Intraperitoneal Injection into Adult Zebrafish

The overall goal of this procedure is to inject a solution into the peritoneal cavity of the adult zebra fish. In this example, we will inject glucose in our lab. We use this procedure to study blood glucose stasis.

This is accomplished by first weighing the fish to determine how much solution to inject per gram of body weight. The second step of the procedure is to anesthetize the fish with cold water. The third step of the procedure is to transfer the fish to the microscope stage and inject.

The final step of the procedure is to return the fish to its tank for recovery. Ultimately, results can be obtained that show changes in glucose homeostasis through measurement of blood glucose levels. Hi, I'm Mary Kinkle from the Laboratory of Victoria Prince in the Department of Organismal Biology and Anatomy at the University of Chicago.

I'm Stephanie Ames also from the Prince Lab. Today we'll show you a procedure for intraperitoneal injection into adult Zebrafish. We use this procedure in our laboratory to study the regulation of blood glucose homeostasis.

So let's get started. To prepare fish for injection, start by transferring the fish to a clean tank and withhold food fast. The fish for 24 hours to empty the intestinal bulb contents and 72 hours to reduce blood glucose to baseline levels.

The density used in this protocol is 10 to 12 mixed sex population. Fish per nine liter tank with three layers of marbles, the marble sequester eggs so that they're not eaten. Remove eggs and waste daily by siphoning the tank.

Bottom next, prepare Cortland salt solution using 124.1 millimolar sodium chloride 5.1 millimolar potassium chloride 2.9 millimolar sodium pyrophosphate, 1.9 millimolar magnesium sulfate hept hydrate, 1.4 millimolar calcium chloride dihydrate 11.9 millimolar sodium bicarbonate, four grams of polyvinyl roone, and 1000 USP units of heparin to a final volume of 100 milliliters with deionized water filter, sterilize, and store at four degrees Celsius. Next, dissolve five grams of glucose in 10 milliliters of cottin salt solution. To make a 0.5 milligrams per microliter glucose solution, filter, sterilize, and store at four degrees Celsius.

Then prepare the surgical table by cutting a soft sponge to approximately 20 millimeter height on the flat face. Make a 10 to 15 millimeter deep cut to hold the fish for injection. Set the sponge into a 60 millimeter Petri dish and set the Petri dish with a sponge into a P 200 pipette tip box lid.

Next, prepare the microscope by covering the microscope base with plastic wrap and paper towels for protection. In case of spills, set the surgical table on top of the paper towel. Pre adjust focus by viewing the surgical table and focusing on the sponge.

Put your finger on top of the sponge and focus on it to minimize further focal adjustment when the fish is on the surgical table. After weigh the fish by filling a 500 milliliter beaker about one third full with fish facility water and tear. On a scale net transfer fish with as little excess water as possible to the beaker and record the weight transfer each weighed fish to a clear labeled tank.

Calculate injection volume based on weight. In this example, we will inject a solution of 0.5 milligrams per gram body weight of glucose dissolved in Cortland salt solution. Since our solution is 0.5 milligrams per microliter, the calculation is one times the fish weighting grams equals the volume to inject in microliters, prepare the syringe and related injection equipment.

This protocol uses a 30 5G beveled steel needle, ultra micro pump and micro four processor and a 10 microliter nano fill micro syringe for glucose injection, fill the syringe with 0.5 milligrams per microliter glucose dissolved in Cortland salt solution. It is important to remove all air bubbles from the syringe mount the syringe on the ultra micro pump and program the injection volume for the first fish by entering the volume in nanoliters, using the numbered keypad on the micro four processor. Lastly, prepare the ice anesthetic by crushing enough ice cubes made from fish facility water.

To fill an ice bucket, put the surgical table into a larger container such as a 2.4 liter rubber made food storage container. Pour some warm facility water into the outer container and the surgical table. Keep a reserve tank of warm facility water nearby.

Attach a thermometer to the outer container. Place the anesthetic outer container plus surgical table adjacent to the microscope. Have the bucket of ice chips nearby.

Begin adding ice chips until the water temperature is 17 degrees Celsius. Do not go below 17 degrees Celsius for this step. Use a net to transfer the fish to the outer container.

At 17 degrees or slightly lower, the fish typically will spread its pectoral fins horizontally, gasp, and have rapid auricular movements. Slowly add ice chips to the container to bring the temperature down to 12 degrees Celsius over the course of several minutes. As the temperature drops, the fish will swim more slowly and finally stop swimming.

Eventually gasping will stop and auricular movements will slow. The fish is ready for injection when it does not react to being handled. As the required temperature is reached, press on the sponge to saturate it.

Keep your fingers in the cold water sufficiently so that they will not warm up the fish and bring it out of anesthesia. During handling with cold fingers, gently transfer the fish to the trough of the sponge. Position the fish with the abdomen up and the gills in the trough.

Quickly transfer the surgical table to the microscope stage. Working quickly, carefully position the needle so that the tip points cran and is closer to the pelvic girdle than to the anus. Insert the needle into the midline between the pelvic fins.

You should be able to feel when the needle is deep to the body wall. As you insert the needle, you can feel the body wall give. When the needle enters the abdominal cavity, inject the appropriate volume using the micro for processor.

Use the foot pedal to inject one microliter per gram of fish body weight after injection, remove the needle and immediately transfer the fish back to its warm water tank for recovery. By releasing the fish from the sponge over the tank water, check the needle under the microscope. Occasionally a scale will be attached and should be removed before the next injection.

After a suitable period of time, proceed with your downstream assay. We have found that a 72 hour fast is required to decrease blood glucose to a baseline level prior to injection. In this figure, the non injected control data shown in green displays blood glucose values not significantly different from vehicle injected controls shown here in red.

However, the blue line displays blood glucose values for glucose injected fish and displays the rise and fall of circulating glucose. Over time, both glucose injected and vehicle injected animals were under surgical anesthesia during the injection procedure, whereas non injected controls were not under surgical anesthesia. We've just shown you how to perform intraperitoneal injection into adult zebrafish.

In our lab, we study changes in blood glucose concentration at various time points following IP injection of a glucose solution. However, this technique we just showed you can be used to inject your solution of interest. When doing this procedure, remember to anesthetize the fish slowly and handle it gently.

So that's it. Thanks for Watching and good luck with your experiments.