Name: electrophysiological investigations of retinogeniculate and corticogeniculate synapse function
Uploaded: 2019-08-07T13:00:00
Description: Watch this Scientific Journal Video about Electrophysiological Investigations of Retinogeniculate and Corticogeniculate Synapse Function at JoVE.com

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...

<ol>
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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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	<tbody>
		<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>
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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 ...

Here we present the protocols for the preparation of acute brain slices containing the lateral geniculate nucleus and electrophysiological investigation of retinal geniculate and cortical geniculate synapse function. Retinal ganglion cell axons enter the lateral geniculate nucleus far away from the cortical inputs. This allows the separate stimulations of retinal geniculate and cortical geniculate synapses, which have very different functional properties.To begin this procedure, prepare two slice chambers, one filled with 100 milliliters of the dissection solution and the other with 100 milliliters of recording solution. Place the two beakers into a water bath at 37 degrees Celsius, and bubble the solutions with carbogen for at least 15 minutes before the dissection. Next fill a plastic beaker with 250 milliliters of near-ice-cold dissection solution, and bubble it with carbogen for at least 15 minutes before use.Then fill a plastic Petri dish with the cold dissection solution for brain dissection. Place a piece of filter paper in the lid of the Petri dish, and fill it with a small amount of dissection solution. Subsequently cool down the dissection chamber of the vibratome.Cut the skin of the head from caudal to nasal, and keep it opened with fingers to expose the skull. To obtain the forebrain, first cut the skull along the posterior/anterior midline. Then cut two more times through the coronal suture and lambdoid suture.Remove the skull between the coronal suture and lambdoid suture to expose the cerebrum. Cut the olfactory bulb, and separate the forebrain from the midbrain with a fine blade. Remove the brain from the skull with a thin spatula, and place it on a filter paper in the lid of the Petri dish.To preserve the integrity of the sensory and cortical inputs to the dorsal lateral geniculate nucleus in the brain slice, separate the two hemispheres with a parasagittal cut at an angle of three to five degrees. Afterward dry the medial lateral planes of the two hemispheres by placing them on a filter paper. Then glue them onto the cutting stage with an angle of 10 to 25 degrees from the horizontal plane.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 pouring too hard might remove the pasted brain. Next keep the buffer tray in the ice-filled tray to maintain the low temperature during dissection.Section 250-micrometer slices with a razor blade at a speed of 0.1 millimeter per second and an amplitude of one millimeter. Transfer the slices to the holding chamber filled with oxygenated dissection solution at 34 degrees Celsius for around 30 minutes, and then allow the slices to recover in the recording solution at 34 degrees Celsius for another 30 minutes. After recovery, remove the slice-holding chamber from the water bath, and keep the slices at room temperature until used in experiments.In this procedure, pull the recording pipettes using borosilicate glass capillaries and a filament puller. Next pull the stimulating pipettes using the same protocol, but break the tip slightly after pulling to increase the diameter. Subsequently fill the recording pipettes with intracellular solution and biocytin, and fill the stimulating pipettes with recording solution.Then place the slices in the recording chamber, and continuously perfuse them with oxygenated recording solution at room temperature. Visualize the slices with an upright microscope equipped with IR-DIC video microscopy. Check all the slices, and select the ones that display intact optic tracts.Place the stimulation pipette on the slice before patching the cell with a recording pipette. To investigate the retinal geniculate synapses, place the stimulating pipette directly on the optic tract where the axon fibers from retinal ganglion cells are bundled. To analyze the cortical geniculate synapses, place the stimulating electrode on the nucleus reticularis thalami, which is rostroventrally adjacent to the dorsal lateral geniculate nucleus.Once the recording pipette is immersed in the recording solution, apply a five-millivolt step to monitor the pipette resistance. Set the holding potential to zero millivolt, and cancel the offset potential so that the holding current is zero picoampere. Approach the cell with a recording pipette while applying positive pressure.When the pipette is in direct contact with the cell membrane, release the positive pressure, and set the holding potential to minus 70 millivolts. Then apply slight negative pressure to allow the cell membrane to attach to the glass pipette such that a gigaohm seal forms. Compensate the pipette capacitance, and open the cell by applying negative pressure pulses.To investigate the synaptic function, apply 0.1-millisecond current pulses via the stimulation pipette. Monitor the series resistance continuously by applying a five-millivolt step. The series resistance can be estimated by dividing five millivolts by the peak amplitude of the evoked current.Use only the cells with a series resistance smaller than 20 megaohms for analysis. For biocytin labeling, maintain the whole-cell configuration for at least five minutes to allow the diffusion of biocytin into the distal dendrites. After recording, gently remove the pipette from the cell so that the soma is not destroyed.Prepare a 24-well cell culture plate. Fill each well with 300 microliters of 4%PFA. Take care to not contaminate anything with PFA that will be in contact with the slices to be recorded.Transfer the slices from the recording chamber to the PFA-containing plate with a rubber pipette, and fix them overnight. The next day, replace the PFA with PBS. Keep the slices in PBS at four degrees Celsius for future processing.To stain the cells, wash the slices with fresh PBS three times. Block nonspecific binding sites by incubating the slices in the blocking solution for two hours at room temperature on an orbital shaker. Then discard the blocking solution, and incubate the slices with streptavidin Alexa 568 diluted in the blocking solution at four degrees Celsius overnight on an orbital shaker.The next day, discard the antibody solution, and wash the slices three times with PBS for 10 minutes each at room temperature on an orbital shaker. Then wash the slices with tap water before mounting. Place the slices from one mouse on one glass slide.Ensure that the stained cells are present on the upper surface of the slices. Absorb the water around the slices with tissues, and then leave the slices to dry for approximately 15 minutes. Subsequently apply two drops of mounting medium on each slice, and mount a cover slip on the slide without producing air bubbles.Store the labeled slices at four degrees Celsius after viewing them with a confocal microscope. This IR-DIC image shows the soma of a dorsal lateral geniculate nucleus relay neuron with a tip of the patch pipette. Biocytin staining enables us to gain a view of the recorded neuron.Different from interneurons, which have bipolar morphology, relay neurons have multipolar dendritic arbors containing more than three primary dendrites. Three-dimensional reconstruction of a biocytin-labeled relay neuron was then generated, which shows a radially symmetric dendritic architecture. Preserve the retinal geniculate inputs and the cortical geniculate inputs in the same brain slices are most important in this protocol.Following this procedure, we can, for example, also investigate the inhibitor inputs from the nucleus reticularis thalami onto dorsal lateral geniculate nucleus neurons. Remember to wear gloves when fixing the brain slices with PFA, as PFA is hazardous.

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Research

JoVE Journal

Neuroscience

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 ...

Electrophysiological Recording in the Brain of Intact Adult Zebrafish

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram recordings. This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments, allowing recording of neural activity. Immobilization of the adult requires a mechanism to deliver dissolved oxygen to the gills in lieu of buccal and opercular movement. With our technique, animals are immobilized and perfused with habitat water to fulfill this requirement. A craniotomy is performed under tricaine methanesulfonate (MS-222; tricaine) anesthesia to provide access to the brain. The primary electrode is then positioned within the craniotomy window to record extracellular brain activity. Through the use of a multitube perfusion system, a variety of pharmacological compounds can be administered to the adult fish and any alterations in the neural activity can be observed. The methodology not only allows for observations to be made regarding changes in neurological activity, but it also allows for comparisons to be made between larval and adult zebrafish. This gives researchers the ability to identify the alterations in neurological activity due to the introduction of various compounds at different life stages.

This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments to allow recordings and manipulation of neural activity in an intact animal.

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram ...

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 ...

Anti-RDL and Anti-mGlutR1 Receptors Antibody Testing in Honeybee Brain Sections using CRISPR-Cas9

Cluster Regularly Interspaced Short Palindromic Repeats (CRISPR)/CRISPR-associated protein 9 (Cas9) is a gene editing technique widely used in studies of gene function. We use this method in this study to check for the specificity of antibodies developed against the insect GABAA receptor subunit Resistance to Dieldrin (RDL) and a metabotropic glutamate receptor mGlutR1 (mGluRA). The antibodies were generated in rabbits against the conjugated peptides specific to fruit flies (Drosophila melanogaster) as well to honeybees (Apis mellifera). We used these antibodies in honeybee brain sections to study the distribution of the receptors in honeybee brains. The antibodies were affinity purified against the peptide and tested with immunoblotting and the classical method of preadsorption with peptide conjugates to show that the antibodies are specific to the corresponding peptide conjugates against which they were raised. Here we developed the CRISPR-Cas9 technique to test for the reduction of protein targets in the brain 48 h after CRISPR-Cas9 injection with guide RNAs designed for the corresponding receptor. The CRISPR-Cas9 method can also be used in behavioral analyses in the adult bees when one or multiple genes need to be modified.

Presented here is a protocol to use the CRISPR-Cas9 system for reducing the production of a protein in the adult honeybee brain to test antibody specificity.

Cluster Regularly Interspaced Short Palindromic Repeats (CRISPR)/CRISPR-associated protein 9 (Cas9) is a gene editing technique widely used in studies of ...