push embryos into the methyl cellulose and orientate so that the lateral side can be imaged.

Using an emission filter of 570 nanometers for UV light, acquire an image of the embryo. Allow the embryo to recover before replacing it in the designated well acquire images for all injected embryos, then continue to incubate at 28.5 degrees Celsius. After acquiring a second round of images at 24 HP, I use image J software to quantify the fluorescence intensity of each injected embryo by opening an image and specifying a region of interest or ROI.

By choosing edit selection, specify and setting the value at 100 square pixels. Position the heart in the center of the ROI and choose analyze, set measurements and set the measurements to include mean gray scale and area. Then take the measurement, quantify the fluorescence for each set of images at three HPI and 24 HPI per embryo and transfer the average gray scale values to a spreadsheet for further processing.

Use appropriate software to perform statistical analysis and compare groups for statistical significance using the student’s T-test as shown here. Injection of RD into bbbs nine Morpho injected fish resulted in 40%of the embryos developing prone fric cysts by five days post fertilization. In addition, the morphin fish exhibited a reduction in the ability to clear the fluorescent dye after 24 HPI compared to controls.

Once must a group of 10 fish can be injected within 15 minutes. After watching this video, you should have a good understanding of how to take advantage of the optical transparency of zebrafish to quantitatively monitor the temporal clearance of a micro injected fluorescent dye as an effective readout of zebrafish kidney function.