Imaging Exocytosis in Retinal Bipolar Cells with TIRF Microscopy

This procedure starts with the dissection of the neural retina of adult goldfish, which is the nervous tissue at the back of the eye. Retinas are placed in an enzymatic solution for 35 minutes, rinsed and ated with a fire polished glass pipette to dissociate cells. Cells are then plated on high refractive glass cover slips and mounted into plastic tissue culture dishes.

Isolated bipolar cells are identified by their distinct morphology to load terminals. We puff on them for 10 seconds with a solution containing FM 1 43 and wait 30 seconds before washing the dye away. After a 30 minute wash cells are patch, clamped and imaged with TIRF microscopy.

Hi, my name is Christina Jovi from the ZEC Lab in the Department of Cellular and Molecular Physiology at Yale University School of Medicine. Today we'll show you a procedure to study exocytosis in living retinal bipolar cells using turf microscopy. We use this procedure in the lab to study mechanisms of synaptic release.

So let's get started. To begin dark adapt a goldfish for at least 30 minutes prior to dissection. Following decapitation and eye removal, place one eye bulb on a piece of filter paper and use the tip of a number 11 scalpel blade to puncture the scleral limbus Inside the punctured hole.

Use a pair of bonna scissors to cut the whole anterior segment away. Next place a small piece of filter paper on top of the remaining optic cup and exert some pressure causing it. To soak up the vitreous humor, lift the filter paper with the retina attached and using a pair of VNA scissors cut the optic nerve.

Place the filter paper containing the retina in a 35 millimeter plastic cultured dish containing high alur solution and use number seven dumont tweezers to peel the retina off the filter paper. Next, using half of an industrial carbon steel single edged blade, cut the retina into four to six pieces and let them sit in the hyaluronidase solution for 20 minutes while waiting For the hyaluronidase to take effect, add five milliliters of L cysteine solution to the papain and let it sit for approximately five to 10 minutes until the liquid becomes transparent. Wash the pieces of retina three times in low calcium ringer solution and let them sit in the papain solution for 30 to 35 minutes.

After 35 minutes incubation in the papain solution. Wash the retinal pieces three times. Transfer the pieces to a 35 millimeter plastic cultured dish containing low calcium ringers, and store the dish at 12 to 14 degrees Celsius until you are ready to begin dissociation of the cells to dissociate the cells.

First, fabricate a dissociation pipette by using a bun and burner to heat up the tip of a glass PEs or pipette and then use anatomical forceps to slightly bend it. Put a piece of retina in a micro centrifuge tube containing low calcium ringer solution and slowly iterate the retina by carefully pipetting it up and down with the fabricated dissociation pipette. Being careful not to produce any air bubbles plate the isolated cells by adding a drop of the retinal suspension to a homemade recording chamber filled with low calcium ringer solution.

The chamber consists of the bottom half of the 35 millimeter plastic culture dish with a circular hole in the middle and a circular cover slip made of 1.78 refractive index glass glued to the bottom. With dissection and bipolar cell isolation complete, we are ready to begin bipolar cell loading and washout out. In this experiment, it is best to use an objective type total internal reflectance fluorescence microscope with a very high numerical aperture objective and a sensitive camera.

In this video, we will be using a 1.65 numerical aperture objective and an electron multiplying charge coupled device. To begin, add a drop of high refractive index liquid to the microscope.Objective. Place the recording chamber on top of the microscope.Objective.

Carefully mount the ground electrode and superfusion exit pipe to the chamber. Let the chamber sit on the microscope for 10 to 20 minutes, allowing the cells to sink and adhere to the bottom. Next, purge the superfusion lines and add washing solution.

Low calcium ringer solution and control ringer solution to the superfusion system. Turn the microscope brightfield light on and search for intact bipolar cells. Slightly tap the microscope to make sure that the neurons are firmly attached to the bottom of the chamber.

Position the superfusion pen close to an intact cell and continuously perfuse the preparation with low calcium ringer solution. Now turn the lights off in the room and add a red long pass Filter to the optical path to minimize excitation of the fluorescent dye. Fill a loading pipette with 10 microliters of fluorescent dye solution.

Mount it in the micro manipulator and then lower it onto the preparation without using the over pressure option until it is at the same focal plane as the selected bilar cell. Position the puffer opening at a distance of around 10 micrometers from the axon terminal. Turn the superfusion system off and puff the dye solution for 10 seconds by turning the pipette over pressure on.

Now turn the over pressure off and without moving the pipette. Wait 30 seconds, turn the superfusion system on and bathe the chamber in washing solution for five minutes. In the meantime, remove the puffer pipette from the bath.

After five minutes, switch the SUPERFUSION system to low calcium ringer solution and perfuse it for 25 to 30 minutes, allowing for the removal of excess dye With bipolar cell loading and washout complete. We are ready to begin patch clamping and T-I-R-F-M imaging. After the washout is complete, change electrode holders to avoid contamination of your patch pipette with dye.

Place the axon terminal in the center of the field of view. Fill a patch pipette with seven microliters of internal solution. Tap it to get rid of air bubbles.

Coat it with molten dental wax and then mount it in the micro manipulator. Turn the pipette over pressure on and slowly lower the pipette onto the preparation. Check the pipette resistance and make any needed corrections.

Next, with the superfusion system set to control ringer solution, create a giga seal between the pipette and the cell by slightly touching the tip of the electrode against the cell body. With the pipette over pressure off, gently apply negative pressure to the electrode while sealed. Select whole cell mode in the amplifier and set the cell holding potential to negative 60 millivolts.

Now use imaging software to select a region of interest that encompasses the whole axon terminal. Turn the bright field light off and by briefly exposing the terminal to the 488 nanometer laser. Find the right focal plane for TIRF imaging.

Next break into the cell by using the zap command of the amplifier while applying slightly negative pressure to the pipette correct for cell capacitance and series resistance. And then apply the voltage protocol while imaging the movement of synaptic vesicles. Wait at least 40 seconds between trials to allow for recovery.

First, we choose a bipolar cell in brightfield microscopy. We then zoom in and load its axon terminal with the fluorescent dye. After a 30 minute wash, the loaded terminal can be imaged inter microscopy along with the movement of fluorescent glutamatergic vesicles.

Here we introduce a rib eye binding peptide through the patch pipette, which allows us to see the position of synaptic ribbons when imaged with a 561 nanometer laser. We then depolarize the cells and image vesicle movement with the 488 nanometer laser. We have just shown you how to dissociate plate and load retinal bipolar cells with a fluorescent dye as well as to patch, clamp these neurons and simultaneously image them with turf microscopy.

When doing this procedure, it's important not to expose the cells too much to the laser. So that's it. Thanks for watching and good luck with your experiments.