Today I will show you how To prepare retinal slices of the tiger salamander for patch clamp recordings. First, I will prepare two kinds of microscope slides. We use slides that have an adhesive well attached to them so that we can use them as fluid chambers.
On the first slide, I place two parallel Bands of vacuum grease. I then take some filter paper and cut it Into rectangular pieces with a raise of light. Now I place one of the pieces on the grease rails and push it down so it's well attached to the slide.
I do the same for the other piece. The second kind of slide has little rubber blocks glued to it. For now, I Just add a little vacuum grease In between the blocks, I also put a new Piece of razor blade in the tissue slicer.
Finally, I set out the tools I will need during the dissection, which are two pairs of forceps, one pair of scissors, and a tool to remove the eye from the head to record light evoked currents. One should perform the dissection under IR light, but for demonstration purposes, I will now work in the light. I first remove the eye and place it the microscope on a piece of tissue paper, which I moistened with ice cold ringer solution.
I then remove extra connective tissue from the eyeball to make sure that it won't damage the eye. I always cut along the Surface. This is what you see through the microscope.
Next, I use a razor blade to make a little slit in the cornea. To do that, I hold Down the eye with the forceps while pushing down the edge of the razor blade on the cornea. I then hold the cornea with my forceps while removing it with the scissors to Take out The lens.
I put some pressure on the edges of the eye cup so that I can easily access the lens with my forceps. Next, I remove the iris. I hold onto the iris with the forceps while cutting along the aura serato until only the eye cup remains.
I then cut the I cup into two rectangular pieces. I first cut it in half and then trim the top and bottom edges. Now I take one of the slides that I prepared earlier, pick up a piece of I cup and place it retina Side down on the filter paper.
I then pull up this sclera so that only the retina remains on the slide. After that, I quickly add, ring our solution to the chamber. Let's repeat The procedure for the second piece of retina.
But this time you can watch through the microscope while putting the retina down on the filter paper. I use the forceps in my left hand to carefully push down on the sclera. Finally, I cut the retina into 200 to 300 micrometer wide slices.
Next, I choose a slice to Use for my experiments. To pick a good slice, I need to rotate the slices by 90 degrees so that I can see the Layers of the retina. I pick a slice that looks undamaged and shows distinct horizontal bands indicating the different layers.
To move the slides from one slide to another, I create a ringers bridge and then carefully move the slides through the solution. When I put down the slice, I push it against the rubber blocks so that the filter paper is oriented vertically to the glass. Now I am ready for patch clamp recording.
We store the intracellular Solution in small portions in the freezer so that we only saw the amount we actually need in one day. Before filling the electrodes, we filter the solution to remove small particles that could clock the electrode tip. The patch electrodes are pulled from bur silicate capillary glass to record from ganglion cells.
I choose a pull a setting that will produce electrode resistances between two and four mega ohms. When I fill the electrodes, I want to make sure that there are no air bubbles in the electrode tip. Flicking the glass with your finger helps to remove bubbles.
This is the electrode holder attached to the head stage. The electro roller contains a silver chloride wire. This is the ground electrode with an niga bridge.
Here's the perfusion pipette. It is connected to an eight channel micro perfusion system. The main perfusion is gravity driven, so it's just a bottle filled with wingers, and the solution trips down through this tube into the recording chamber.
To be able to record in the dark, I have an IR camera attached to the side port of the microscope, which is connected to a TV screen. On the top port, I connected an image projector, which allows me to project visual stimuli onto the retina. And finally, the patch clamp amplifier.
I now place the retinal slice under the microscope and add the ground electrode to the bath to set up the main profusion. And next, Add the tube from the gravity profusion system. The glass tube on the other side is connected to vacuum suction, which keeps the fluid level constant.
I also make sure that a suction tube is connected to The electro shoulder. Next, I position the micro perfusion pipette. I lower it in the bath Following with the microscope objective.
Once I get close to the slice, I pull it back a little bit so that the solution will hit the slice. When I turn on the perfusion, I should see the slice move slightly like here. Let's now look for a good cell to record from.
We are looking for ganglion cells, which are at the bottom of the slice. Now I get the electrode ready before entering the bath with the electrode, I add some positive pressure. Once I found the electrode under the Microscope, I lower it down under its right above the slice to patch onto a cell.
I switched to the 40 x objective. Now I zero the baseline current and check the electrode resistance. The seal test is A sequence of 10 millivolt pulses, which helps me to see whether I established a tight seal between the electrode and the cell.
I now lower my electrode on the target cell. Once I see a dimple on the cell Body, I release the positive pressure from the tube, which establishes a tight seal. The resistance is now so high so that the current trace looks flat.
The little spikes are capacitance artifacts from the electrode. These can be compensated for like this, so now they're gone. So now I will try to break into the cell.
To do that, I carefully put little suction pulses on the tube connected to the electrode shoulder, and if everything works out right, you see the current trace change like this. The spikes you can see now are crossed by the membrane capacitance and also can be compensated for this now is a preparation that was prepared in the dark. Let's see whether we can record light evoked responses.
The trace you can see on the oscilloscope is the current response of a ganglion cell voltage clamped at minus 60 millivolts. The light stimuli we use are bright and dark bars presented on a steady uniform background. Our lab use software shows the complete record of one trial.
One can see that this is an off cell as a response to the onset of a dark bar and the offset of a bright bar. The white line on the top indicates the duration of the stimulus. We can now run experiments in which we perfuse different pharmacological agents.
For example, we can block GABAergic and Gly synergic inhibition. After waiting for less than one minute, we can Usually start to see the effects of the drug here. The response became stronger and more prolonged.
The retinal slice Preparation has been used widely to record the synaptic currents from different cells in the retina, and if one adds a fluorescent dye to the electrode solution, the cell type can be easily confirmed. Anatomically, here we use the larval TGA salamander as an animal model. The fact that salamanders are cold-blooded animals and have large retinal cells provides advantageous recording conditions, but the same technique can be applied to mammalian preparations.
