Name: Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques (Video) | JoVE Journal
Uploaded: 2015-02-11T14:00:00.000+00:00
Duration: 10 min 30 s
Description: Learn patch clamp techniques to record light-evoked postsynaptic responses in dark-adapted mouse retinal neurons. This method enables precise electrophysiological analysis of retinal circuitry.

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5 minutes

This work was supported by NIH R01 EY020533, WSU Startup Fund, and RPB grants.

Ethics Statement: Procedures involving animal subjects were approved by the Institutional Animal Care and Use Committee (IACUC) at Wayne State University.
1. Preparation of Experimental Solution
<ol>
	<li>Prepare the dissecting solution 1 day to 1 week before the actual experiment. Use a HEPES buffer solution for retinal dissection because of its strong buffering ability at lower temperatures16. Mix all chemicals as follows (in mM): 115 NaCl, 2.5 KCl, 2.5 CaCl2, 1.0 MgCl2, 10 HEPES, 28 glucose. Adjust pH to 7.4 with NaOH. Keep the solution in a refrigerator up to 1 week.</li>
	<li>The recording solution is Ames&#8217; medium, which is an artificial cerebral spinal fluid (aCSF) designed for retinal preparations17. By the day of the experiment, weigh out Ames&#8217; powder (4.4 g for 500 ml solution) in a tube. Also, weigh out NaHCO3 (0.95 g for 500 ml solution) and mix with the Ames&#8217; powder.</li>
	<li>Prepare the intracellular pipette solution for patch clamp recordings by the day of the experiment. Mix all chemicals as follows (in mM): 111 K-gluconate, 1.0 CaCl2, 10 HEPES, 1.1 EGTA, 10 NaCl, 1.0 MgCl2, 5 ATP-Mg, and 1.0 GTP-Na. Adjust pH to 7.2 with KOH. Filter the pipette solution with a conventional syringe filter. Make ~500 &#181;l aliquots and store them in a freezer or a deep freezer.</li>
</ol>
2. Preparation for the Day of Experiment
<ol>
	<li>The night before the experiment, place a mouse in a cage (C57BL/6J or similar background, 4 - 8 weeks old, male) in a dark space for O/N dark adaptation.</li>
	<li>Preparation for dissection:
	<ol>
		<li>Fill dissecting solution (100 - 200 ml) in a glass beaker, place it on ice, and bubble with oxygen for at least 10 min.</li>
		<li>Start oxygenation of the dark box for retinal preparation storage.</li>
		<li>Prepare plastic coverslips with grease rails for retinal slice preparations and place each of them in a 35 mm plastic dish.</li>
		<li>Attach a new razor blade to a chopper (tissue slicer).</li>
		<li>Cut a filter membrane into two halves. This is for retinal slicing.</li>
		<li>Align the dissecting tools and transfer pipettes near the dissecting microscope. The work area must be well organized so that each tool can be easily accessed in the dark.</li>
	</ol>
	</li>
	<li>Preparation for patch clamp recordings:
	<ol>
		<li>Dissolve Ames&#8217; powder and NaHCO3 in distilled water (Ames 4.4 g and NaHCO3 0.95g/500 ml ddH2O). Bubble the Ames&#8217; medium with mixed oxygen (95% O2 and 5% CO2) for at least 30 min while heating to a recording temperature level (~35 &#176;C). Adjust the pH to 7.4 with NaHCO3 while still bubbling.</li>
		<li>Thaw the internal pipette solution during the dissection. After it is thawed, add 0.01% sulforhodamine B for intracellular staining.</li>
	</ol>
	</li>
</ol>
3. Retinal Dissection
<ol>
	<li>In a dark room, euthanize the mouse using carbon dioxide and bilateral pneumothorax. After 2 - 3 min of using the carbon dioxide, the mouse will lose consciousness.</li>
	<li>When the mouse no longer responds to a tail pinch, quickly enucleate the eyes and place them in a cold dissecting solution in a plastic dish. Perform the bilateral pneumothorax to ensure the animal&#8217;s death. Quickly store the eye in the dish in the dark box. Use an infrared viewer to conduct the procedure in the dark. 
	NOTE: Alternatively, isoflurane, decapitation, or cervical dislocation can be used for euthanasia. The method of euthanasia should follow the guidelines of the National Institute of Health (NIH) and/or the Institutional Animal Care and Use Committee.</li>
	<li>In a dark room, adjust the dissecting microscope and infrared illumination. Under the microscope, transfer an eye from its dish into a 10 cm plastic dish with the cooled dissecting solution. Continuously bubble oxygen through tubing placed on the bottom of the plastic dish. Adjust the flow volume so it is not too low, but ensure that it is not high enough to disturb the dissection.</li>
	<li>Cornea and lens removal:
	<ol>
		<li>Make an incision on the top of the cornea with a micro-surgical knife. Holding the optic nerve at the other end of the eyeball prevents the eye from moving. Extend the cut using a pair of surgical scissors. Cut out the cornea with some sclera around it.</li>
		<li>Grab the lens with a fine forceps and slowly pull the lens out. If the vitreous is too sticky to remove the lens, cut the vitreous with a pair of scissors.</li>
		<li>Once the eye-cup is made, gently pour cold, oxygenated dissecting solution into the eyecup with a small transfer pipette. 
		NOTE: The eyeball quickly deflates after an incision is made in the cornea. It is important to maintain the shape of the eye as deforming the eye during this procedure will damage the retina and cause retinal detachment.</li>
	</ol>
	</li>
	<li>Identifying the dorsal and ventral sides of the retina: 
	NOTE: Because of uneven distribution of the green and the ultraviolet (UV) cones18, identifying a specific retinal area is important for light response recordings.
	<ol>
		<li>Identify the line going across the retinal eye cup passing near the optic nerve head. This line passes mostly across the ventral side of the optic nerve; the side including the optic nerve is the dorsal side, while the other is ventral 19. See this landmark clearly under infrared viewers.</li>
		<li>Depending on the purpose of the experiment, make a cut on the unnecessary side. This will be useful after isolating the retina from the eye cup.</li>
	</ol>
	</li>
	<li>Vitreous removal:
	<ol>
		<li>Optionally, incubate the eye-cup with hyaluronidase (0.5 mg/ml) for 15 min.</li>
		<li>Remove the vitreous with extra-fine forceps. The vitreous is mainly attached to the bottom and the outer edge of the eye-cup. Gently remove it without touching or poking the retina. As the vitreous might be tightly attached to the eyecup, remove gently to avoid retinal detachment from the eye-cup. Continue to pull until no tension is felt. 
		NOTE: This step is particularly important for dissecting the mouse retina; however, it is not a problem for the retinas of other species such as the rat or salamander.</li>
	</ol>
	</li>
	<li>Isolate the retina from the eye-cup. Grab the sclera and gently peel off the retina using the backside of forceps. 
	NOTE: Whole-mount retinal preparations can be made from the isolated retina. Make three to four cuts at the edges and flatten the retina, or cut the retina into several pieces.</li>
	<li>Trim the retina and discard the unnecessary half of the retina (step 3.5). In the same plastic dish, cut the remaining retina into two pieces. For each piece of the retina, cut off the edges to make a retinal slab, and trim the folding edges.</li>
	<li>Transfer a retinal slab onto a glass plate using a large transfer pipette (~2 ml). Suck up the excess solution with a small pipette, use a piece of filter paper to flatten the slab, and then quickly place a piece of the filter membrane on top of the tissue.
	<ol>
		<li>Using a small transfer pipette, place a drop of dissecting solution on the filter membrane. Wait ~15 sec until the solution spreads over the membrane. During this time, the retinal tissue will stick to the filter membrane.</li>
		<li>Pour more cold dissecting solution under and around the filter. The retinal tissue with the filter membrane will float. Grab the filter membrane with a forceps and place it in the slicing chamber. Pour dissecting solution over the tissue, and store the preparation in the dark box. 
		NOTE: The procedure described here is critical. It is important to perform these steps quickly to ensure that the retinal tissue sticks to the filter membrane and does not dry out.</li>
	</ol>
	</li>
	<li>Using a chopper, cut the retinal tissue into slices. Around 200 &#956;m thickness is feasible for patch clamp studies.</li>
	<li>Place a plastic coverslip with grease rails in the slicing chamber. Transfer a retinal slice on top of the grease rails on a plastic coverslip. Rotate the slice 90&#176; so that the transverse section is visible. Press the filter membrane down onto the coverslip, then, cover the sides of the filter paper with grease.
	<ol>
		<li>After the slice is immobilized on a plastic coverslip, grab the coverslip and transfer it to a 35 mm plastic dish. Pour cold dissecting solution onto the retinal preparation and submerge the preparation. Store each dish in the dark box, which should be continuously oxygenated. 
		NOTE: The entire procedure needs to be done carefully so that the filter membrane is not deformed. Otherwise, the retinal slice easily separates from the filter membrane.</li>
	</ol>
	</li>
</ol>
4. Patch Clamp Recordings from Retinal Slice Preparation
<ol>
	<li>Recording preparation:
	<ol>
		<li>Prime the perfusion tubes with Ames&#8217; medium. Allow all the bubbles to pass through when filling the tubing.</li>
		<li>Make patch clamp recording pipettes with a puller. To fill a pipette with the pipette solution, dip in the backside of a pipette for 1 min until the pipette tip is backfilled. Next, fill ~1/3 of the pipette using a micro-pipette filler. Store each pipette in a moist pipette box.</li>
		<li>Turn on the equipment for patch clamp recordings; including the computer, amplifier, CCD camera, and microscope.</li>
	</ol>
	</li>
	<li>Shut off the room light. Place a retinal slice preparation onto the microscope stage chamber using an infrared viewer. After it is immobilized, begin continuous perfusion. Set the perfusion temperature at 33 to 37 &#176;C.</li>
	<li>View the slice surface with the CCD camera. Focus on the target location where the target cell types reside. Select a healthy-looking cell for patch clamp recording (i.e., it should have smooth-looking surface and a good cell shape).</li>
	<li>Place a recording pipette in a pipette holder. Advance the pipette to the slice preparation. When it is close to the slice preparation (~2 mm above), find the tip of the pipette with the microscope. Once the pipette tip is visible under the microscope, move the tip down towards the target cell.</li>
	<li>Set the amplifier. Adjust the pipette at 0 mV/0 pA (zeroing) and start a continuous electric pulse of ~5 mV at ~10 Hz. Check the pipette resistance; ideal resistance is between 5 and 10 M&#8486; for most retinal neurons.</li>
	<li>Start blowing out from the tip. Use a mouthpiece, or use a syringe, to apply positive pressure when the pipette dips into the bath solution. Continuously blow out the internal solution until the tip is on the surface of the target cell.
	<ol>
		<li>When the positive pressure makes a small dimple on the cell surface, advance the tip slightly and stop blowing out. Check the pipette resistance, which should increase. If it is continuously increasing, leave it and monitor the resistance until it reaches &#62;1 G&#8486; (gigaseal). If the resistance does not spontaneously increase, gently apply negative pressure until it becomes a gigaseal.</li>
	</ol>
	</li>
	<li>After the gigaseal is achieved, change the holding potential to -70 mV. Then, intermittently apply negative pressure to rupture the membrane inside the pipette tip. When the whole-cell configuration is made, the pipette resistance can be between 500 M&#8486; and 1 G&#8486;, and the capacitive current is seen. Sometimes spontaneous postsynaptic currents can be observed.</li>
	<li>Record the I-V relationship from -80 to +40 mV. Different types of voltage-gated channels are activated depending on the cell type.</li>
	<li>Record light-evoked synaptic currents or voltages.</li>
</ol>

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The authors have nothing to disclose.

Good recordings can only be achieved with good retinal preparations and well-designed patch clamp setups. Although all the steps described above are important, the discussion highlights some critical steps both for the dissection and recordings. 
For dissection, two things are especially important: cooling and oxygenation. After enucleating the eye, quickly remove the front part of the eye in a dissecting chamber with oxygen-bubbled, cooled dissecting solution, and pour cold solution into the ...

The retina is one of the unique parts of the nervous system. As an accessible part of the brain, its synaptic architecture has been well characterized. In addition, the functions of this neural network can be examined with a physiological stimulus: light. If the retinal tissue is isolated in a dark room with appropriate procedures, neurons in the tissue will respond to light. This preparation has been used to study visual signal processing and elucidate various synaptic mechanisms and neural network functions, as well as disease mechanisms. 
Light responses in retinal neurons have been recorded for decades. Early studies used sharp electrodes to make intracellular recordings from mudpuppy retinal neurons1. In the 1980s, the patch clamp technique was invented2, and soon became a popular method among vision researchers3,4. Single cell recordings from lower vertebrates, including mudpuppy and fish retinal neurons, were popular methods that contributed to the elucidation of visual signal processing mechanisms5,6. 
After genetic mutation techniques were developed, the mouse retina became a more popular model for vision researchers7-9. The mammalian retina is more attractive than that of lower vertebrates because it is evolutionarily closer to the human retina, and there is an opportunity to use disease models. However, mouse retinal cells are small and fragile10, and making retinal preparations and conducting patch clamp recordings in a dark room is challenging. As technology has improved, diverse approaches have become available to study visual signaling mechanisms such as imaging studies11 and the electroretinogram (ERG)12. Nevertheless, single cell recording with the patch clamp method is still important because it is highly temporally and spatially sensitive compared to other methods. Therefore, we have continuously conducted patch clamp recordings and improved our methods to investigate visual signal processing in mouse retinal slice preparations13-15.
In this video tutorial, the protocols are presented with important tips. Good recordings can only be achieved with good preparation. Practicing animal dissection and building a sturdy patch clamp rig will enable most researchers to achieve successful recordings.

<table><tbody><tr><td>Mice (28-60 days old, male)</td><td>Jackson laboratory</td><td>C57BL/6J strain</td><td></td></tr><tr><td>Ames&#039; medium powder</td><td>Sigma</td><td>A1420</td><td>excellent</td></tr><tr><td>Stereo microscope</td><td>Nikon</td><td>SMZ745</td><td>excellent</td></tr><tr><td>Dissecting tool: forceps</td><td>Dumont</td><td>#4, #5, #55</td><td>excellent</td></tr><tr><td>Dissecting tool: scissors</td><td>Roboz</td><td>RS-5605</td><td>excellent</td></tr><tr><td>Dissecting tool: surgery knife</td><td>Surgistar</td><td>7514</td><td>excellent</td></tr><tr><td>Razor blade (for chopper)</td><td>EMS</td><td>71970</td><td>excellent</td></tr><tr><td>Chopper</td><td>handmade</td><td></td><td></td></tr><tr><td>Infrared viewer</td><td>Night Owl Optics</td><td>NOBG1</td><td>It shows bright view. Focusing small objects is an issue.</td></tr><tr><td>Puller</td><td>Sutter</td><td>P-1000</td><td>excellent; makes consistent size pipettes.</td></tr><tr><td>Dark box</td><td>Pelican</td><td>dark box</td><td>excellent</td></tr><tr><td>Patch clamp system</td><td>Scientifica</td><td>slice scope 2000</td><td>Excellent setup. Most key components are included in one package. Micromanipulators are excellent.</td></tr><tr><td>Amplifier</td><td>Molecular Devices</td><td>multiclamp 700B</td><td>Excellent and easy control.</td></tr><tr><td>Acquiring software</td><td>Molecular Devices</td><td>pClamp software</td><td>Excellent and easy control.</td></tr><tr><td>Light source (LED)</td><td>Cool LED</td><td>pE-2 4 channel system</td><td>Excellent</td></tr><tr><td>CCD camera</td><td>Q-imaging</td><td>Retiga 2000</td><td>Excellent</td></tr><tr><td>Faraday cage</td><td>handmade</td><td></td><td></td></tr></tbody></table>

recording light-evoked postsynaptic responses in neurons in dark-adapted, mouse retinal slice preparations using patch clamp techniques

Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques

The retina is the gateway to the visual system. To understand visual signal processing mechanisms, we investigate retinal neural network functions. Retinal neurons in the network comprise of numerous subtypes. More than 10 subtypes of bipolar cells, ganglion cells, and amacrine cells have been identified by morphological studies. Multiple subtypes of retinal neurons are thought to encode distinct features of visual signaling, such as motion and color, and form multiple neural pathways. However, the functional roles of each neuron in visual signal processing are not fully understood. The patch clamp method is useful to address this fundamental question. Here, a protocol to record light-evoked synaptic responses in mouse retinal neurons using patch clamp recordings in dark-adapted conditions is provided. The mouse eyes are dark-adapted O/N, and retinal slice preparations are dissected in a dark room using infrared illumination and viewers. Infrared light does not activate mouse photoreceptors and thus preserves their light responsiveness. Patch clamp is used to record light-evoked responses in retinal neurons. A fluorescent dye is injected during recordings to characterize neuronal morphological subtypes. This procedure enables us to determine the physiological functions of each neuron in the mouse retina.

A representative slice preparation is shown in Figure 1. The slice preparation is in a straight position, showing photoreceptors to ganglion cells in a flat surface, and no detachment from the filter paper. If a slice is tilted, only part of the preparation is in focus, which makes it difficult to identify an appropriate cell for patch clamping. For recordings, it is important to select a good soma, which usually has a shiny surface, an even round shape, and no visible dark plaques. After the whole cell ...

We will demonstrate how to prepare retinal slices from the mouse eye and record light responses in retinal neurons. The entire procedure is conducted in dark-adapted conditions.

The retina is the gateway to the visual system. To understand visual signal processing mechanisms, we investigate retinal neural network functions. ...

The overall goal of this experiment is to obtain light response recordings from dark adapted retinal neurons. This is accomplished by first harvesting a mouse eye. The second step is to isolate the retina and cut it for slice preparations.Next, the retinal preparation is mounted and viewed on a microscope to find a target cell for patch clamp recordings. After establishing a whole cell patch clamp configuration, a light stimulus is applied in the vicinity of the cell. Finally, as simultaneous electrophysiological response can be observed through acquisition software.The main advantage of this technique of existing methods like KRC imaging, is that the recording sensitivity is high, and that recording cells can be morphologically identified. Generally, individuals new to this method will struggle because working in total darkness can be difficult and disorienting for those unaccustomed to doing so. To begin a range, the dissecting tools and transfer pipettes near the dissecting microscope.The work area will be well organized so that each tool can be easily accessed in the dark. When ready, Dawn an infrared viewer to conduct the procedure in the dark. After euthanasia quickly in nucleate the eyes and place them in a dish of cold dissecting solution, kept in a dark box working under a microscope.Transfer an eye into a dish with cool dissecting solution. Continuously bubbled with oxygen. To remove the cornea and lens, hold the optic nerve at the other end of the eyeball to keep it from moving and make an incision on the top of the cornea with a microsurgical knife.Extend the cut using a pair of surgical scissors and remove the cornea, leaving some sclera surrounding it. Next, grab the lens with fine forceps and slowly pull it out. Once the eye cup is made, gently pour cold oxygenated dissecting solution into the eye cup with a small transfer pipette.Work quickly to maintain the shape of the retina and use gentle pressure to identify the dorsal and ventral sides of the retina. Use infrared viewers to identify the line going across the retinal eye cup, passing near the optic nerve head. This line passes mostly across the ventral side of the optic nerve and the side, including the optic nerve, is the dorsal side.Depending on the experiment, make a cut on either the ventral or dorsal side. This marks the unnecessary side and is a useful marker moving forward. Next, gently remove the vitreous with extra fine forceps without touching the retina.The vitreous is mainly attached to the bottom and the outer edge of the eye cup. As the vitreous might be attached tightly to the eye cup, proceed gently to remove retinal detachment. Continue to pull until no tension is felt.Once the retina is isolated, grab the sclera and gently peel off the retina. Using the backside of the forceps, trim the retina and discard the unnecessary half in the same plastic dish. Cut the remaining retina into two pieces for each piece.Cut off the edges to make a retinal slab and trim the folding edges. Next, use a large transfer pipette to transfer a retinal slab onto a glass plate. Suck up the excess solution with a small pipette and use a piece of filter paper to flatten the slab.Gently place a piece of the filter membrane on top of the tissue using a small transfer pipette. Place a drop of dissecting solution on the filter membrane. Wait about 15 seconds while the solution spreads and sticks the retinal tissue to the filter membrane.Once stuck together, pour more cold dissecting solution under and around the filter until the membrane floats. Next, place the filter membrane into the slicing chamber and cover with dissecting solution. Working quickly to ensure that the membrane does not dry out, use the chopper to cut the retinal tissue into slices.Place a plastic cover slip with grease rails into the slicing chamber. Transfer a retinal slice on top of the grease rails and rotate the slice 90 degrees so that the transverse section is visible. Press the filter membrane down onto the cover slip before covering the sides of the filter paper with grease.After the slices immobilized on a plastic cover slip, transfer it to a 33 millimeter plastic dish. Submerge the preparation with coal dissecting solution and store each dish in a dark box that is continuously oxygenated. For patch clamp recordings.Prime the profusion tubes with aims medium and allow all the bubbles to pass from the tubing. After making recording pipettes with a glass polar, backfill the tip with pipette solution. Next, use a micro pipette filler to fill about one third of the pipette once filled, store each pipette in a moist pipette box.Next, turn on the equipment for patch plant recordings, including the computer amplifier CCD camera and microscope working in dark conditions. Place a retinal slice preparation onto the microscope stage with the aid of an infrared viewer. After it is immobilized, begin continuous perfusion.Set the perfusion temperature to 33 to 37 degrees Celsius. View the slice surface with a CCD camera. Focus on the target location.With the target cell types reside, select a healthy looking cell for a patch clat recording. Once a healthy cell is identified, place a recording pipette in a pipette holder and advance the pipette to the slice preparation. When it is close to the slice preparation, find the tip of the pipette with a microscope.Once the pipette tip is visible under the microscope, move the tip down towards the target cell. Next, set the amplifier and adjust the pipette at pika amp. Five volts per Nana amp.Start a continuous electric pulse of approximately five millivolts and approximately 10 hertz and check the pipette resistance. An ideal resistance is between five and 12 mega ohms. For most retinal neurons.When ready, use a mouthpiece or syringe to blow out the internal solution until the tip of the pipette is on the surface of the target cell. When the positive pressure makes a small dimple on the cell surface, advance the tip slightly and stop blowing out. If the pipette resistance is continuously increasing, leave it and monitor the resistance until it reaches greater than one giga ohm.If the resistance does not spontaneously increase, gently apply negative pressure until it becomes a giga seal. After the giga seal is achieved, change the holding potential to negative 70 millivolts then intermittently apply negative pressure to rupture the membrane inside the pipette tip. When the whole cell configuration is made, the pipette resistance can be between 500 mega ohms and one giga ohm, and the capacitive current is observable.Record the IV relationship from negative 80 to 40 millivolts. Different types of voltage gated channels will be activated depending on the cell type. Lastly, record light evoked synaptic currents or voltages.This slice preparation viewed at 10 x shows both photoreceptors and ganglion cells with no detachment from the filter paper. When this preparation is viewed with a 60 x objective, each cell layer is clearly observed. Light evoked excitatory postsynaptic potentials from retinal neurons were evoked in response to a step light.The peak amplitude and decay time vary depending on the cell type and subtype. The cell type and its morphological subtype were revealed by sulfur labeling after physiological recordings After its development. This technique paved the way for scientists in the field of retina research to study the functions of natural visual signaling, as well as the contributions of synaptic mechanisms.

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Research

JoVE Journal

Neuroscience

This video shows the process of whole-cell voltage clamp recordings in the retinal slice of the aquatic tiger salamander. We demonstrate the preparation of the slice as well as how to perform patch clamp recordings during visual stimulation of the retina.

Whole-cell Recordings of Light Evoked Excitatory Synaptic Currents in the Retinal Slice

Here we describe a procedure for generating dark-adapted slices of the mouse retina for electrophysiological recordings.

Patch Clamp Recordings from Mouse Retinal Neurons in a Dark-adapted Slice Preparation

This article provides a description of how to dissect and record from the isolated retinal preparation in mouse. In particular, we describe how to record light responses from a fluorescently labeled ganglion cell population and subsequently identify and analyze its morphology.<br...

An Isolated Retinal Preparation to Record Light Response from Genetically Labeled Retinal Ganglion Cells

We describe a relatively simple method of transretinal electroretinogram (ERG) recordings for obtaining rod and cone photoresponses from intact mouse retina. This approach takes advantage of the block of synaptic transmission from photoreceptors to isolate their light responses and record them using field electrodes placed across the isolated flat-mounted retina.

Transretinal ERG Recordings from Mouse Retina: Rod and Cone Photoresponses

This video demonstrates a whole-cell patch-clamp technique for electrophysiological recording from a flat-mount mouse retina. A thin glass pipette with an electrode is used to approach starburst amacrine cells (SACs) within the retina. After forming a seal, suction is applied to rupture the membrane and form the whole-cell patch-clamp configuration. Finally, the flow of ions through the SAC synaptic receptors is measured using the configuration.

Establishing a Whole-Cell Patch Clamp for Electrophysiological Recording from a Flat-Mount Mouse Retina

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Photostimulation and Whole-Cell Patch Clamp Recordings of Neurons in Mouse Hippocampal Slices

This video demonstrates the recording of light-evoked post-synaptic responses in a dark-adapted mouse retinal tissue slice preparation. The retinal slice is placed in a recording chamber, and a recording pipette is used to establish electrical continuity with a retinal ganglion cell. Light pulses are applied to generate excitatory post-synaptic potentials in the ganglion cells, which are recorded by the pipette.

Light-Evoked Postsynaptic Responses in Neurons of Mouse Retinal Slices

We describe the preparation of thin retinal slices from aquatic tiger salamanders (Ambystoma tigrinum) and explain how we use these slices to study synaptic processing in the retina by obtaining dual whole-cell voltage clamp recordings from photoreceptors and second-order horizontal and bipolar cells.

Simultaneous Whole-cell Recordings from Photoreceptors and Second-order Neurons in an Amphibian Retinal Slice Preparation

This protocol demonstrates how to perform whole-cell patch clamp recording on retinal neurons from a flat-mount preparation.

Patch Clamp Recording of Starburst Amacrine Cells in a Flat-mount Preparation of Deafferentated Mouse Retina

We developed a horizontal slice preparation of the adult mammalian retina with the plane of sectioning parallel to the retinal surface. Dendritic fields of nonradially oriented retinal neurons were not truncated, allowing the study of signal processing with patch-clamp and imaging techniques.

Preparation of Horizontal Slices of Adult Mouse Retina for Electrophysiological Studies

In this paper, we discuss three brain preparations used for whole cell patch clamp recording to study the retinotectal circuit of Xenopus laevis tadpoles. Each preparation, with its own specific advantages, contributes to the experimental tractability of the Xenopus tadpole as a model to study neural circuit function.

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

Learn patch clamp techniques to record light-evoked postsynaptic responses in dark-adapted mouse retinal neurons. This method enables precise electrophysiological analysis of retinal circuitry.

Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques (Video) | JoVE Journal

Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques

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