Electroretinogram Analysis of the Visual Response in Zebrafish Larvae

The overall goal of this procedure is to illustrate the steps necessary to perform electrographic analysis on larval zebra fish. This is accomplished by first generating the appropriately shaped and sized glass micro pipettes. The second step is to carefully set up equipment critical to the experiment, such as reference and recording electrodes.

Next, the larval zebrafish is anesthetized and properly oriented relative to the stimulation and recording equipment. The final step is the proper placement of the glass micro electrode onto the cornea of the larval zebra fish. Ultimately larval zebra fish.

ERG analysis is useful for investigating cone visual function in the all cone vertebrate retina in vivo. This method is very useful for answering key questions in the field of vision science, such as the identification of signaling pathways that contribute to visual sensitivity and light adaptation in cones. Visual demonstration of this method is critical as the proper positioning of the recording micro electrode onto the specimen is difficult to learn due to the small size of the larval zebrafish eye.

To begin this procedure, pull several micro pipettes using fire polished boro silicate glass capillaries with filament. Check each micro pipette under a microscope with an appropriate grad ruler to ensure that the tips diameter is 10 to 15 micrometers and even in appearance. Create a working solution on the day of the experiment by diluting 10 x ringer solution to one x.

With deionized distilled water filter, sterilize the solution to ensure that particulates will not block the micro pipette tip. Then oxygenated with 95%oxygen, 5%carbon dioxide for 10 minutes. Cap the solution tightly afterwards to ensure that the solution remains oxygenated.

Now place a movable plastic platform with a viscoelastic urethane polymer shock absorbing bottom on the anti vibration table under the light source. Next, position the camera with a magnetized stand and aim it at the movable plastic platform. Then position the micro manipulator of the recording micro electrode on a second magnetized stand.

Ensure that the camera and micro manipulator are not disturbed by the movement of other equipment, and do not block illumination from the light source. Afterward, connect the camera to a video monitor and position it towards the platform. Ensure that the setup is properly grounded with copper wire.

In this step, cut a small rectangle of dry PVA sponge that will fit snugly in a 35 millimeter Petri dish with a clean razor blade. Make an additional cut into the sponge to accommodate the reference electrode. Then use a chemical resistant marker to Mark A.Small dot on the sponge where the larva will be placed.

After that, soak the PVA sponge in ringer solution until saturated. Remove and blot it quickly on a paper towel two to three times before placing it in a clean 35 millimeter Petri dish. Subsequently, position the Petri dish containing the sponge on a plastic platform such that the mark can be visualized by the camera.

Next, chloride the electrodes by soaking them in six to 9%sodium chloride. Then air dry them on a Kim wipe for five minutes. Place the silver, silver chloride pellet of the reference electrode into or under the sponge depending on the style of the cut made, and attach the reference electrode lead to the recording system.

Next, attach approximately 40 centimeters of tubing to a five milliliter non lure lock syringe and fill the syringe with ringer solution. Afterward, fill a one milliliter non lure lock syringe with ringer solution. Using a microfill, carefully fill the micro electrode holder and prevent the formation of bubbles.

Then attach the five milliliter syringe to the pressure port of the micro electrode holder with tubing using the microfill and one milliliter syringe filled with ringer solution. Fill the glass micro pipette from the tip and ensure that no bubbles are present. Next, attach the glass micro pipette to the micro electrode holder and be careful to keep the electrode wire straight.

Once secured, use the five milliliter syringe to carefully force the ringer solution through the micro electrode until a tiny amount of solution is visible on the tip. Now carefully place the recording micro electrode in the micro manipulator and attach the lead to the recording system to check for system noise. Place the reference electrode and the tip of the recording micro electrode in a 35 millimeter Petri dish filled with ringer solution.

Check the electrical noise levels of the setup with an oscilloscope or a built-in feature of the ERG apparatus. Noise levels should be no more than plus or minus 10 microvolts from the baseline. In this procedure, cut several one square centimeter paper towel squares, anesthetize three to five larvae in a 35 millimeter Petri dish by adding trica to a final concentration of 0.02%and waiting one to two minutes.

After that, carefully transfer an individual larva onto a paper towel square under a dissecting stereoscope with minimal illumination. Check the position of the larvae choosing a candidate that is dorsal side up with an occluded eye for recording sessions that last longer than 30 minutes. Keep the larva moist by glazing the body with 3%methylcellulose using a fine camel hair brush.

Then transfer the paper towel square with the larva to the damp PVA sponge. Next, apply a continuous stream of water saturated 100%oxygen over the larvae by bubbling the gas through an airstone in a sidearm flask containing distilled water. Position the humidified oxygen from the flask sidearm near the larvas head with tubing under minimal illumination.

Use the micro manipulator and camera to position the micro electrode tip at the midpoint between the nasal and coddle ends of the eye and gently press onto the dorsal limit of the cornea. Allow the larvae to adapt to the dark for five to 10 minutes. Then record the test flash responses to light provided from an LED light source or optical stimulator.

This figure shows a typical ERG recording in a five DPF larval zebra fish. The dark adapted fish were exposed to white LED light for 20 milliseconds with intensities ranging from one to 5, 000 canula per square meter with the onset of the light stimulus at zero seconds. The negative potential a wave is difficult to distinguish, whereas the positive potential B wave is the dominant peak of the wave form.

This inset shows the averaged B wave response amplitudes that have been fit using the NACA rushed in equation. This figure shows a typical ERG recording in a dark adapted five DPF larval zebra fish treated with a PB.The fish are exposed to LED white light for 20 milliseconds with intensities ranging from one to 5, 000 canula per square meter, and the isolated cone mass receptor potential is the dominant element of the waveform. This inset shows the averaged response amplitudes of a PB treated larval zebra fish that had been fit using the NACA rushed in equation.

This figure shows a typical ERG recording in a five D-P-F-A-P-B treated larval zebra fish. Using a dual flash paradigm. The dark adapted fish are exposed to successive 20 millisecond white light.

LED flashes with an intensity of 1000 canula per square meter, and the ISI between the flashes is two seconds. The onset of each light stimulus is indicated here. This figure shows the ratio of the maximum isolated cone mass receptor potential response of the second stimulus to that of the initial stimulus with increasing ISI, which indicates the progressive recovery of photoreceptor sensitivity.

While attempting this procedure is important to remember that the quality of the recordings is dictated by the integrity of the electrode corneal connection and the health of the larvae. The development of this technique has greatly enhanced the ability of researchers in the vision sciences to explore cone adaptation and recovery in the vertebrate retina in vivo.