Name: tuning in the hippocampal theta band in vitro: methodologies for recording from the isolated rodent septohippocampal circuit
Uploaded: 2017-08-02T13:00:00
Description: Watch this Scientific Journal Video about Tuning in the Hippocampal Theta Band In Vitro: Methodologies for Recording from the Isolated Rodent Septohippocampal Circuit at JoVE.com

This work was supported by the Canadian Institutes of Health Research and Natural Sciences.

All procedures have been performed according to protocols and guidelines approved by the McGill University Animal Care Committee and the Canadian Council on Animal Care.
1. Acute Hippocampus In Vitro Preparation
NOTE: Isolating the intact hippocampal preparation involves three major steps: (1) Preparation of solutions and equipment, (2) Dissection of the hippocampus and (3) Setting up the fast perfusion rate system necessary for generation of intrinsic theta ...

<ol>
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While electrophysiological recordings from acute hippocampal slices constitute a standard in vitro technique, the methods presented here differ substantially from the classic approach. Unlike the thin slice preparations where specific cell layers are visible at the surface and can be examined directly, the intact hippocampal preparations are more akin to in vivo configurations where electrodes are lowered into targeted brain regions while crossing through individual layers. The integrity of the hippocam...

Hippocampal theta oscillations (4 - 12 Hz) are amongst the most predominant forms of rhythmic activity in the mammalian brain and are believed to play key roles in cognitive functions such as processing of spatiotemporal information and formation of episodic memories1,2,3. While several in vivo studies that highlight the relationship of theta-modulated place-cells with spatial navigation and lesion studies, as well as clinical evidence, support the view that hippocampal theta oscillations are involved in memory formation4,5,6, the mechanisms associated with generation of hippocampal theta oscillations are still not fully understood. Early in vivo investigations suggested that theta activity depended mainly on extrinsic oscillators, in particular rhythmic input from afferent brain structures such as the septum and entorhinal cortex7,8,9,10. A role for intrinsic factors - internal connectivity of hippocampal neural networks together with the properties of hippocampal neurons - was also postulated based on in vitro observations11,12,13,14,15,16,17,18. However, apart from a few landmark studies19,20,21, difficulties in developing approaches that could replicate physiologically realistic population activities in simple in vitro slice preparations have, for a long time, delayed more detailed experimental examination of the intrinsic abilities of the hippocampus and related areas to self-generate&#160;theta oscillations.
An important downside of the standard in vitro thin-slice experimental setting is that the 3D cellular and synaptic organization of brain structures is usually compromised. This means that many forms of concerted network activities based on spatially distributed cell assemblies, ranging from localized groups (&#8804;1 mm radius) to populations of neurons spread across one or more brain areas (&#62;1 mm), cannot be supported. Given these considerations, a different type of approach was needed to study how theta oscillations emerge in the hippocampus and propagate to related cortical and subcortical output&#160;structures.
In recent years, the initial development of the &#34;complete septo-hippocampal&#34; preparation to examine bidirectional interactions of the two structures22, and the ensuing evolution of the &#34;isolated hippocampus&#34; preparation, have revealed that intrinsic theta oscillations occur spontaneously in the hippocampus lacking external rhythmic input23. The value of these approaches lies on the initial insight that the whole functional structure of these regions had to be preserved in order to function as a theta rhythm generator in vitro22.

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			<td height="21" style="height:21px;width:319px;">Reagents</td>
			<td style="width:135px;"></td>
			<td style="width:119px;"></td>
			<td style="width:247px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sodium Chloride</td>
			<td>Sigma Aldrich</td>
			<td>S9625</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sucrose</td>
			<td>Sigma Aldrich</td>
			<td>S9378</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sodium Bicarbonate</td>
			<td>Sigma Aldrich</td>
			<td>S5761</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">NaH2PO4 - sodium phosphate monobasic</td>
			<td>Sigma Aldrich</td>
			<td>S8282</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Magnesium sulfate</td>
			<td>Sigma Aldrich</td>
			<td>M7506</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Potassium Chloride</td>
			<td>Sigma Aldrich</td>
			<td>P3911</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">D-(+)-Glucose</td>
			<td>Sigma Aldrich</td>
			<td>G7528</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Calcium chloride dihydrate</td>
			<td>Sigma Aldrich</td>
			<td>C5080</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sodium Ascorbate</td>
			<td>Sigma Aldrich</td>
			<td>A7631-25G</td>
			<td></td>
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		<tr height="30">
			<td height="30" style="height:30px;width:319px;">Name</td>
			<td style="width:135px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td>Comments</td>
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			<td height="21" style="height:21px;">Equipment</td>
			<td></td>
			<td></td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Standard Dissecting Scissors</td>
			<td>Fisher Scientific</td>
			<td>08-951-25</td>
			<td>brain extraction</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Scalpel Handle #4, 14cm</td>
			<td>WPI</td>
			<td>500237</td>
			<td>brain extraction</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Filter forceps, flat jaws, straight (11cm)</td>
			<td>WPI</td>
			<td>500456</td>
			<td>brain extraction</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Paragon Stainless Steel Scalpel Blades #20</td>
			<td>Ultident</td>
			<td>02-90010-20</td>
			<td>brain extraction</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Fine Point Curved Dissecting Scissors</td>
			<td>Thermo Fisher Scientific</td>
			<td>711999</td>
			<td>brain extraction</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Teflon (PTFE) -coated thin spatula</td>
			<td>VWR</td>
			<td>82027-534</td>
			<td>hippocampal preparation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Hayman Style Microspatula</td>
			<td>Fisher Scientific</td>
			<td>21-401-25A</td>
			<td>hippocampal preparation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Lab spoon</td>
			<td>Fisher Scientific</td>
			<td>14-375-20</td>
			<td>hippocampal preparation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Borosilicate Glass Pasteur Pipets</td>
			<td>Fisher Scientific</td>
			<td>13-678-20A</td>
			<td>hippocampal preparation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Droper</td>
			<td>Fisher Scientific</td>
			<td></td>
			<td>hippocampal preparation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Razor blades Single edged</td>
			<td>VWR</td>
			<td>55411-055</td>
			<td>hippocampal preparation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Lens paper (4X6 inch)</td>
			<td>VWR</td>
			<td>52846-001</td>
			<td>hippocampal preparation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Glass petri dishes (100 x 20 mm)</td>
			<td>VWR</td>
			<td>25354-080</td>
			<td>hippocampal preparation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Plastic tray for ice; size 30 x 20 x 5 cm</td>
			<td>n.a.</td>
			<td>n.a.</td>
			<td>hippocampal preparation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Single Inline Solution Heater</td>
			<td>Warner Instruments</td>
			<td>SH-27B</td>
			<td>perfusion system</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Aquarium air stones for bubbling</td>
			<td>n.a.</td>
			<td>n.a.</td>
			<td>perfusion system</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Tygon E-3603 tubing (ID 1/16 OD 1/8)</td>
			<td>Fisherbrand</td>
			<td>14-171-129</td>
			<td>perfusion system</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Electric Skillet</td>
			<td>Black &#38; Decker</td>
			<td>n.a.</td>
			<td>perfusion system</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">95% O2/5% CO2 gas mixture (carbogen)&#160;</td>
			<td>Vitalaire</td>
			<td>SG466204A</td>
			<td>perfusion system</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Glass bottles/flasks (4 x 1 L)</td>
			<td>n.a.</td>
			<td>n.a.</td>
			<td>perfusion system</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Submerged recording Chamber</td>
			<td>custom design (FM)</td>
			<td>n.a.</td>
			<td style="width:247px;">Commercial alternative may be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Glass pipettes (1.5 / 0.84 OD/ID (mm) )</td>
			<td>WPI</td>
			<td>1B150F-4</td>
			<td>electrophysiology</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Hum Bug 50/60 Hz Noise Eliminator</td>
			<td>Quest Scientific</td>
			<td>Q-Humbug</td>
			<td>electrophysiology</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Multiclamp 700B patch-clamp amplifier</td>
			<td>Molecular devices</td>
			<td>MULTICLAMP</td>
			<td>electrophysiology</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Multiclamp 700B Commander Program</td>
			<td>Molecular devices</td>
			<td>MULTICLAMP</td>
			<td>electrophysiology</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Digital/Analogue converter</td>
			<td>Molecular devices</td>
			<td>DDI440</td>
			<td>electrophysiology</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">PCLAMP10</td>
			<td>Molecular devices</td>
			<td>PCLAMP10</td>
			<td>electrophysiology</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Vibration isolation table&#160;</td>
			<td>Newport</td>
			<td>n.a.</td>
			<td>electrophysiology</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Micromanipulators (manually operated )</td>
			<td>Siskiyou&#160;</td>
			<td>MX130</td>
			<td>electrophysiology (LFP)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Micromanipulators (automated)</td>
			<td>Siskiyou&#160;</td>
			<td>MC1000e</td>
			<td>electrophysiology (patch)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Audio monitor&#160;</td>
			<td>A-M Systems</td>
			<td>Model 3300</td>
			<td>electrophysiology</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Micropipette/Patch pipette puller</td>
			<td>Sutter</td>
			<td>P-97</td>
			<td>electrophysiology</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Custom-built upright fluorescence microscope</td>
			<td>Siskiyou</td>
			<td>n.a.</td>
			<td>Imaging</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Analogue video camera</td>
			<td>COHU</td>
			<td>4912-2000/0000</td>
			<td>Imaging</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Digital frame grabber with imaging software</td>
			<td>EPIX, Inc</td>
			<td>PIXCI-SV7</td>
			<td>Imaging</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Olympus 2.5x objective</td>
			<td>Olympus</td>
			<td>MPLFLN</td>
			<td>Imaging</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Olympus 40x water immersion objective</td>
			<td>Olympus</td>
			<td>UIS2 LUMPLFLN</td>
			<td>Imaging</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Custom-made light-emitting diode (LED) system&#160;</td>
			<td>custom</td>
			<td>n.a.</td>
			<td>optogenetic stimulation (Amhilon et al., 2015)</td>
		</tr>
		<tr height="30">
			<td height="30" style="height:30px;width:319px;">Name</td>
			<td style="width:135px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Animals</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:319px;">PV::Cre (KI) mice</td>
			<td style="width:135px;">Jackson Laboratory</td>
			<td style="width:119px;">stock number 008069</td>
			<td style="width:247px;">Allow&#160; Cre-directed gene expression in PV interneurons</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:319px;">Constitutive-conditional Ai9 mice (R26-lox-stop-lox-tdTomato (KI))</td>
			<td style="width:135px;">Jackson Laboratory</td>
			<td style="width:119px;">stock number 007905</td>
			<td style="width:247px;">Express TdTomato following Cre-mediated recombination</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:319px;">Ai32 mice (R26-lox-stop-lox-ChR2(H134R)-EYFP</td>
			<td style="width:135px;">Jackson Laboratory</td>
			<td style="width:119px;">stock 
			number 012569</td>
			<td style="width:247px;">Express the improved channelrhodopsin-2/EYFP fusion protein following exposure to Cre recombinase</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">PVChY mice</td>
			<td>In house breeding</td>
			<td>n.a.</td>
			<td style="width:247px;">Offspring obtained from cross-breeding the PV-Cre line with Ai32 mice (R26-lox-stop-lox-ChR2(H134R)-EYFP</td>
		</tr>
	</tbody>
</table>

tuning in the hippocampal theta band in vitro: methodologies for recording from the isolated rodent septohippocampal circuit

This protocol outlines the procedures for preparing and recording from the isolated whole hippocampus, of WT and transgenic mice, along with recent improvements in methodologies and applications for the study of theta oscillations. A simple characterization of the isolated hippocampal preparation is presented whereby the relationship between internal hippocampal theta oscillators is examined together with the activity of pyramidal cells, and GABAergic interneurons, of the cornu ammonis-1 (CA1) and subiculum (SUB) areas. Overall, we show that the isolated hippocampus is capable of generating intrinsic theta oscillations in vitro and that rhythmicity generated within the hippocampus can be precisely manipulated by optogenetic stimulation of parvalbumin-positive (PV) interneurons. The in vitro isolated hippocampal preparation offers a unique opportunity to use simultaneous field and intracellular patch-clamp recordings from visually-identified neurons to better understand the mechanisms underlying theta rhythm generation.

This section illustrates examples of results that can be obtained by studying theta oscillations in the mouse isolated hippocampal preparation in vitro. The dissection procedure for extracting the isolated hippocampus is illustrated in Figure 1. Using this preparation, intrinsic theta oscillations can be examined during placement of multiple field electrodes, recording overall activity and synchronized synaptic inputs to neuronal populations in diffe...

Watch this Scientific Journal Video about Tuning in the Hippocampal Theta Band In Vitro: Methodologies for Recording from the Isolated Rodent Septohippocampal Circuit at JoVE.com

Tuning in the Hippocampal Theta Band In Vitro: Methodologies for Recording from the Isolated Rodent Septohippocampal Circuit

Here, we present a protocol for recording rhythmic neuronal network theta and gamma oscillations from an isolated whole hippocampal preparation. We describe the experimental steps from extraction of the hippocampus to details of field, unitary and whole-cell patch clamp recordings as well as optogenetic pacing of the theta rhythm.

Tuning in the Hippocampal Theta Band In Vitro: Methodologies for Recording from the Isolated Rodent Septohippocampal Circuit

This protocol outlines the procedures for preparing and recording from the isolated whole hippocampus, of WT and transgenic mice, along with recent ...

The overall goal of this protocol is to present procedures for extracting the whole hippocampal preparation and exploring the generation of rhythmic neuronal network using field, unitary and patch-clamp recordings as well as optogenetic stimulation. This method can help answer key questions in the neuroscience field of learning and memory by greatly facilitating the study of cellular and synaptic mechanisms that underlie rhythmic oscillations in the hippocampus. The main advantage of this technique is that it uses an optimized preparation to investigate the circuits in detailed oscillations, which play a crucial role in hippocampal dependent memory information.To begin this procedure, place the brain onto the dissection dish in an upright position, remove the cerebellum with a razor blade, then cut the brain in half along the mid sagittal plane and return the two isolated hemispheres to the holding chamber. Next, place the single hemisected brain upright onto the dissection dish. Rotate the dish until the mid sagittal structures are facing the experimenter and the embedded outline of the septal complex is visible as a thin pear shaped layer of tissue interior to the thalamus.Then, insert a coated spatula beneath the septum and move the tip down until the dissection dish is reached, sever the fibers that connect the septal area cordially. Now, repeat the same operation along the anterior edge of the septum and cut the fibers that connect to the frontal part of the brain. With the spatula lightly holding the inner part of the cortex just above the hippocampus in an upright position, use the microspatula to carefully pull down the thalamic, hypothalamic and the remaining brain stem nuclei.Subsequently, use the spatula to cut off and remove the pulled away tissue. Afterward, insert the coated spatula in the lateral ventricle underneath the rostral end of the dorsal hippocampus. Hold the spatula horizontally, aligned with the mid-sagittal plane of the hemisected brain and slide it through the smooth contour of the ventricular walls, until the tip emerges cordially.Hold the spatula beneath the hippocampus and lightly press it onto the connecting fibers along the inside layer where the hippocampus joins with the overlying cortex, then, apply the microspatula on the external side of this layer and press it against the coated spatula to slice through the connecting fibers. To complete the extraction, rotate the dissection dish and insert the coated spatula beneath the ventral hippocampus. Lightly hold the hippocampus with the spatula and cut through the endocrinal connections using a slicing motion against the spatula.Once isolation is complete, keep the hippocampus resting on the dish and add a drop of ice cold sucrose solution to keep it cool. Carefully trim off any remaining cortex and fibers and separate the hippocampus from the septum by gently applying a razor blade to the fornix. After that, transfer the preparation to room temperature sucrose solution and let it recover for 15 to 30 minutes before transferring to the recording chamber.In this step, set up the gravity fed perfusion system to enable continuous high speed in flow of oxygenated ACSF. Next, stop the ACSF flow and transfer the hippocampus to the recording chamber, using the wide end of a glass pipette. Allow the sucrose saturated preparation to sink and settle at the bottom.Place the preparation in the center of the recording chamber, with the smooth surface of CA1 and subiculum on top. Stabilize the hippocampus with small weights of the septal and temporal extremities and restart the ACSF flow. In this procedure, lower the LFP electrode to the surface of the hippocampus.Advance the LFP electrode through the parameter layer and observe the increase in extracellular spiking activity as single unit discharge from individual neurons is detected. Lower the electrode further and note that spiking begins to fade again as the tip crosses into the radiatom. Observe that a clearly visible network oscillation in the theta frequency range, becomes apparent and reaches maximum amplitude as the recording location is lowered through the radiatom.To test the spatial properties of spontaneous theta oscillations across the CA1 region, place a second LFP electrode into a CA1 site and observe that CA1 theta oscillations synchronize over large distances. To test the properties of theta oscillations across hippocampal layers, leave a reference LFP electrode in a CA1 radiatom site. Starting from just above the stratum oriens, lower a second electrode into the parameter cell layer and through the radiatom.Observe a gradual inversion of the LFP signal across the parameter layer. To test gamma oscillations and theta gamma coupling in the intact hippocampus, place a field electrode at the CA1 subiculum border and lower it until it sits at the interface between the parameter and molecular layers. At this level, a field potential displaying clear gamma oscillations with changes in amplitude that phase locked to the local theta rhythm can be recorded.Then, adjust the scaling to observe the slow time scale of theta gamma coupling. Note that gamma bursts occur in two distinct frequency bands which can be revealed by band during the ongoing LFP signal, in the slow and fast gamma range. For whole cell patch clamp recording during in vitro hippocampal theta oscillations, use fluorescence video microscopy with low and high power magnification to visualize tdTomato positive interneurons located near the surface of a hippocampal preparation from a PV tom mouse.Under low magnification view of the hippocampus, place an LFP electrode in CA1 subiculum to monitor theta oscillations while preparing for patch clamp experiments. Then, switch to 40x magnification and immerse the objective over the target region. Lower it until the top layers become visible, under the fluorescence microscopy, select the fluorescent PV tom cell and approach with a patch pipette filled with standard intercellular solution.Once in whole cell configuration, examine the physiological properties of the identified PV cell during spontaneous hippocampal oscillations. Observe the intecellular membrane potential recording from PV cells that are characterized from fast spiking behavior and bursts of action potentials synchronized to the ongoing CA1 subiculum theta rhythm. In this procedure, place an LFP electrode in the CA1 subiculum area and patch a nearby parameter cell in the isolated hippocampus of a mouse, expressing the blue light sensitive excitatory opcin, CHR2 in PV interneurons.Place an optic fiber light guide above the hippocampal preparation and center it on the recorded region. Use blue light from a LED source for optic genetic stimulation, which consists of 10 to 20 millisecond light pulses or sin wave voltage commands delivered at theta frequencies. In current clamp, characterize the activity of the recorded cell during spontaneous theta oscillations.Then, start the stimulation protocol and record the light responses. Observe that field oscillations and synaptic activity and the recorded neuron, become increasingly synchronized during optogenetic stimulation and that rhythmic pacing of PV cells results in a robust control of both frequency and power of theta oscillations. Shown here, is the electrophysiological activity recorded from a parameter cell during theta oscillations.Current clamp traces shows spontaneous but not rhythmic firing at rest and inhibitory post synaptic potentials that were not clearly synchronized with the slowly emerging LFP oscillation. Voltage clamp recordings show that the corresponding inhibitory post synaptic currents have the reversal potential at around minus 70 millivolts. And here, is the electrophysiological activity recorded from a fast spiking fluorescent PV interneuron during theta oscillations.In current clamp, this cell was spontaneously firing at rest and was strongly driven by rhythmic excitatory post synaptic potentials that were phased locked with the stable LFP oscillation. In voltage clamp recordings, the excitatory post synaptic current reversal potential was approximately at zero millivolts. Once mastered, this technique can be performed in two to three hours, while attempting this procedure, it's important to remember to oxygenate a solution vigorously and use a perfusion system allowing high speed but steady flow of armed ACSF over the preparation during the electrophysiological recordings.Following this procedure, other methods such as optogenetics stimulation or silencing of specific cell type populations, can be used in order to answer additional questions such as identifying the cellular subtypes that form theta oscillators in the hippocampus. After its development, this technique paved a way for researchers in the field of neuroscience to systematically explore the dynamics of theta oscillations across the septal temporal axis of the hippocampus in vitro. After watching this video, you should have a good understanding of how to extract a whole hippocampus preparation in order to explore the function of rhythmic neuronal networks using field, unit and patch clamp recordings, as well as optogenetic stimulation.

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Research

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Neuroscience

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

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 ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses specifically on antisense oligonucleotides (ASOs), though the techniques shown in the video here can also be used to deliver a plethora of other drugs to the CNS. Antisense oligonucleotides (ASOs) have the capability to knockdown sequence-specific targets 1 as well as shift isoform ratios of specific genes 2. To achieve widespread gene knockdown or splicing in the CNS of mice, the ASOs can be delivered into the brain using two separate routes of administration, both of which we demonstrate in the video.
The first uses Alzet osmotic pumps, connected to a catheter that is surgically implanted into the lateral ventricle. This allows the ASOs to be continuously infused into the CNS for a designated period of time. The second involves a single bolus injection of a high concentration of ASO into the right lateral ventricle. Both methods use the mouse cerebral ventricular system to deliver the ASO to the entire brain and spinal cord, though depending on the needs of the study, one method may be preferred over the other.

We describe a method to target drugs to the central nervous system by either implanting a catheter or performing a bolus injection into the right lateral ventricle in mice. We focus specifically on the delivery of antisense oligonucleotides. This technique is readily adaptable to other drugs and to rats.

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses ...

An In-vitro Preparation of Isolated Enteric Neurons and Glia from the Myenteric Plexus of the Adult Mouse

The enteric nervous system is a vast network of neurons and glia running the length of the gastrointestinal tract that functionally controls gastrointestinal motility. A procedure for the isolation and culture of a mixed population of neurons and glia from the myenteric plexus is described. The primary cultures can be maintained for over 7 days, with connections developing among the neurons and glia. The longitudinal muscle strip with the attached myenteric plexus is stripped from the underlying circular muscle of the mouse ileum or colon and subjected to enzymatic digestion. In sterile conditions, the isolated neuronal and glia population are preserved within the pellet following centrifugation and plated on coverslips. Within 24-48 hr, neurite outgrowth occurs and neurons can be identified by pan-neuronal markers. After two days in culture, isolated neurons fire action potentials as observed by patch clamp studies. Furthermore, enteric glia can also be identified by GFAP staining. A network of neurons and glia in close apposition forms within 5 - 7 days. Enteric neurons can be individually and directly studied using methods such as immunohistochemistry, electrophysiology, calcium imaging, and single-cell PCR. Furthermore, this procedure can be performed in genetically modified animals. This methodology is simple to perform and inexpensive. Overall, this protocol exposes the components of the enteric nervous system in an easily manipulated manner so that we may better discover the functionality of the ENS in normal and disease states.

An In-vitro Preparation of Isolated Enteric Neurons and Glia from the Myenteric Plexus of the Adult Mouse

We demonstrate a cell culture protocol for the direct study of neuronal and glial components of the enteric nervous system. A neuron/glia mixed culture on coverslips is prepared from the myenteric plexus of adult mouse providing the ability to examine individual neuron and glia function by electrophysiology, immunohistochemical, etc.

The enteric nervous system is a vast network of neurons and glia running the length of the gastrointestinal tract that functionally controls ...

Simultaneous Electrophysiological Recording and Micro-injections of Inhibitory Agents in the Rodent Brain

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing system was developed that allows for the injection of a drug while recording electrophysiological signals from the affected neuronal population. This method provides a simple and economical alternative to commercial solutions. A glass pipette was modified by combining it with a hypodermic needle and a silver filament. The injectrode is attached to commercial microsyringe pump for drug delivery. This results in a technique that provides real-time pharmacodynamics feedback through multi-unit extracellular signals originating from the site of drug delivery. As a proof of concept, we recorded neuronal activity from the superior colliculus elicited by flashes of light in rats, concomitantly with delivery of drugs through the injectrode. The injectrode recording capacity permits the functional characterization of the injection site favoring precise control over the localization of drug delivery. Application of this method also extends far beyond what is demonstrated here, as the choice of chemical substance loaded into the injectrode is vast, including tracing markers for anatomic experiments.

Here, we craft a glass pipette with dual functions: inhibition of deep brain structures by microinjections of drugs and real-time monitoring of their effects through simultaneous electrophysiological recordings.

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing ...

Utilizing Combined Methodologies to Define the Role of Plasma Membrane Delivery During Axon Branching and Neuronal Morphogenesis

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular guidance cues like netrin-1 and results in dramatic increases to the surface area of the axonal plasma membrane. Netrin-1-dependent axon branching likely involves temporal and spatial control of plasma membrane expansion, the components of which are supplied through exocytic vesicle fusion. These fusion events are preceded by formation of SNARE complexes, comprising a v-SNARE, such as VAMP2 (vesicle-associated membrane protein 2), and plasma membrane t-SNAREs, syntaxin-1 and SNAP25 (synaptosomal-associated protein 25). Detailed herein isa multi-pronged approach used to examine the role of SNARE mediated exocytosis in axon branching. The strength of the combined approach is data acquisition at a range of spatial and temporal resolutions, spanning from the dynamics of single vesicle fusion events in individual neurons to SNARE complex formation and axon branching in populations of cultured neurons. This protocol takes advantage of established biochemical approaches to assay levels of endogenous SNARE complexes and Total Internal Reflection Fluorescence (TIRF) microscopy of cortical neurons expressing VAMP2 tagged with a pH-sensitive GFP (VAMP2-pHlourin) to identify netrin-1 dependent changes in exocytic activity in individual neurons. To elucidate the timing of netrin-1-dependent branching, time-lapse differential interference contrast (DIC) microscopy of single neurons over the order of hours is utilized. Fixed cell immunofluorescence paired with botulinum neurotoxins that cleave SNARE machinery and block exocytosis demonstrates that netrin-1 dependent axon branching requires SNARE-mediated exocytic activity.

Light microscopy techniques coupled with biochemical assays elucidate the involvement of SNARE-mediated exocytosis in netrin-dependent axon branching. This combination of techniques permits identification of molecular mechanisms controlling axon branching and cell shape change.

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular ...

Recording Gamma Band Oscillations in Pedunculopontine Nucleus Neurons

Synaptic efferents from the PPN are known to modulate the neuronal activity of several intralaminar thalamic regions (e.g., the centrolateral/parafascicular; Cl/Pf nucleus). The activation of either the PPN or Cl/Pf nuclei in vivo has been described to induce the arousal of the animal and an increment in gamma band activity in the cortical electroencephalogram (EEG). The cellular mechanisms for the generation of gamma band oscillations in Reticular Activating System (RAS) neurons are the same as those found to generate gamma band oscillations in other brains nuclei. During current-clamp recordings of PPN neurons (from parasagittal slices from 9 - 25 day-old rats), the use of depolarizing square steps rapidly activated voltage-dependent potassium channels that prevented PPN neurons from being depolarized beyond -25 mV.
Injecting 1 - 2 sec long depolarizing current ramps gradually depolarized PPN membrane potential resting values towards 0 mV. However, injecting depolarizing square pulses generated gamma-band oscillations of membrane potential that showed to be smaller in amplitude compared to the oscillations generated by ramps. All experiments were performed in the presence of voltage-gated sodium channels and fast synaptic receptors blockers. It has been shown that the activation of high-threshold voltage-dependent calcium channels underlie gamma-band oscillatory activity in PPN neurons. Specific methodological and pharmacological interventions are described here, providing the necessary tools to induce and sustain PPN subthreshold gamma band oscillation in vitro.

The pedunculopontine nucleus (PPN) is located in the brainstem and its neurons are maximally activated during waking and rapid eye movement (REM) sleep brain states. This work describes the experimental approach to record in vitro gamma band subthreshold membrane oscillation in PPN neurons.

Synaptic efferents from the PPN are known to modulate the neuronal activity of several intralaminar thalamic regions (e.g., the ...

Recording Synaptic Plasticity in Acute Hippocampal Slices Maintained in a Small-volume Recycling-, Perfusion-, and Submersion-type Chamber System

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful analysis of synaptic transmission modulation when performing field potential or intracellular recordings. This manuscript describes methodological aspects that might be helpful in improving experimental conditions for the maintenance of acute brain slices and for recording field excitatory postsynaptic potentials in a commercially available submersion chamber with an outflow-carbogenation unit. The outflow-carbogenation helps to stabilize the oxygen level in experiments that rely on the recycling of a small buffer reservoir to enhance the cost-efficiency of drug experiments. In addition, the manuscript presents representative experiments that examine the effects of different carbogenation modes and stimulation paradigms on the activity-dependent synaptic plasticity of synaptic transmission.

This protocol describes the stabilization of the oxygen level in a small volume of recycled buffer and methodological aspects of recording activity-dependent synaptic plasticity in submerged acute hippocampal slices.

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful ...

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 ...