Forebrain Electrophysiological Recording in Larval Zebrafish

The overall goal of this procedure is to obtain extracellular field recordings from the forebrain of an intact larval zebra fish. This is accomplished by first immobilizing the larval zebra fish in an aros block. Next, a recording micro electrode is placed in the forebrain under direct visualization.

The final step is to record forebrain electrical activity. Ultimately, results can be used to monitor abnormal electrographic seizure activity in intact zebrafish larvae. I first had the idea for this method when I was seeking to demonstrate that seizures could be elicited in larval zebrafish Zebrafish husbandry follow standard procedures described previously.Briefly.

Adult zebrafish are set up in breeding tanks with dividers in place. When the lights in the room come on the following morning, the dividers are removed and the fish are allowed 20 to 60 minutes of undisturbed mating time. Eggs are then collected from the tanks, transferred to a Petri dish with egg water and cleared of debris with a transfer pipette now place the eggs in an incubator, and in about two days, the eggs will hatch.

Once hatched, remove the corion and any other debris with a transfer pipette then return the larvae to the incubator until they reach the desired developmental time point. In preparation, pull 1.2 millimeter outer diameter bo silicate glass capillaries into needles using a micro pipette puller. Store the needles in a 150 millimeter Petri dish over putty ramps.

Now pack, load the micro electrode with the two molar sodium chloride solution. Using a one liter syringe with a four millimeter syringe filter and attached filament, tap the bolus near the needle tip until there are few or no bubbles remaining. Prepare a fresh solution of 1.2%LMP agar in egg water and keep it melted in a 37 degree Celsius bath.

Next, prepare a cover slip with the appropriate recording chamber and place it in a freezer for five to 10 minutes. This example uses a low profile open diamond bath imaging chamber in a clean Petri dish plate submerge one larval seabra, fish in a droplet of egg water using a transfer pipette. Next, mix in a droplet of solution containing an anesthetic and paralyzing agent for five to 10 minutes.

Monitor the zebra fish for loss of movement. Then remove the cover slip and recording chamber from the freezer. Place the chamber on the stereo microscope stage to the chamber.

Using a transfer pipette, add several milliliters of agros to the fish in water and mix them together Using a micro pipette tip or fine blunt needle. Position the larvae so that the dorsal aspect of the fish is exposed to the agros still surface. After the agros hardens, use a flat spatula to transfer the agro block with the larvae to a recording chamber set up on an electrophysiology rig.

At the rig, add two to five milliliters of zebrafish recording media to the recording chamber. If drug changes are necessary, perfuse the chamber with a recording solution at a rate of approximately one milliliter per minute, no oxygenation is necessary. Next, insert a micro electrode into the amplifier head stage mounted on a three-dimensional micro manipulator.

Once loaded, check that the microm manipulator has a good range of movement and adjustment. Then bring the micro electrode tip into the plane of view of the microscope high off the stage, and using the course adjustment, lower it to a point just above and slightly in front of the head of the zebrafish. With the micro electrode in place, put the amplifier in current clamp mode and zero the electrode in the field of view.

Use the course adjustment to lower the electrode tip so that it just touches the surface of the zebrafish slightly in front of the forebrain. Then with the fine adjustment step, advance the electrode tip until it punctures the skin of the zebra fish. Once punctured, allow the micro pipette to settle slowly advance the electrode several microns into the forebrain.

Now use aScope software to record electrical activity. In current clamp mode, acquire data at a sampling rate of 10 kilohertz. The fish remains viable for 24 hours of recording to apply drugs, add them in two to five milliliter volumes and wait 30 to 45 minutes for the drugs to diffuse through the agar and into the zebra fish.

Four brain. A static bath configuration is acceptable, making continuous perfusion unnecessary. Using these methods, extracellular recordings can also be obtained from the optic tum.

Electrographic seizure-like discharges were recorded in the forebrain of eight day post fertilization, zebrafish larvae, large amplitude multis spike bursts were revoked by a bath application of 40 millimolar pilocarpine, A convulsant drug. Similar bursts were produced with a convulsant drug. Ritoxin applied at one millimolar.

A spontaneous burst discharge was observed from a genetically modified zebra fish at three days post fertilization. In this animal, morpho oligonucleotide injections at the one to two cell stage were used to knock down expression of a gene for tuberous sclerosis complex, a pediatric form of epilepsy associated with seizures and autism. If performed correctly, this procedure can be completed in 30 minutes or less.