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.

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		<tr height="62">
			<td height="62" style="height:62px;width:223px;">Amplifier&#160;</td>
			<td style="width:98px;">HEKA Elektronik</td>
			<td style="width:123px;">EPC 10 USB Double patch clamp amplifier</td>
			<td style="width:172px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:223px;">Biocytin</td>
			<td>Sigma-Aldrich</td>
			<td style="width:123px;">B4261-250MG</td>
			<td></td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;">CaCl2</td>
			<td style="width:98px;">EMSURE</td>
			<td style="width:123px;">1.02382.1000</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">choline chloride</td>
			<td>Sigma-Aldrich</td>
			<td style="width:123px;">C1879-1KG</td>
			<td></td>
		</tr>
		<tr height="62">
			<td height="62" style="height:62px;">Confocal Laser Scanning Microscope</td>
			<td style="width:98px;">Leica Microsystems</td>
			<td style="width:123px;">TCS SP5</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">CsCl</td>
			<td style="width:98px;">EMSURE</td>
			<td style="width:123px;">1.02038.0100</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">Cs-gluconate</td>
			<td style="width:98px;">Self-prepared</td>
			<td style="width:123px;"></td>
			<td>Since there was no commercial Cs-gluconate, we prepared it by ourselves&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">D-600&#160;</td>
			<td>Sigma-Aldrich</td>
			<td style="width:123px;">M5644-50MG</td>
			<td>methoxyverapamil hydrochloride</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:223px;">D-APV&#160;</td>
			<td style="width:98px;">Biotrend&#160;</td>
			<td style="width:123px;">BN0085-100</td>
			<td style="width:172px;">NMDA-receptor antagonist</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:223px;">Digital camera for microscope</td>
			<td style="width:98px;">Olympus</td>
			<td style="width:123px;">XM10</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">EGTA</td>
			<td style="width:98px;">SERVA</td>
			<td style="width:123px;">11290.02</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:223px;">Forene</td>
			<td style="width:98px;">Abbvie</td>
			<td style="width:123px;">2594.00.00</td>
			<td style="width:172px;">isoflurane</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Glucose</td>
			<td>Sigma-Aldrich</td>
			<td style="width:123px;">49159-1KG</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">HEPES</td>
			<td style="width:98px;">ROTH</td>
			<td style="width:123px;">9105.2</td>
			<td></td>
		</tr>
		<tr height="62">
			<td height="62" style="height:62px;width:223px;">High Current Stimulus Isolator</td>
			<td style="width:98px;">World Precision Instruments</td>
			<td style="width:123px;">A385</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">KCl</td>
			<td style="width:98px;">EMSURE</td>
			<td style="width:123px;">1.04936.1000</td>
			<td></td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;">MgCl2</td>
			<td style="width:98px;">EMSURE</td>
			<td style="width:123px;">1.05833.0250</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:223px;">Micromanipulators</td>
			<td style="width:98px;">Luigs &#38; Neumann</td>
			<td style="width:123px;">SM7</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:223px;">Miroscope</td>
			<td style="width:98px;">Olympus</td>
			<td style="width:123px;">BX51</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">mounting medium&#160;</td>
			<td style="width:98px;">ThermoFisher Scientific</td>
			<td>P36930</td>
			<td>Prolong Gold Invitrogen</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">NaCl</td>
			<td style="width:98px;">ROTH</td>
			<td align="right" style="width:123px;">3957.1</td>
			<td></td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;">NaH2PO4</td>
			<td style="width:98px;">EMSURE</td>
			<td style="width:123px;">1.06346.1000</td>
			<td></td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;">NaHCO3</td>
			<td style="width:98px;">EMSURE</td>
			<td style="width:123px;">1.06329.1000</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Pipette</td>
			<td style="width:98px;">Hilgenberg</td>
			<td style="width:123px;">1807502</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:223px;">Puller</td>
			<td style="width:98px;">Sutter&#160;</td>
			<td style="width:123px;">P-1000</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:223px;">razor blade&#160;</td>
			<td style="width:98px;">Personna&#160;</td>
			<td>60-0138</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:223px;">Semiautomatic Vibratome</td>
			<td style="width:98px;">Leica&#160; Biosystems</td>
			<td style="width:123px;">VT1200S</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:223px;">SR 95531 hydrobromide&#160;</td>
			<td style="width:98px;">Biotrend&#160;</td>
			<td style="width:123px;">AOB5680-10</td>
			<td style="width:172px;">GABAA-receptor antagonist&#160;</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 ...

electrophysiological-investigations-retinogeniculate

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

6.0K Views.  University Medical Center of the Johannes Gutenberg University Mainz. 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.

Video: Electrophysiological Investigations of Retinogeniculate and Corticogeniculate Synapse Function

Studying Synaptic Vesicle Pools using Photoconversion of Styryl Dyes

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The vesicles are then retrieved from the plasma membrane (endocytosis) and grouped together with the general pool of vesicles within the nerve terminal, until they undergo a new exo- and endocytosis cycle (vesicle recycling). These processes have been studied using a variety of techniques such as electron microscopy, electrophysiology recordings, amperometry and capacitance measurements. Importantly, during the last two decades a number of fluorescently labeled markers emerged, allowing optical techniques to track vesicles in their recycling dynamics. One of the most commonly used markers is the styryl or FM dye 1; structurally, all FM dyes contain a hydrophilic head and a lipophilic tail connected through an aromatic ring and one or more double bonds (Fig. 1B). A classical FM dye experiment to label a pool of vesicles consists in bathing the preparation (Fig. 1Ai) with the dye during the stimulation of the nerve (electrically or with high K+). This induces vesicle recycling and the subsequent loading of the dye into recently endocytosed vesicles (Fig. 1Ai-iii). After loading the vesicles with dye, a second round of stimulation in a dye-free bath would trigger the FM release through exocytosis (Fig. 1Aiv-v), process that can be followed by monitoring the fluorescence intensity decrease (destaining).

Although FM dyes have contributed greatly to the field of vesicle recycling, it is not possible to determine the exact localization or morphology of individual vesicles by using conventional fluorescence microscopy. For that reason, we explain here how FM dyes can also be used as endocytic markers using electron microscopy, through photoconversion. The photoconversion technique exploits the property of fluorescent dyes to generate reactive oxygen species under intense illumination. Fluorescently labeled preparations are submerged in a solution containing diaminobenzidine (DAB) and illuminated. Reactive species generated by the dye molecules oxidize the DAB, which forms a stable, insoluble precipitate that has a dark appearance and can be easily distinguished in electron microscopy 2,3. As DAB is only oxidized in the immediate vicinity of fluorescent molecules (as the reactive oxygen species are short-lived), the technique ensures that only fluorescently labeled structures are going to contain the electron-dense precipitate. The technique thus allows the study of the exact location and morphology of actively recycling organelles.

FM dyes have been of invaluable help in the understanding of synaptic dynamics. FMs are normally followed under the fluorescent microscope during different stimulation conditions. However, photoconversion of FM dyes combined with electron microscopy allows the visualization of distinct synaptic vesicle pools, among other ultrastructure components, in synaptic boutons.

The fusion of synaptic vesicles with the plasma membrane (exocytosis) is a required step in neurotransmitter release and neuronal communication. The ...

Electrophysiological Measurements and Analysis of Nociception in Human Infants

Pain is an unpleasant sensory and emotional experience. Since infants cannot verbally report their experiences, current methods of pain assessment are based on behavioural and physiological body reactions, such as crying, body movements or changes in facial expression. While these measures demonstrate that infants mount a response following noxious stimulation, they are limited: they are based on activation of subcortical somatic and autonomic motor pathways that may not be reliably linked to central sensory processing in the brain. Knowledge of how the central nervous system responds to noxious events could provide an insight to how nociceptive information and pain is processed in newborns.
The heel lancing procedure used to extract blood from hospitalised infants offers a unique opportunity to study pain in infancy. In this video we describe how electroencephalography (EEG) and electromyography (EMG) time-locked to this procedure can be used to investigate nociceptive activity in the brain and spinal cord.
This integrative approach to the measurement of infant pain has the potential to pave the way for an effective and sensitive clinical measurement tool.

The assessment and treatment of pain in infants is difficult because infants cannot verbally report their experience. In this video we describe quantitative electrophysiological methods and analysis techniques that can be used to measure the response to noxious events from the infant nervous system.

Pain is an unpleasant sensory and emotional experience. Since infants cannot verbally report their experiences, current methods of pain assessment are ...

Investigations on Alterations of Hippocampal Circuit Function Following Mild Traumatic Brain Injury

Traumatic Brain Injury (TBI) afflicts more than 1.7 million people in the United States each year and even mild TBI can lead to persistent neurological impairments 1. Two pervasive and disabling symptoms experienced by TBI survivors, memory deficits and a reduction in seizure threshold, are thought to be mediated by TBI-induced hippocampal dysfunction 2,3. In order to demonstrate how altered hippocampal circuit function adversely affects behavior after TBI in mice, we employ lateral fluid percussion injury, a commonly used animal model of TBI that recreates many features of human TBI including neuronal cell loss, gliosis, and ionic perturbation 4-6.
	Here we demonstrate a combinatorial method for investigating TBI-induced hippocampal dysfunction. Our approach incorporates multiple ex vivo physiological techniques together with animal behavior and biochemical analysis, in order to analyze post-TBI changes in the hippocampus. We begin with the experimental injury paradigm along with behavioral analysis to assess cognitive disability following TBI. Next, we feature three distinct ex vivo recording techniques: extracellular field potential recording, visualized whole-cell patch-clamping, and voltage sensitive dye recording. Finally, we demonstrate a method for regionally dissecting subregions of the hippocampus that can be useful for detailed analysis of neurochemical and metabolic alterations post-TBI.
	These methods have been used to examine the alterations in hippocampal circuitry following TBI and to probe the opposing changes in network circuit function that occur in the dentate gyrus and CA1 subregions of the hippocampus (see Figure 1). The ability to analyze the post-TBI changes in each subregion is essential to understanding the underlying mechanisms contributing to TBI-induced behavioral and cognitive deficits. 
	The multi-faceted system outlined here allows investigators to push past characterization of phenomenology induced by a disease state (in this case TBI) and determine the mechanisms responsible for the observed pathology associated with TBI.

A multi-faceted approach to investigating functional changes to hippocampal circuitry is explained. Electrophysiological techniques are described along with the injury protocol, behavioral testing and regional dissection method. The combination of these techniques can be applied in similar fashion for other brain regions and scientific questions.

Traumatic Brain Injury (TBI) afflicts more than 1.7 million people in the United States each year and even mild TBI can lead to persistent neurological ...

Simultaneous Electrophysiological Recording and Calcium Imaging of Suprachiasmatic Nucleus Neurons

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in regulating the intracellular calcium concentration. Changing environmental conditions, such as the light-dark cycle, can modify neuronal and neural network activity and the expression of a family of circadian clock genes within the suprachiasmatic nucleus (SCN), the location of the master circadian clock in the mammalian brain. Excitatory synaptic transmission leads to an increase in the postsynaptic Ca2+ concentration that is believed to activate the signaling pathways that shifts the rhythmic expression of circadian clock genes. Hypothalamic slices containing the SCN were patch clamped using microelectrodes filled with an internal solution containing the calcium indicator bis-fura-2. After a seal was formed between the microelectrode and the SCN neuronal membrane, the membrane was ruptured using gentle suction and the calcium probe diffused into the neuron filling both the soma and dendrites. Quantitative ratiometric measurements of the intracellular calcium concentration were recorded simultaneously with membrane potential or current. Using these methods it is possible to study the role of changes of the intracellular calcium concentration produced by synaptic activity and action potential firing of individual neurons. In this presentation we demonstrate the methods to simultaneously record electrophysiological activity along with intracellular calcium from individual SCN neurons maintained in brain slices.

Procedures are described to perform simultaneous recordings of membrane potential or current and changes of intracellular calcium concentration. Suprachiasmatic nucleus neurons are filled with the calcium indicator bis-fura-2 using a patch clamp electrode in the whole cell patch clamp configuration.

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in ...

Imaging Serotonergic Fibers in the Mouse Spinal Cord Using the CLARITY/CUBIC Technique

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal cord of the majority of these fiber systems have not been thoroughly investigated in any species. Serotonergic fibers, which project to the spinal cord, have been studied in rats and opossums on histological sections and their functional significance has been deduced based on their fiber termination pattern in the spinal cord. With the development of CLARITY and CUBIC techniques, it is possible to investigate this fiber system and its distribution in the spinal cord, which is likely to reveal previously unknown features of serotonergic supraspinal pathways. Here, we provide a detailed protocol for imaging the serotonergic fibers in the mouse spinal cord using the combined CLARITY and CUBIC techniques. The method involves perfusion of a mouse with a hydrogel solution and clarification of the tissue with a combination of clearing reagents. Spinal cord tissue was cleared in just under two weeks, and the subsequent immunofluorescent staining against serotonin was completed in less than ten days. With a multi-photon fluorescent microscope, the tissue was scanned and a 3D image was reconstructed using Osirix software.

Supraspinal projections are important for pain perception and other behaviors, and serotonergic fibers are one of these fiber systems. The present study focused on the application of the combined CLARITY/CUBIC protocol to the mouse spinal cord in order to investigate the termination of these serotonergic fibers.

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal ...

Evaluation of Synapse Density in Hippocampal Rodent Brain Slices

In the brain, synapses are specialized junctions between neurons, determining the strength and spread of neuronal signaling. The number of synapses is tightly regulated during development and neuronal maturation. Importantly, deficits in synapse number can lead to cognitive dysfunction. Therefore, the evaluation of synapse number is an integral part of neurobiology. However, as synapses are small and highly compact in the intact brain, the assessment of absolute number is challenging. This protocol describes a method to easily identify and evaluate synapses in hippocampal rodent slices using immunofluorescence microscopy. It includes a three-step procedure to evaluate synapses in high-quality confocal microscopy images by analyzing the co-localization of pre- and postsynaptic proteins in hippocampal slices. It also explains how the analysis is performed and gives representative examples from both excitatory and inhibitory synapses. This protocol provides a solid foundation for the analysis of synapses and can be applied to any research investigating the structure and function of the brain.

A protocol for accurately identifying and analyzing synapses in hippocampal slices using immunofluorescence is outlined in this article.

In the brain, synapses are specialized junctions between neurons, determining the strength and spread of neuronal signaling. The number of synapses is ...

Visualization of Thalamocortical Axon Branching and Synapse Formation in Organotypic Cocultures

Axon branching and synapse formation are crucial processes for establishing precise neuronal circuits. During development, sensory thalamocortical (TC) axons form branches and synapses in specific layers of the cerebral cortex. Despite the obvious spatial correlation between axon branching and synapse formation, the causal relationship between them is poorly understood. To address this issue, we recently developed a method for simultaneous imaging of branching and synapse formation of individual TC axons in organotypic cocultures.
This protocol describes a method which consists of a combination of an organotypic coculture and electroporation. Organotypic cocultures of the thalamus and cerebral cortex facilitate gene manipulation and observation of axonal processes, preserving characteristic structures such as laminar configuration. Two distinct plasmids encoding DsRed and EGFP-tagged synaptophysin (SYP-EGFP) were co-transfected into a small number of thalamic neurons by an electroporation technique. This method allowed us to visualize individual axonal morphologies of TC neurons and their presynaptic sites simultaneously. The method also enabled long-term observation which revealed the causal relationship between axon branching and synapse formation.

This protocol describes a method for simultaneous imaging of thalamocortical axon branching and synapse formation in organotypic cocultures of the thalamus and cerebral cortex. Individual thalamocortical axons and their presynaptic terminals are visualized by a single cell electroporation technique with DsRed and GFP-tagged synaptophysin.

Axon branching and synapse formation are crucial processes for establishing precise neuronal circuits. During development, sensory thalamocortical (TC) ...

Healthy Brain-pituitary Slices for Electrophysiological Investigations of Pituitary Cells in Teleost Fish

Electrophysiological investigations of pituitary cells have been conducted in numerous vertebrate species, but very few in teleost fish. Among these, the clear majority have been performed on dissociated primary cells. To improve our understanding of how teleost pituitary cells, behave in a more biologically relevant environment, this protocol shows how to prepare viable brain-pituitary slices using the small freshwater fish medaka (Oryzias latipes). Making the brain-pituitary slices, pH and osmolality of all solutions were adjusted to values found in body fluids of freshwater fish living at 25 to 28 &#176;C. Following slice preparation, the protocol demonstrates how to conduct electrophysiological recordings using the perforated whole-cell patch-clamp technique. The patch-clamp technique is a powerful tool with unprecedented temporal resolution and sensitivity, allowing investigation of electrical properties from intact whole cells down to single ion channels. Perforated patch is unique in that it keeps the intracellular environment intact preventing regulatory elements in the cytosol from being diluted by the patch pipette electrode solution. In contrast, when performing traditional whole-cell recordings, it was observed that medaka pituitary cells quickly lose their ability to fire action potentials. Among the various perforation techniques available, this protocol demonstrates how to achieve perforation of the patched membrane using the fungicide Amphotericin B.

The article describes an optimized protocol for making viable brain-pituitary tissue slices, using the teleost fish medaka (Oryzias latipes), followed by electrophysiological recordings of pituitary cells using the patch-clamp technique with the perforated patch configuration.

Electrophysiological investigations of pituitary cells have been conducted in numerous vertebrate species, but very few in teleost fish. Among these, the ...

In Vivo Calcium Imaging of Lateral-line Hair Cells in Larval Zebrafish

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory stimuli that mechanically deflect apical protrusions called hair bundles. Deflection opens mechanotransduction (MET) channels in hair bundles, leading to an influx of cations, including calcium. This cation influx depolarizes the cell and opens voltage-gated calcium channels located basally at the hair-cell presynapse. In mammals, hair cells are encased in bone, and it is challenging to functionally assess these activities in vivo. In contrast, larval zebrafish are transparent and possess an externally located lateral-line organ that contains hair cells. These hair cells are functionally and structurally similar to mammalian hair cells and can be functionally assessed in vivo. This article outlines a technique that utilizes a genetically encoded calcium indicator (GECI), GCaMP6s, to measure stimulus-evoked calcium signals in zebrafish lateral-line hair cells. GCaMP6s can be used, along with confocal imaging, to measure in vivo calcium signals at the apex and base of lateral-line hair cells. These signals provide a real-time, quantifiable readout of both mechanosensation- and presynapse-dependent calcium activities within these hair cells. These calcium signals also provide important functional information regarding how hair cells detect and transmit sensory stimuli. Overall, this technique generates useful data about relative changes in calcium activity in vivo. It is less well-suited for quantification of the absolute magnitude of calcium changes. This in vivo technique is sensitive to motion artifacts. A reasonable amount of practice and skill are required for proper positioning, immobilization, and stimulation of larvae. Ultimately, when properly executed, the protocol outlined in this article provides a powerful way to collect valuable information about the activity of hair-cells in their natural, fully integrated states within a live animal.

The zebrafish is a model system that has many valuable features including optical clarity, rapid external development, and, of particular importance to the field of hearing and balance, externally located sensory hair cells. This article outlines how transgenic zebrafish can be used to assay both hair-cell mechanosensation and presynaptic function in toto.

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory ...

Purification of the Dendritic Filopodia-rich Fraction

Dendritic filopodia are thin and long protrusions based on the actin filament, and they extend and retract as if searching for a target axon. When the dendritic filopodia establish contact with a target axon, they begin maturing into spines, leading to the formation of a synapse. Telencephalin (TLCN) is abundantly localized in dendritic filopodia and is gradually excluded from spines. Overexpression of TLCN in cultured hippocampal neurons induces dendritic filopodia formation. We showed that telencephalin strongly binds to an extracellular matrix molecule, vitronectin. Vitronectin-coated microbeads induced phagocytic cup formation on neuronal dendrites. In the phagocytic cup, TLCN, TLCN-binding proteins such as phosphorylated Ezrin/Radixin/Moesin (phospho-ERM), and F-actin are accumulated, which suggests that components of the phagocytic cup are similar to those of dendritic filopodia. Thus, we developed a method for purifying the phagocytic cup instead of dendritic filopodia. Magnetic polystyrene beads were coated with vitronectin, which is abundantly present in the culture medium of hippocampal neurons and which induces phagocytic cup formation on neuronal dendrites. After 24 h of incubation, the phagocytic cups were mildly solubilized with detergent and collected using a magnet separator. After washing the beads, the binding proteins were eluted and analyzed by silver staining and Western blotting. In the binding fraction, TLCN and actin were abundantly present. In addition, many proteins identified from the fraction were localized to the dendritic filopodia; thus, we named the binding fraction as the dendritic filopodia-rich fraction. This article describes details regarding the purification method for the dendritic filopodia-rich fraction.

In this protocol, we introduce a method for purifying the dendritic filopodia-rich fraction from the phagocytic cup-like protrusion structure on cultured hippocampal neurons by taking advantage of the specific and strong affinity between a dendritic filopodial adhesion molecule, TLCN, and an extracellular matrix molecule, vitronectin.

Dendritic filopodia are thin and long protrusions based on the actin filament, and they extend and retract as if searching for a target axon. When the ...