This work has been funded by the German Research Foundation (DFG) within the Collaborative Research Center (SFB) 1134 &#34;Functional Ensembles&#34; (J.v.E. and X.C.) and the Research Grant EN948/1-2 (J.v.E.).

All the experiments were approved by the Governmental Supervisory Panel on Animal Experiments of Rhineland-Palatinate.
1. Solutions
<ol>
	<li>Dissection solution
	<ol>
		<li>To reduce excitotoxicity, prepare a choline-based solution to be used during the dissection as presented here (in mM): 87 NaCl, 2.5 KCl, 37.5 choline chloride, 25 NaHCO3, 1.25 NaH2PO4, 0.5 CaCl2, 7 MgCl2, and25 glucose. Prepare the dissection solution less than 1 week before the experiment.</li>
	</ol>
	</li>
	<li>Recording solution
	<ol>
		<li>Prepare an artificial cerebral spinal fluid (ACSF) solution containing (in mM): 125 NaCl, 25 NaHCO3, 1.25 NaH2PO4, 2.5 KCl, 2 CaCl2, 1 MgCl2, and 25 glucose. Add 50 &#181;M D-APV and 10 &#181;M SR 95531 hydrobromide to block N-methyl-D-aspartate (NMDA) and GABAA receptors, respectively.</li>
	</ol>
	</li>
	<li>Intracellular solution
	<ol>
		<li>Prepare an intracellular solution containing (in mM): 35 Cs-gluconate, 100 CsCl, 10 HEPES, 10 EGTA, and 0.1 D-600 (methoxyverapamil hydrochloride). Adjust the pH to 7.3 with CsOH. Filter, aliquot, and store the stock solution at -20 &#176;C until use.</li>
	</ol>
	</li>
</ol>
2. Dissection
<ol>
	<li>Prepare two slice chambers, of which one is filled with 100 mL of the dissection solution and the other with 100 mL of recording solution. Place the two beakers into a water bath (37 &#176;C) and bubble the solutions with carbogen for at least 15 min before the dissection.</li>
	<li>Fill a plastic beaker with 250 mL of near ice-cold (but not frozen) dissection solution. Bubble with carbogen at least 15 min before use.</li>
	<li>Fill a plastic Petri dish (100 mm x 20 mm), in which the brain dissection will be performed, with the cold dissection solution. Place a piece of filter paper into the lid of the Petri dish.</li>
	<li>Cool down the dissection chamber of the vibratome.</li>
	<li>Anesthetize a mouse with 2.5% isoflurane, e.g., in the mouse cage. As soon as the mouse does not respond to a tail pinch, decapitate the mouse near the cervical medulla and immerse the head into the ice-cold dissection solution in the base of the Petri dish.</li>
	<li>Cut the skin of the head from caudal to nasal and keep it laterally with fingers to expose the skull. To take out the forebrain, first cut the skull along with the posterior-anterior midline, and then cut two more times through the coronal suture and lambdoid suture (Figure 1A).</li>
	<li>Remove the skull between the coronal suture and lambdoid suture such that the cerebrum is exposed. Cut the olfactory bulb and separate the forebrain from the midbrain with a fine blade (Figure 1B). Remove the brain from the skull with a thin spatula and place it onto the filter paper in the lid of the Petri dish. 
	NOTE: Steps 2.6 and 2.7 should be performed as fast as possible to reduce cell death.</li>
	<li>To preserve the integrity of the sensory and cortical inputs to the dLGN in the brain slice, separate the two hemispheres with a parasagittal cut with an angle of 3&#8722;5&#176; (Figure 1C,D). Dry the mediolateral planes of the two hemispheres by placing them on the filter paper and then glue them onto the cutting stage with an angle of 10&#8722;25&#176; from the horizontal plane (Figure 1E).</li>
	<li>Place the stage in the center of the metal buffer tray and gently pour the rest of the ice-cold dissection solution into the buffer tray. Perform this step carefully since a strong pour might remove the pasted brain (Figure 1F).</li>
	<li>Place the buffer tray into the ice-filled tray, which helps to maintain the low temperature during dissection. Cut 250 &#181;m slices with a razor blade at a speed of 0.1 mm/s and an amplitude of 1 mm.</li>
	<li>Keep slices in the holding chamber filled with oxygenated dissection solution at 34 &#176;C for around 30 min and then allow the slices to recover in the recording solution at 34 &#176;C for another 30 min.</li>
	<li>After recovery, remove the slice holding chamber from the water bath and keep slices at room temperature (RT) until used in experiments.</li>
</ol>
3. Electrophysiology
<ol>
	<li>Pull pipettes using borosilicate glass capillaries and a filament puller. Maintain the resistance of recording pipettes at 3&#8722;4 M&#937;. Pull stimulating pipettes using the same protocol but break the tip slightly after pulling to increase the diameter. Fill recording pipettes with the intracellular solution and biocytin, while fill stimulating pipettes with the recording solution.</li>
	<li>Place the slices into the recording chamber and continuously perfuse the slices with oxygenated recording solution at RT. Check all the slices and select the ones that display an intact optic tract.</li>
	<li>Visualize the slice and cells with an upright microscope equipped with infrared differential interference contrast (IR-DIC) video microscopy. 
	NOTE: Relay neurons are differentiated from interneurons by their larger soma size and more primary dendrites (branches emerging from the soma). Interneurons display bipolar morphology with a smaller soma.</li>
	<li>Place the stimulating pipette onto the slice before patching the cell with the recording pipette. To investigate retinogeniculate synapses, place the stimulating pipette directly onto the optic tract, where the axon fibers from retinal ganglion cells are bundled (Figure 2A). To analyze corticogeniculate synapses, place the stimulating electrode on the nucleus reticularis thalami, which is rostroventrally adjacent to the dLGN (Figure 2B).</li>
	<li>Once the recording pipette is immersed into the recording solution, apply a 5 mV voltage step to monitor the pipette resistance. Set the holding potential to 0 mV and cancel the offset potential so that the holding current is 0 pA.</li>
	<li>Approach the cell with the recording pipette while applying positive pressure. When the pipette is in direct contact to the cell membrane, release the positive pressure, and set the holding potential to -70 mV. Apply a slight negative pressure to allow the cell membrane to attach to the glass pipette such that a gigaohm seal forms (resistance &#62;1 G&#937;). Compensate the pipette capacitance and open the cell by application of negative pressure pulses.</li>
	<li>To investigate the synaptic function, apply 0.1 ms current pulses (ca. 30 &#181;A) via the stimulation pipette. If no synaptic responses are observed, the stimulating pipettes may have to be replaced. Be careful during placing the stimulating pipettes to a different position such that the recorded cell is not lost. 
	NOTE: In the example recordings shown in Figure 2, synaptic short-term plasticity was investigated by stimulating the optic tract or corticogeniculate fibers twice with 30 ms inter-stimulus intervals. The paired-pulse ratio (PPR) was calculated by dividing the amplitude of the second excitatory postsynaptic current (EPSC) by that of the first EPSC (EPSC2/EPSC1). Each stimulation protocol was repeated 20 times. The EPSC amplitude was measured from average currents of the 20 sweeps. The time interval between each repetition was least 5 s to avoid short-term plasticity induced by the repetitions.</li>
	<li>Monitor the series resistance continuously by applying a 5-mV voltage step. Only use the cells with the series resistance smaller than 20 M&#937; for analysis.</li>
</ol>
4. Biocytin labeling
<ol>
	<li>Maintain the whole-cell configuration for at least 5 min to allow the diffusion of biocytin into the distal dendrites.</li>
	<li>After recording, gently remove the pipette from the cell so that the soma is not destroyed. 
	NOTE: If more than one cell is recorded on one slice, their somas should be sufficiently away from their neighboring cells. Ensure that the dendrites from different cells do not interfere with each other during imaging.</li>
	<li>Prepare a 24-well cell culture plate. Fill each well with 300 &#181;L of 4% paraformaldehyde (PFA). Take care to not contaminate anything with PFA that will be in contact with the slices to be recorded.</li>
	<li>Transfer the slices from the recording chamber to the PFA-containing plate with a rubber pipette and fix the slices overnight. Ensure that the pipette is always prevented from PFA contamination. 
	CAUTION: Always wear gloves and use a chemical hood when manipulating PFA.</li>
	<li>After fixing slices overnight in PFA, replace PFA with phosphate-buffered saline (PBS). Store the slices in PBS at 4 &#176;C for future processing (less than 2 weeks).</li>
	<li>Wash the slices with fresh PBS 3 times (10 min each wash).</li>
	<li>Block nonspecific binding sites by incubating the slices in the blocking solution (0.2% non-ionic surfactant and 5% bovine serum albumin in PBS) for 2 h at RT on an orbital shaker.</li>
	<li>Discard the blocking solution. Incubate the slices with streptavidin-Alexa 568 diluted into the blocking solution (1:1,000) at 4 &#176;C overnight on an orbital shaker.</li>
	<li>Discard the antibody solution and wash slices 3 times with PBS for 10 min at RT on an orbital shaker. Wash the slices with tap water before mounting.</li>
	<li>Put the slices from one mouse on one glass slide. Ensure that the stained cells are present on the top surface of the slices. Absorb the water around the slices with tissues and then leave the slices to dry for approximately 15 min.</li>
	<li>Apply two drops of mounting medium on each slice. Mount a coverslip on the slide without producing air bubbles.</li>
	<li>Store the labeled slices at 4 &#176;C after visualization with a confocal microscope.</li>
</ol>
5. Cellular imaging and reconstruction
<ol>
	<li>Scan slices with a confocal microscope and use the associated software for analysis.</li>
	<li>Excite the fluorescence at 561 nm.</li>
	<li>Image the slices with 0.7 numerical aperture (NA), oil-immersion objective.</li>
	<li>Adjust the targeted cell in the middle of the view and apply a 2.5-times magnification.</li>
	<li>Render the Z-stack datasets to scan the whole neuron with the following parameters: format = 2,550 x 2,550 pixels, scan frequency = 400 Hz, z number = 40.Voxel size was around 0.1211 x 0.1211 x 1.8463 &#181;m3.</li>
	<li>Semi-automatically trace the neurons using a neuronal reconstruction software (e.g., NeuronStudio). An example of a 3D traced relay neuron is shown in Figure 3B.</li>
</ol>

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The authors have nothing to disclose.

We describe an improved protocol based on a previously published method3, which allows for the investigation of the high probability of release retinogeniculate synapses and low probability of release corticogeniculate synapses from the same slice. This is of great importance since these two inputs interact with each other to modulate the visual signal transmission: retinogeniculate inputs are the main excitatory drive of relay neurons, whereas corticothalamic inputs function as a modulator, which...

Relay neurons of the lateral geniculate nucleus integrate and relay visual information to the visual cortex. These neurons receive excitatory input from ganglion cells via retinogeniculate synapses, which provide the main excitatory drive for relay neurons. In addition, relay neurons receive excitatory inputs from cortical neurons via corticogeniculate synapses. Moreover, relay neurons receive inhibitory inputs from local interneurons and GABAergic neurons of the nucleus reticularis thalami1. The nucleus reticularis thalami is present like a shield between thalamus and cortex such that fibers projecting from cortex to thalamus and in the opposite direction must go through the nucleus reticularis thalami2.
Retinogeniculate inputs and corticogeniculate inputs display distinct synaptic properties3,4,5,6,7,8. Retinogeniculate inputs form large terminals with multiple release sites9,10. In contrast, corticogeniculate inputs display small terminals with single release sites7. In addition, retinogeniculate synapses efficiently drive action potentials of relay neurons despite constituting only 5&#8722;10% of all synapses on relay neurons3,8,11. Corticogeniculate synapses, on the other hand, serve as a modulator of retinogeniculate transmissions by controlling the membrane potential of relay neurons12,13.
These two main excitatory inputs to relay neurons are also functionally different. One prominent difference is the short-term depression of retinogeniculate synapses and the short-term facilitation of corticogeniculate synapses3,5,8. Short-term plasticity refers to a phenomenon in which synaptic strength changes when the synapse is repeatedly active within a time period of few milliseconds to several seconds. Synaptic release probability is an important factor underlying short-term plasticity. Synapses, with a low initial release probability, display short-term facilitation due to the buildup of Ca2+ in the presynapse and consequently an increase in the release probability is observed upon repeated activity. In contrast, synapses with high release probability usually display short-term depression due to the depletion of ready-releasable vesicles14. In addition, desensitization of postsynaptic receptors contributes to the short-term plasticity in some high-release probability synapses8,15. High release probability and desensitization of &#945;-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptors contribute to the prominent short-term depression of retinogeniculate synapses. In contrast, low-release probability underlies the short-term facilitation of corticogeniculate synapses.
In mice, the optic tract enters the dorsal lateral geniculate nucleus (dLGN) from the caudolateral site, whereas corticogeniculate fibers enter the dLGN rostroventrally. The distance between the two inputs allows for the investigation of the individual properties of two very different excitatory inputs impinging onto the same cell. Here, we build on and improve a previously described dissection method in which retinogeniculate and corticogeniculate fibers are preserved in acute brain slices3. We, then, describe the electrophysiological investigation of relay neurons and stimulation of retinogeniculate and corticogeniculate fibers with extracellular stimulation electrodes. Finally, we provide a protocol for the filling of relay neurons with biocytin and subsequent anatomical analysis.

<table><tbody><tr><td>Amplifier&amp;nbsp;</td><td>HEKA Elektronik</td><td>EPC 10 USB Double patch clamp amplifier</td><td></td></tr><tr><td>Biocytin</td><td>Sigma-Aldrich</td><td>B4261-250MG</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>EMSURE</td><td>1.02382.1000</td><td></td></tr><tr><td>choline chloride</td><td>Sigma-Aldrich</td><td>C1879-1KG</td><td></td></tr><tr><td>Confocal Laser Scanning Microscope</td><td>Leica Microsystems</td><td>TCS SP5</td><td></td></tr><tr><td>CsCl</td><td>EMSURE</td><td>1.02038.0100</td><td></td></tr><tr><td>Cs-gluconate</td><td>Self-prepared</td><td></td><td>Since there was no commercial Cs-gluconate, we prepared it by ourselves&amp;nbsp;</td></tr><tr><td>D-600&amp;nbsp;</td><td>Sigma-Aldrich</td><td>M5644-50MG</td><td>methoxyverapamil hydrochloride</td></tr><tr><td>D-APV&amp;nbsp;</td><td>Biotrend&amp;nbsp;</td><td>BN0085-100</td><td>NMDA-receptor antagonist</td></tr><tr><td>Digital camera for microscope</td><td>Olympus</td><td>XM10</td><td></td></tr><tr><td>EGTA</td><td>SERVA</td><td>11290.02</td><td></td></tr><tr><td>Forene</td><td>Abbvie</td><td>2594.00.00</td><td>isoflurane</td></tr><tr><td>Glucose</td><td>Sigma-Aldrich</td><td>49159-1KG</td><td></td></tr><tr><td>HEPES</td><td>ROTH</td><td>9105.2</td><td></td></tr><tr><td>High Current Stimulus Isolator</td><td>World Precision Instruments</td><td>A385</td><td></td></tr><tr><td>KCl</td><td>EMSURE</td><td>1.04936.1000</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>EMSURE</td><td>1.05833.0250</td><td></td></tr><tr><td>Micromanipulators</td><td>Luigs &amp;amp; Neumann</td><td>SM7</td><td></td></tr><tr><td>Miroscope</td><td>Olympus</td><td>BX51</td><td></td></tr><tr><td>mounting medium&amp;nbsp;</td><td>ThermoFisher Scientific</td><td>P36930</td><td>Prolong Gold Invitrogen</td></tr><tr><td>NaCl</td><td>ROTH</td><td>3957.1</td><td></td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>EMSURE</td><td>1.06346.1000</td><td></td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>EMSURE</td><td>1.06329.1000</td><td></td></tr><tr><td>Pipette</td><td>Hilgenberg</td><td>1807502</td><td></td></tr><tr><td>Puller</td><td>Sutter&amp;nbsp;</td><td>P-1000</td><td></td></tr><tr><td>razor blade&amp;nbsp;</td><td>Personna&amp;nbsp;</td><td>60-0138</td><td></td></tr><tr><td>Semiautomatic Vibratome</td><td>Leica&amp;nbsp; Biosystems</td><td>VT1200S</td><td></td></tr><tr><td>SR 95531 hydrobromide&amp;nbsp;</td><td>Biotrend&amp;nbsp;</td><td>AOB5680-10</td><td>GABAA-receptor antagonist&amp;nbsp;</td></tr></tbody></table>

electrophysiological investigations of retinogeniculate and corticogeniculate synapse function

The lateral geniculate nucleus is the first relay station for the visual information. Relay neurons of this thalamic nucleus integrate input from retinal ganglion cells and project it to the visual cortex. In addition, relay neurons receive top-down excitation from the cortex. The two main excitatory inputs to the relay neurons differ in several aspects. Each relay neuron receives input from only a few retinogeniculate synapses, which are large terminals with many release sites. This is reflected by the comparably strong excitation, the relay neurons receive, from retinal ganglion cells. Corticogeniculate synapses, in contrast, are simpler with few release sites and weaker synaptic strength. The two synapses also differ in their synaptic short-term plasticity. Retinogeniculate synapses have a high release probability and consequently display a short-term depression. In contrast, corticogeniculate synapses have a low release probability. Corticogeniculate fibers traverse the reticular thalamic nuclei before entering the lateral geniculate nucleus. The different locations of the reticular thalamic nucleus (rostrally from the lateral geniculate nucleus) and optic tract (ventro-laterally from the lateral geniculate nucleus) allow stimulating corticogeniculate or retinogeniculate synapses separately with extracellular stimulation electrodes. This makes the lateral geniculate nucleus an ideal brain area where two excitatory synapses with very different properties impinging onto the same cell type, can be studied simultaneously. Here, we describe a method to investigate the recording from relay neurons and to perform detailed analysis of the retinogeniculate and corticogeniculate synapse function in acute brain slices. The article contains a step-by-step protocol for the generation of acute brain slices of the lateral geniculate nucleus and steps for recording activity from relay neurons by stimulating the optic tract and corticogeniculate fibers separately.

The slice preparation of dLGN containing the retinogeniculate and corticogeniculate pathways is shown under a 4x objective (Figure 2). Axons of retinal ganglion cells bundle together in the optic tract (Figure 2). The stimulating pipette was placed on the optic tract to induce retinogeniculate synapse-mediated current (Figure 2A) and on nucleus reticularis thalami to induce corticogeniculate synapses...

Watch this Scientific Journal Video about Electrophysiological Investigations of Retinogeniculate and Corticogeniculate Synapse Function at JoVE.com

Electrophysiological Investigations of Retinogeniculate and Corticogeniculate Synapse Function

Here, we present protocols for the preparation of acute brain slices containing the lateral geniculate nucleus and the electrophysiological investigation of retinogeniculate and corticogeniculate synapses function. This protocol provides an efficient way to study synapses with the high-release and low-release probability in the same acute brain slices.

The lateral geniculate nucleus is the first relay station for the visual information. Relay neurons of this thalamic nucleus integrate input from retinal ...

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6.0K Views.  University Medical Center of the Johannes Gutenberg University Mainz. Hier presenteren we de protocollen voor de voorbereiding van acute hersenplakjes die de laterale geniale kern en elektrofysiologisch onderzoek van retinalegeniculate en corticale geniculate synapsfunctie bevatten. Retinale ganglion cel axonen voer de laterale geniale kern ver weg van de corticale ingangen. Dit maakt de afzonderlijke stimulaties van retinalegeniculate en corticale geniale synapsen mogelijk, die zeer verschillende functionele eigenschappen hebben.Om deze procedure te beg...

Video: Electrophysiological Investigations of Retinogeniculate and Corticogeniculate Synapse Function

Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Visual stimuli are detected and conveyed over a wide dynamic range of light intensities and frequency changes by specialized neurons in the vertebrate retina. Two classes of retinal neurons, photoreceptors and bipolar cells, accomplish this by using ribbon-type active zones, which enable sustained and high-throughput neurotransmitter release over long time periods. ON-type mixed bipolar cell (Mb) terminals in the goldfish retina, which depolarize to light stimuli and receive mixed rod and cone photoreceptor input, are suitable for the study of ribbon-type synapses both due to their large size (~10-12 &mu;m diameter) and to their numerous lateral and reciprocal synaptic connections with amacrine cell dendrites. Direct access to Mb bipolar cell terminals in goldfish retinal slices with the patch-clamp technique allows the measurement of presynaptic Ca2+ currents, membrane capacitance changes, and reciprocal synaptic feedback inhibition mediated by GABAA and GABAC receptors expressed on the terminals. Presynaptic membrane capacitance measurements of exocytosis allow one to study the short-term plasticity of excitatory neurotransmitter release 14,15. In addition, short-term and long-term plasticity of inhibitory neurotransmitter release from amacrine cells can also be investigated by recordings of reciprocal feedback inhibition arriving at the Mb terminal 21. Over short periods of time (e.g. ~10 s), GABAergic reciprocal feedback inhibition from amacrine cells undergoes paired-pulse depression via GABA vesicle pool depletion 11. The synaptic dynamics of retinal microcircuits in the inner plexiform layer of the retina can thus be directly studied.
The brain-slice technique was introduced more than 40 years ago but is still very useful for the investigation of the electrical properties of neurons, both at the single cell soma, single dendrite or axon, and microcircuit synaptic level 19. Tissues that are too small to be glued directly onto the slicing chamber are often first embedded in agar (or placed onto a filter paper) and then sliced 20, 23, 18, 9. In this video, we employ the pre-embedding agar technique using goldfish retina. Some of the giant bipolar cell terminals in our slices of goldfish retina are axotomized (axon-cut) during the slicing procedure. This allows us to isolate single presynaptic nerve terminal inputs, because recording from axotomized terminals excludes the signals from the soma-dendritic compartment. Alternatively, one can also record from intact Mb bipolar cells, by recording from terminals attached to axons that have not been cut during the slicing procedure. Overall, use of this experimental protocol will aid in studies of retinal synaptic physiology, microcircuit functional analysis, and synaptic transmission at ribbon synapses.

Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Here we describe a protocol for the preparation of agar-embedded retinal slices that are suitable for electrophysiology and Ca2+ imaging. This method allows one to study ribbon-type synapses in retinal microcircuits using direct patch-clamp recordings of single presynaptic nerve terminals.

Visual stimuli are detected and conveyed over a wide dynamic range of light intensities and frequency changes by specialized neurons in the vertebrate ...

Implementing Dynamic Clamp with Synaptic and Artificial Conductances in Mouse Retinal Ganglion Cells

Ganglion cells are the output neurons of the retina and their activity reflects the integration of multiple synaptic inputs arising from specific neural circuits. Patch clamp techniques, in voltage clamp and current clamp configurations, are commonly used to study the physiological properties of neurons and to characterize their synaptic inputs. Although the application of these techniques is highly informative, they pose various limitations. For example, it is difficult to quantify how the precise interactions of excitatory and inhibitory inputs determine response output. To address this issue, we used a modified current clamp technique, dynamic clamp, also called conductance clamp 1, 2, 3 and examined the impact of excitatory and inhibitory synaptic inputs on neuronal excitability. This technique requires the injection of current into the cell and is dependent on the real-time feedback of its membrane potential at that time. The injected current is calculated from predetermined excitatory and inhibitory synaptic conductances, their reversal potentials and the cell's instantaneous membrane potential. Details on the experimental procedures, patch clamping cells to achieve a whole-cell configuration and employment of the dynamic clamp technique are illustrated in this video article. Here, we show the responses of mouse retinal ganglion cells to various conductance waveforms obtained from physiological experiments in control conditions or in the presence of drugs. Furthermore, we show the use of artificial excitatory and inhibitory conductances generated using alpha functions to investigate the responses of the cells.

This video article illustrates the set-up, the procedures to patch cell bodies and how to implement dynamic clamp recordings from ganglion cells in whole-mount mouse retinae. This technique allows the investigation of the precise contribution of excitatory and inhibitory synaptic inputs, and their relative magnitude and timing to neuronal spiking.

Ganglion cells are the output neurons of the  retina and their activity reflects the integration of multiple synaptic inputs  arising from specific neural ...

Electrophysiological and Morphological Characterization of Neuronal Microcircuits in Acute Brain Slices Using Paired Patch-Clamp Recordings

The combination of patch clamp recordings from two (or more) synaptically coupled neurons (paired recordings) in acute brain slice preparations with simultaneous intracellular biocytin filling allows a correlated analysis of their structural and functional properties. With this method it is possible to identify and characterize both pre- and postsynaptic neurons by their morphology and electrophysiological response pattern. Paired recordings allow studying the connectivity patterns between these neurons as well as the properties of both chemical and electrical synaptic transmission. Here, we give a step-by-step description of the procedures required to obtain reliable paired recordings together with an optimal recovery of the neuron morphology. We will describe how pairs of neurons connected via chemical synapses or gap junctions are identified in brain slice preparations. We will outline how neurons are reconstructed to obtain their 3D morphology of the dendritic and axonal domain and how synaptic contacts are identified and localized. We will also discuss the caveats and limitations of the paired recording technique, in particular those associated with dendritic and axonal truncations during the preparation of brain slices because these strongly affect connectivity estimates. However, because of the versatility of the paired recording approach it will remain a valuable tool in characterizing different aspects of synaptic transmission at identified neuronal microcircuits in the brain.

Patch-clamp recordings and simultaneous intracellular biocytin filling of synaptically coupled neurons in acute brain slices allow a correlated analysis of their structural and functional properties. The aim of this protocol is to describe the essential technical steps of electrophysiological recording from neuronal microcircuits and their subsequent morphological analysis.

The combination of patch clamp recordings from two (or more) synaptically coupled neurons (paired recordings) in acute brain slice preparations with ...

Simultaneous Recording of Electroretinography and Visual Evoked Potentials in Anesthetized Rats

The electroretinogram (ERG) and visual evoked potential (VEP) are commonly used to assess the integrity of the visual pathway. The ERG measures the electrical responses of the retina to light stimulation, while the VEP measures the corresponding functional integrity of the visual pathways from the retina to the primary visual cortex following the same light event. The ERG waveform can be broken down into components that reflect responses from different retinal neuronal and glial cell classes. The early components of the VEP waveform represent the integrity of the optic nerve and higher cortical centers. These recordings can be conducted in isolation or together, depending on the application. The methodology described in this paper allows simultaneous assessment of retinal and cortical visual evoked electrophysiology from both eyes and both hemispheres. This is a useful way to more comprehensively assess retinal function and the upstream effects that changes in retinal function can have on visual evoked cortical function.

This protocol describes simultaneous measurement of electroretinogram and visual evoked potentials in anesthetized rats.

The electroretinogram (ERG) and visual evoked potential (VEP) are commonly used to assess the integrity of the visual pathway. The ERG measures the ...

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 ...

Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex

Studying the physiological properties of specific synapses in the brain, and how they undergo plastic changes, is a key challenge in modern neuroscience. Traditional in vitro electrophysiological techniques use electrical stimulation to evoke synaptic transmission. A major drawback of this method is its nonspecific nature; all axons in the region of the stimulating electrode will be activated, making it difficult to attribute an effect to a particular afferent connection. This issue can be overcome by replacing electrical stimulation with optogenetic-based stimulation. We describe a method for combining optogenetics with in vitro patch-clamp recordings. This is a powerful tool for the study of both basal synaptic transmission and synaptic plasticity of precise anatomically defined synaptic connections and is applicable to almost any pathway in the brain. Here, we describe the preparation and handling of a viral vector encoding channelrhodopsin protein for surgical injection into a pre-synaptic region of interest (medial prefrontal cortex) in the rodent brain and making of acute slices of downstream target regions (lateral entorhinal cortex). A detailed procedure for combining patch-clamp recordings with synaptic activation by light stimulation to study short- and long-term synaptic plasticity is also presented. We discuss examples of experiments that achieve pathway- and cell-specificity by combining optogenetics and Cre-dependent cell labeling. Finally, histological confirmation of the pre-synaptic region of interest is described along with biocytin labeling of the post-synaptic cell, to allow further identification of the precise location and cell type.

Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex

Here we present a protocol describing viral transduction of discrete brain regions with optogenetic constructs to permit synapse-specific electrophysiological characterization in acute rodent brain slices.

Studying the physiological properties of specific synapses in the brain, and how they undergo plastic changes, is a key challenge in modern neuroscience. ...

Studying Bipolar and Horizontal Neuronal Cells in the Murine Split Retina

Split Retina as an Improved Flatmount Preparation for Studying Inner Nuclear Layer Neurons in Vertebrate Retina

Bipolar cells and horizontal cells of the vertebrate retina are the first neurons to process visual information after photons are detected by photoreceptors. They perform fundamental operations such as light adaptation, contrast sensitivity, and spatial and color opponency. A complete understanding of the precise circuitry and biochemical mechanisms that govern their behavior will advance visual neuroscience research and ophthalmological medicine. However, current preparations for examining bipolar and horizontal cells (retinal whole mounts and vertical slices) are limited in their capacity to capture the anatomy and physiology of these cells. In this work, we present a method for removing photoreceptor cell bodies from live, flatmount mouse retinas, providing enhanced access to bipolar and horizontal cells for efficient patch clamping and rapid immunolabeling. Split retinas are prepared by sandwiching an isolated mouse retina between two pieces of nitrocellulose, then gently peeling them apart. The separation splits the retina just above the outer plexiform layer to yield two pieces of nitrocellulose, one containing the photoreceptor cell bodies and another containing the remaining inner retina. Unlike vertical retina slices, the split retina preparation does not sever the dendritic processes of inner retinal neurons, allowing for recordings from bipolar and horizontal cells that integrate the contributions of gap junction-coupled networks and wide-field amacrine cells. This work demonstrates the versatility of this preparation for the study of horizontal and bipolar cells in electrophysiology, immunohistochemistry, and in situ hybridization experiments.

Author Spotlight: Using the Split Retina Technique for Enhanced Access and Accelerated Experiments 

This work presents an alternative flatmount retina preparation in which the removal of photoreceptor cell bodies enables faster antibody diffusion and improved patch pipette access to inner retinal neurons for immunohistochemistry, in situ hybridization, and electrophysiology experiments.

Bipolar cells and horizontal cells of the vertebrate retina are the first neurons to process visual information after photons are detected by ...

Synaptic Circuits Analysis and Protein Localization in the Mouse Retina

Investigating Retinal Circuits and Molecular Localization by Pre&#45;Embedding Immunoelectron Microscopy

The retina comprises numerous cells forming diverse neuronal circuits, which constitute the first stage of the visual pathway. Each circuit is characterized by unique features and distinct neurotransmitters, determining its role and functional significance. Given the intricate cell types within its structure, the complexity of neuronal circuits in the retina poses challenges for exploration. To better investigate retinal circuits and cross-talk, such as the link between cone and rod pathways, and precise molecular localization (neurotransmitters or neuropeptides), such as the presence of substance P-like immunoreactivity in the mouse retina, we employed a pre-embedding immunoelectron microscopy (immuno-EM) method to explore synaptic connections and organization. This approach enables us to pinpoint specific intercellular synaptic connections and precise molecular localization and could play a guiding role in exploring its function. This article describes the protocol, reagents used, and detailed steps, including (1) retina fixation preparation, (2) pre-embedding immunostaining, and (3) post-fixation and embedding.

Author Spotlight: Understanding the Ultrastructural Basis of Retinal Synaptic Connectivity and Neurotransmitter Localization in Mice 

This protocol outlines the detailed steps of pre-embedding immunoelectron microscopy, with a focus on exploring synaptic circuits and protein localization in the retina.

The retina comprises numerous cells forming diverse neuronal circuits, which constitute the first stage of the visual pathway. Each circuit is ...

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 ...

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Neurophysiology

Imaging and Optical Techniques

An Electrophysiological Study of Retinogeniculate and Corticogeniculate Synapses in Mouse Brain Slices

Source: Chen, X., et al. <a href="https://app.jove.com/v/59680">Electrophysiological Investigations of Retinogeniculate and Corticogeniculate Synapse Function.</a> J. Vis. Exp. (2019).
The video describes the electrophysiological recording of retinogeniculate and corticogeniculate synapses in mouse brain slices. The slices containing the dorsal lateral geniculate nucleus (dLGN) are placed in a recording chamber. A stimulating pipette is positioned on either the optic tract to stimulate retinal ganglion cells (RGCs) and investigate retinogeniculate synapses or the nucleus reticularis thalami to stimulate cortical neurons and investigate corticogeniculate synapses. Upon stimulation, the RGCs and cortical neurons release excitatory neurotransmitters, which produce synaptic currents in the LGN relay neurons. These currents are recorded using a patch-clamp recording pipette.

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
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Electrophysiological Investigations of Retinogeniculate and Corticogeniculate Synapse Function

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