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

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 ...

Watch this Scientific Journal Video about Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques at JoVE.com

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

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. ...

recording-light-evoked-postsynaptic-responses-neurons-dark-adapted

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JoVE Journal

Neuroscience

8.4K Views.  Wayne State University School of Medicine. 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.

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

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

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of intracellular events that lead to closure of cyclic-nucleotide-gated channels in the cell membrane. The resulting change in membrane potential leads in turn to reductions in the amount of neurotransmitter release from both rod and cone synaptic terminals. To measure how the light-evoked change in photoreceptor membrane potential leads to downstream activity in the retina, scientists have made electrophysiological recordings from retinal slice preparations for decades1,2. In the past these slices have been cut manually with a razor blade on retinal tissue that is attached to filter paper; in recent years another method of slicing has been developed whereby retinal tissue is embedded in low gelling temperature agar and sliced in cool solution with a vibrating microtome3,4. This preparation produces retinal slices with less surface damage and very robust light-evoked responses. Here we document how this procedure can be done under infrared light to avoid bleaching the visual pigment.

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

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of ...

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

The first steps in vertebrate vision take place when light stimulates the rod and cone photoreceptors of the retina 1. This information is then segregated into what are known as the ON and OFF pathways. The photoreceptors signal light information to the bipolar cells (BCs), which depolarize in response to increases (On BCs) or decreases (Off BCs) in light intensity. This segregation of light information is maintained at the level of the retinal ganglion cells (RGCs), which have dendrites stratifying in either the Off sublamina of the inner plexiform layer (IPL), where they receive direct excitatory input from Off BCs, or stratifying in the On sublamina of the IPL, where they receive direct excitatory input from On BCs. This segregation of information regarding increases or decreases in illumination (the On and Off pathways) is conserved and signaled to the brain in parallel.
The RGCs are the output cells of the retina, and are thus an important cell to study in order to understand how light information is signaled to visual nuclei in the brain. Advances in mouse genetics over recent decades have resulted in a variety of fluorescent reporter mouse lines where specific RGC populations are labeled with a fluorescent protein to allow for identification of RGC subtypes 2 3 4 and specific targeting for electrophysiological recording. Here, we present a method for recording light responses from fluorescently labeled ganglion cells in an intact, isolated retinal preparation. This isolated retinal preparation allows for recordings from RGCs where the dendritic arbor is intact and the inputs across the entire RGC dendritic arbor are preserved. This method is applicable across a variety of ganglion cell subtypes and is amenable to a wide variety of single-cell physiological techniques.

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.

The first steps in vertebrate vision take place when light stimulates the rod and cone photoreceptors of the retina 1.  This information is then ...

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

We use the whole-cell patch clamp technique to study the synaptic circuitry that underlies visual information processing in the retina. In this video, we will guide you through the process of performing whole-cell recordings of light evoked currents of individual cells in the retinal slice preparation. We use the aquatic tiger salamander as an animal model. We begin by describing the dissection of the eye and show how slices are mounted for electrophysiological recordings. Once the slice is placed in the recording chamber, we demonstrate how to perform whole-cell voltage clamp recordings. We then project visual stimuli onto the photoreceptors in the slice to elicit light-evoked current responses. During the recording we perfuse the slice with pharmacological agents, whereby an 8-channel perfusion system allows us to quickly switch between different agents.  The retinal slice preparation is widely used for patch clamp recordings in the retina, in particular to study amacrine or bipolar cells, which are not accessible in a whole-mount preparation.

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.

We use the whole-cell patch clamp technique to study the synaptic circuitry that underlies visual information processing in the retina. In this video, we ...

Biology

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

One of the central tasks in retinal neuroscience is to understand the circuitry of retinal neurons and how those connections are responsible for shaping the signals transmitted to the brain. Photons are detected in the retina by rod and cone photoreceptors, which convert that energy into an electrical signal, transmitting it to other retinal neurons, where it is processed and communicated to central targets in the brain via the optic nerve. Important early insights into retinal circuitry and visual processing came from the histological studies of Cajal1,2 and, later, from electrophysiological recordings of the spiking activity of retinal ganglion cells - the output cells of the retina3,4.
A detailed understanding of visual processing in the retina requires an understanding of the signaling at each step in the pathway from photoreceptor to retinal ganglion cell. However, many retinal cell types are buried deep in the tissue and therefore relatively inaccessible for electrophysiological recording. This limitation can be overcome by working with vertical slices, in which cells residing within each of the retinal layers are clearly visible and accessible for electrophysiological recording.
Here, we describe a method for making vertical sections of retinas from larval tiger salamanders (Ambystoma tigrinum). While this preparation was originally developed for recordings with sharp microelectrodes5,6, we describe a method for dual whole-cell voltage clamp recordings from photoreceptors and second-order horizontal and bipolar cells in which we manipulate the photoreceptor's membrane potential while simultaneously recording post-synaptic responses in horizontal or bipolar cells. The photoreceptors of the tiger salamander are considerably larger than those of mammalian species, making this an ideal preparation in which to undertake this technically challenging experimental approach. These experiments are described with an eye toward probing the signaling properties of the synaptic ribbon - a specialized synaptic structure found in a only a handful of neurons, including rod and cone photoreceptors, that is well suited for maintaining a high rate of tonic neurotransmitter release7,8 - and how it contributes to the unique signaling properties of this first retinal synapse.

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.

One of the central tasks in retinal neuroscience  is to understand the circuitry of retinal neurons and how those connections are  responsible for shaping ...

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

The mammalian retina is a layered tissue composed of multiple neuronal types. To understand how visual signals are processed within its intricate synaptic network, electrophysiological recordings are frequently used to study connections among individual neurons. We have optimized a flat-mount preparation for patch clamp recording of genetically marked neurons in both GCL (ganglion cell layer) and INL (inner nuclear layer) of mouse retinas. Recording INL neurons in flat-mounts is favored over slices because both vertical and lateral connections are preserved in the former configuration, allowing retinal circuits with large lateral components to be studied. We have used this procedure to compare responses of mirror-partnered neurons in retinas such as the cholinergic starburst amacrine cells (SACs).

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

The mammalian retina is a layered tissue composed of multiple neuronal types. To understand how visual signals are processed within its intricate synaptic ...

Preparation of Horizontal Slices of Adult Mouse Retina for Electrophysiological Studies

Vertical slice preparations are well established to study circuitry and signal transmission in the adult mammalian retina. The plane of sectioning in these preparations is perpendicular to the retinal surface, making it ideal for the study of radially oriented neurons like photoreceptors and bipolar cells. However, the large dendritic arbors of horizontal cells, wide-field amacrine cells, and ganglion cells are mostly truncated, leaving markedly reduced synaptic activity in these cells. Whereas ganglion cells and displaced amacrine cells can be studied in a whole-mounted preparation of the retina, horizontal cells and amacrine cells located in the inner nuclear layer are only poorly accessible for electrodes in whole retina tissue.
To achieve maximum accessibility and synaptic integrity, we developed a horizontal slice preparation of the mouse retina, and studied signal transmission at the synapse between photoreceptors and horizontal cells. Horizontal sectioning allows&#160;(1) easy and unambiguous visual identification of horizontal cell bodies for electrode targeting, and (2) preservation of the extended horizontal cell dendritic fields, as a prerequisite for intact and functional cone synaptic input to horizontal cell dendrites.
Horizontal cells from horizontal slices exhibited tonic synaptic activity in the dark, and they responded to brief flashes of light with a reduction of inward current and diminished synaptic activity. Immunocytochemical evidence indicates that almost all cones within the dendritic field of a horizontal cell establish synapses with its peripheral dendrites. The horizontal slice preparation is therefore well suited to study the physiological properties of horizontally extended retinal neurons as well as sensory signal transmission and integration across selected synapses.

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.

Vertical slice preparations are well established to study circuitry and signal transmission in the adult mammalian retina. The plane of sectioning in ...

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

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic tectum, is a popular model to study how neural circuits self-assemble. The ability to carry out whole cell patch clamp recordings from tectal neurons and to record RGC-evoked responses, either in vivo or using a whole brain preparation, has generated a large body of high-resolution data about the mechanisms underlying normal, and abnormal, circuit formation and function. Here we describe how to perform the in vivo preparation, the original whole brain preparation, and a more recently developed horizontal brain slice preparation for obtaining whole cell patch clamp recordings from tectal neurons. Each preparation has unique experimental advantages. The in vivo preparation enables the recording of the direct response of tectal neurons to visual stimuli projected onto the eye. The whole brain preparation allows for the RGC axons to be activated in a highly controlled manner, and the horizontal brain slice preparation allows recording from across all layers of the tectum.

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

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.

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic ...

Light-Evoked Postsynaptic Responses in Neurons of Mouse Retinal Slices

Source: Hellmer, C. B., et al. <a href="https://app.jove.com/v/52422">Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques.</a> J. Vis. Exp. (2015).
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.

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1. Preparation of Experimental ...

JoVE Encyclopedia of Experiments

Neurophysiology

Imaging and Optical Techniques

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

Source: Richevaux, L., et al. <a href="https://app.jove.com/v/59534">In Vivo Intracerebral Stereotaxic Injections for Optogenetic Stimulation of Long-Range Inputs in Mouse Brain Slices.</a> J. Vis. Exp. (2019)
The video demonstrates a protocol to study synaptic interactions and neuronal responses in a transfected mouse hippocampal brain slice. It involves illuminating long-range sensory neurons to trigger action potentials and recording the resulting excitatory post-synaptic potentials in the recorded neuron using the whole-cell patch-clamp technique.

All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.
1. Solutions for acute slice ...

Electrophysiology Techniques

Optogenetic Stimulation and Electrophysiological Recording of Synaptic Events in Rat Brain Slices

Source: Kinnavane, L., et al. <a href="https://app.jove.com/v/63077">Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex.</a> J. Vis. Exp. (2022)
This video demonstrates the procedure for studying synaptic efficiency in the lateral entorhinal cortex (LEC) by optogenetically stimulating medial prefrontal cortex (mPFC) axons and recording the responses of pyramidal neurons in the LEC. This method combines patch-clamp recordings with synaptic activation by light stimulation to study synaptic plasticity.

All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.
1. Electrophysiology and ...

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

Summary

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Chapters in this video

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

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

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

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

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

Preparation of Horizontal Slices of Adult Mouse Retina for Electrophysiological Studies

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

Light-Evoked Postsynaptic Responses in Neurons of Mouse Retinal Slices

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

Optogenetic Stimulation and Electrophysiological Recording of Synaptic Events in Rat Brain Slices