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 oscillations. In this protocol, the timely performance of procedures &#8211; from dissection to recording &#8211; is particularly important because the isolated hippocampus constitutes such a dense, but delicate, preparation that maintaining functional connectivity of the structure in vitro requires great care. Preparing everything beforehand ensures that an adequate level of perfusion is available as early as possible to minimize cell damage and maintain physiological function.
<ol>
	<li>Sucrose-based artificial Cerebral Spinal Fluid (aCSF) solution
	<ol>
		<li>Prepare 1X stock sucrose solution to be used for dissection and post-dissection incubation. Combine all components listed in Table 1, except for CaCl2, into 1 L of deionized water, and store at 4 &#176;C for up to 2 weeks.</li>
	</ol>
	</li>
	<li>Standard aCSF solution
	<ol>
		<li>Prepare standard aCSF for high flow rate perfusion of the isolated hippocampus during electrophysiological recordings. To make the concentrated (5X) stock aCSF, dissolve all the components listed in Table 2, except CaCl2, in 2 L of deionized water and store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Prepare the equipment
	<ol>
		<li>Before starting dissection, set up the bench area with an ice-filled plastic tray (30&#160;x 20&#160;x 5 cm), carbogen connected tubing lines and three glass Petri dishes (10 x 2 cm) (See Figure 1).</li>
		<li>Add 300 mL of sucrose solution (1X stock) to a 500 mL beaker, bubble it with carbogen (95% O2, 5% CO2) and add Ca2+ [1.2 mM].</li>
		<li>Transfer 60 mL of this sucrose solution to a glass Petri dish and leave at room temperature. Place the remainder in a 4 &#176;C freezer for 10 - 15 min.</li>
		<li>Bubble the ice-cold sucrose solution with carbogen throughout the dissection.</li>
		<li>Fill a second Petri dish (cold holding chamber) with sucrose solution (4 &#176;C) and keep on ice.</li>
		<li>Add 30 mL of ice-cold sucrose solution to a 50 mL beaker.</li>
	</ol>
	</li>
</ol>
2. Whole Hippocampus Dissection
NOTE: The method for dissecting the isolated hippocampus is essentially identical to the one developed and described originally22, but with additional details and changes regarding the perfusion rate and recording techniques.
<ol>
	<li>Brain dissection
	<ol>
		<li>Anesthetize mouse (P20 - 35) under a fume hood with a small paper towel soaked with 1 mL of isoflurane (100%) that is added into the cage.</li>
		<li>Decapitate the anesthetized mouse and quickly submerge the head in ice-cold sucrose solution (50 mL beaker, ~ 5 s). Transfer onto a paper towel.</li>
		<li>Use a scalpel to make a medial skin incision (caudal to rostral) and push skin on the sides to expose the cranium.</li>
		<li>Cut open the skull with a small scissors and use a spatula to rapidly extract the brain and drop it into the Petri dish containing ice-cold sucrose solution. Allow 1-2 min for the brain to cool.</li>
		<li>While the brain is recovering, add 2 - 3 mL of sucrose solution onto a lens paper covered Petri (dissection) dish on ice.</li>
	</ol>
	</li>
	<li>Isolating the hippocampus
	<ol>
		<li>Place the brain onto the dissection dish in an upright position.</li>
		<li>Remove the cerebellum and frontal cortex with a razor blade. Cut the brain in half along the midsagittal plane and return the two isolated hemispheres to the holding chamber. Allow a further 1 - 2 min for recovery.</li>
		<li>Place single hemisected brain upright onto the dissection dish.</li>
		<li>Rotate the dish until midsagittal structures face the observer and the embedded outline of the septal complex is visible as a thin pear-shaped layer of tissue anterior to the thalamus.</li>
		<li>Hold the hemi-sected brain steady with the microspatula and place the small end of the polytetrafluoroethylene (PTFE)-coated spatula immediately behind (caudal to) the septal area.</li>
		<li>Insert the coated spatula beneath the septum and move the tip down until the dissection dish is reached. Sever the fibers that connect the septal area caudally. Repeat the same operation along the anterior edge of the septum and cut the fibers that connect to the frontal part of the brain.</li>
		<li>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 remaining brain stem nuclei. Use the spatula to cut off and remove the pulled away tissue.</li>
		<li>Turn the isolated hemisphere on its side and identify the outline of the hippocampus.</li>
		<li>Insert the coated spatula in the lateral ventricle, underneath the rostral end of the dorsal hippocampus.</li>
		<li>Hold the spatula horizontally aligned with the midsagittal plane of the hemi-sected brain and slide it through the smooth contour of ventricular walls until the tip emerges caudally.</li>
		<li>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. Apply the microspatula on the external side of this layer and slide against the coated spatula to slice through the connecting fibers.</li>
		<li>To complete the extraction, rotate the dissection dish and insert the coated spatula beneath the ventral hippocampus. Lightly hold the inside of the cortical fold with the spatula and cut through the entorhinal connections using a slicing motion against the microspatula. 
		NOTE: The whole septohippocampal complex can be removed by placing the coated spatula under the entire hippocampus, and pulling out the remaining brain tissue from underneath.</li>
		<li>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 through the fornix. 
		NOTE: If patch clamp recordings are to be performed from pyramidal cells (see Section 4.6 Optogenetic control), the isolated hippocampus can be transversally cut at a 45&#176; angle to expose the pyramidal layer of CA1 (&#34;angle-cut&#34; preparation), without compromising the local network circuitry in the remaining portion of the hippocampus.</li>
		<li>Transfer the preparation to room temperature sucrose solution and let it recover for 15 - 30 min before transferring to the recording chamber.</li>
	</ol>
	</li>
</ol>
3. Set Up the Fast Perfusion for Recording the Isolated Hippocampus
<ol>
	<li>Set up the gravity-fed perfusion system to enable continuous high-speed inflow of oxygenated aCSF at 20 - 25 mL/min.
	<ol>
		<li>Prepare aCSF (1X) flasks in advance and immerse them in a warming bath (~30 &#176;C) at least 15 min before start of experiment. Turn on the in-line solution heater system (30 &#176;C).</li>
		<li>Use aquarium bubblers and tubing lines connected to a carbogen tank to oxygenate aCSF throughout the experiment, and add CaCl2 [2 mM] prior to the start of perfusion.</li>
		<li>Stop 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.</li>
		<li>Place the preparation in the center of the recording chamber (in line with the inlet-outlet axis) with the smooth surface of CA1 and SUB on top. Stabilize the hippocampus with small weights at the septal and temporal extremities and restart aCSF flow.</li>
	</ol>
	</li>
</ol>
4. Electrophysiology in the Isolated Hippocampus
<ol>
	<li>Extracellular recording of in vitro hippocampal theta oscillations
	<ol>
		<li>For extracellular monitoring of Local Field Potential (LFP) or single-unit activity from the in vitro isolated hippocampus, use glass micropipettes (1 - 3 M&#937;) filled with aCSF. Record low-noise AC-coupled field potential signals with a patch clamp amplifier of gain x1000, online band-pass filtering 0.1 - 500 Hz and a sampling rate of 5-20 kHz. 
		NOTE: The custom-built microscope used in this protocol is equipped with low- (2.5X) and high-magnification (40X) objectives for low-magnification view of the hippocampus, as well as infrared video microscopy and epifluorescence for single cell recognition. The components of this system include the upright fluorescence microscope, fluorescence filter cubes, an analogue video camera and digital frame grabber with imaging software, an on-stage custom perfusion chamber (Figure 2A, inset i) and high-precision micromanipulators for neuronal recordings and optic fiber placement.</li>
		<li>Use the camera mounted on the microscope and connected to a computer display to inspect the hippocampal preparation under low magnification (2.5X) and rapidly check that there are no undercuts or damage to the external surface. The diagonally oriented arrangement of alveus fibers along the smooth surface of CA1 should be visible (Figure 2A, inset ii) when shining transmitted light through the hippocampus.</li>
		<li>Use the low-magnification wide-field objective to view the isolated hippocampus and the placement of LFP electrodes in specific recording locations. 
		NOTE: A layout of the isolated hippocampal preparation with multiple electrodes placed in different recording locations is illustrated in Figure 2A.</li>
		<li>Lower an LFP electrode into the bath until it touches the surface (alveus) of the CA1 area. 
		NOTE: This usually produces a fast transient in the AC-coupled recording similar to what happens when the electrode comes in contact with the recording media. An audio monitor can be useful to listen to the LFP channel signal after this step. 
		NOTE: Oftentimes, early signs of activity will only appear after 10 - 15 min, therefore it is important to not advance quickly into the preparation. If after this period the surface LFP electrode is still not picking up activity, advance a little further down through the stratum oriens and pause periodically to see if any form of activity arises while approaching the principal cell layer. If nothing is detected after a further 5 - 10 min, carefully retract the electrode and place it elsewhere along the same region.</li>
		<li>Advance the LFP electrode through the pyramidal layer. Observe that extracellular spiking activity initially increases as single-unit discharge from individual neurons (mostly pyramidal and inhibitory basket cells) is detected. Lower the electrode further and note that spiking begins to fade again as the tip crosses into the radiatum. Observe that a clearly visible network oscillation in the theta-frequency range (4 - 12 Hz) becomes apparent and reaches maximum amplitude (~100 - 300 &#181;V) as the recording location is lowered through the radiatum.</li>
	</ol>
	</li>
	<li>Theta oscillations across a single region
	<ol>
		<li>To test the spatial properties of spontaneous theta oscillations across the CA1 region, place the tip of a reference LFP electrode in a medial CA1 site (medially located along the longitudinal septo-temporal axis) and lower it into the radiatum as described previously.</li>
		<li>Place a second LFP electrode into a CA1 site (200 - 800 &#181;m septal or temporal from the first LFP) and observe that CA1 theta oscillations can synchronize over relatively large distances.</li>
	</ol>
	</li>
	<li>Theta oscillations across layers
	<ol>
		<li>To test the properties of theta oscillations across hippocampal layers, leave a reference LFP electrode in a CA1 radiatum site. Starting from just above the stratum oriens, lower a second electrode into the pyramidal cell layer, and through the radiatum. Observe a gradual inversion of the LFP signal (relative phase reversal) across the pyramidal layer. 
		NOTE: For representative examples of these types of activity, see Figure 2B. Examples of laminar/depth profiles and Current Source Density (CSD) analysis illustrating the site of phase reversal in isolated hippocampi have been published previously23,24,25.</li>
	</ol>
	</li>
	<li>Gamma oscillations and theta-gamma coupling in the intact hippocampus
	<ol>
		<li>Place a field electrode at the CA1/SUB border and lower it until it sits at the interface between the pyramidal and molecular layers. At this level, a field potential displaying clear gamma oscillations with changes in amplitude that phase-lock to the local theta rhythm can be recorded24. Adjust 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-pass filtering the ongoing LFP signal in the slow (25 - 50 Hz) and fast gamma (150 - 250 Hz) range (See Figure 2C for representative filtered traces).</li>
	</ol>
	</li>
	<li>Whole-cell patch clamp recording during in vitro hippocampal theta oscillations 
	NOTE: In this part of the protocol, the goal is to obtain visually guided whole-cell patch clamp recordings from identified interneurons in isolated hippocampi from transgenic PV-TOM mice expressing the fluorescent protein, tdTomato, under the control of the PV promoter (for more details see materials and Ref 26).
	<ol>
		<li>Use fluorescence video-microscopy with low (2.5X) and high power (40X) magnification to visualize tdTomato-positive interneurons located near the surface of a hippocampal preparation from a PV-TOM mouse (Figure 3). Note that while PV-TOM neurons can be successfully visualized and targeted for patch through the alveus (up to a 100 &#181;m), non-fluorescent pyramidal cells can also be reached relatively easily when the hippocampus is prepared with the angle-cut technique (see Section 2.2.14 note).</li>
		<li>Under low magnification view of the hippocampus, place an LFP electrode in CA1/SUB to monitor theta oscillations while preparing for patch clamp experiments.</li>
		<li>Switch to 40X magnification and immerse the objective over the target region; Lower it until the top layers become visible. Manipulate the position of the microscope to ensure sufficient open access for patch pipette approach from the opposite side.</li>
		<li>Under fluorescence microscopy, select a fluorescent PV-TOM cell and approach it with a patch pipette (2 - 3 M&#937;) filled with standard intracellular solution.</li>
		<li>Use a mouth piece to apply light positive pressure to the pipette and monitor the resistance as the electrode is lowered onto the cell. When the tip encounters the target cell, the pipette resistance increases and a clear dimple appears as positive pressure pushes on the membrane. Release the pressure and check that the current pulse flattens as a seal starts to form. Immediately clamp to a hyperpolarized potential (-70 mV), and once a G&#937; seal is achieved, rupture the cell membrane with a small amount of negative pressure applied through the pipette holder.</li>
		<li>Once in whole-cell configuration, examine the physiological properties of identified PV cell during spontaneous hippocampal oscillations (Figure 3B).</li>
		<li>Observe that intracellular membrane potential recording from PV cells are characterized by fast-spiking behavior and bursts of action potentials synchronized to the ongoing CA1/SUB theta rhythm.</li>
		<li>Switch to voltage-clamp and hold the cell at different membrane potentials to characterize the ratio of excitatory versus inhibitory synaptic currents generated by the intrinsic rhythmic activity. An example of patch clamp recording from a pyramidal cell is shown in Figure 3A for comparison.</li>
	</ol>
	</li>
	<li>Optogenetic control in the isolated hippocampus 
	NOTE: For optogenetic control of hippocampal network activity, a cell-type specific strategy involving rhythmic activation of PV-containing GABAergic cells is used here as these cells were shown to play a major role in synchronizing principal cell populations in the isolated hippocampus25. Selective expression of the excitatory channelrhodopsin-2 (ChR2) in PV cells is achieved by crossing the PV-Cre mouse line with mice constitutively expressing ChR2 Cre-dependently (Ai32), thereby generating PV-ChY mice. Alternatively, viral vector constructs (e.g., AAVdj-ChETA-eYFP) can be injected into the hippocampus of PV-Cre or PV-TOM mice (P15-16) to drive the expression of the excitatory opsin in CA1/SUB interneurons (Figure 4A). 
	NOTE: This strategy has been described previously25.
	<ol>
		<li>Place an LFP electrode in the CA1/SUB area and patch a nearby pyramidal cell in the isolated hippocampus of a mouse expressing the blue-light sensitive excitatory opsin ChR2 in PV interneurons.</li>
		<li>Place an optic fiber light guide (0.2 - 1 mm diameter) above the hippocampal preparation and center it on the recorded region. Use blue light (473 nm) from a LED source for optogenetic stimulation, consisting of light pulses (10 - 20 ms, triggered by a transistor-transistor logic signal) or sine wave voltage commands delivered at theta frequencies (4 - 12 Hz). 
		NOTE: The custom LED-based light stimulation assembly has been described elsewhere25.</li>
		<li>Switch to current clamp and characterize the activity of the pyramidal during spontaneous theta oscillations.</li>
		<li>Start the stimulation protocol and record the light responses.Observe that field oscillations and synaptic activity in 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 (Figure 4).</li>
	</ol>
	</li>
</ol>

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Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Excitatory+Inputs+Determine+Phase-Locking+Strength+and+Spike-Timing+of+CA1+Stratum+Oriens/Alveus+Parvalbumin+and+Somatostatin+Interneurons+during+Intrinsically+Generated+Hippocampal+Theta+Rhythm.">Excitatory Inputs Determine Phase-Locking Strength and Spike-Timing of CA1 Stratum Oriens/Alveus Parvalbumin and Somatostatin Interneurons during Intrinsically Generated Hippocampal Theta Rhythm.</a> J Neurosci. 36 (25), 6605-6622 (2016).</li><li>Gu, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NMDA-dependent+phase+synchronization+between+septal+and+temporal+CA3+hippocampal+networks.">NMDA-dependent phase synchronization between septal and temporal CA3 hippocampal networks.</a> J Neurosci. 33 (19), 8276-8287 (2013).</li><li>Jackson, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reversal+of+theta+rhythm+flow+through+intact+hippocampal+circuits.">Reversal of theta rhythm flow through intact hippocampal circuits.</a> Nat Neurosci. 17 (10), 1362-1370 (2014).</li><li>Gonzalez-Burgos, G., Lewis, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=GABA+neurons+and+the+mechanisms+of+network+oscillations:+implications+for+understanding+cortical+dysfunction+in+schizophrenia.">GABA neurons and the mechanisms of network oscillations: implications for understanding cortical dysfunction in schizophrenia.</a> Schizophr Bull. 34 (5), 944-961 (2008).</li></ol>

The authors declare no competing commercial or financial interests.

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 hippocampus is preserved together with functional connectivity and properties of local neuronal populations. This provides a complex and powerful tool to investigate small and large-scale network oscillations in the hippocampus. For example, the combination of prewired circuitry and cell-type specific properties in the network produces intrinsically and spontaneously generated rhythmic theta and gamma oscillations that mimic important features of hippocampal activity in vivo. Using the methods presented in this protocol, the hippocampus can be extracted quickly from the rodent brain and remain viable for several hours in a fast flowing aCSF environment. Basic electrophysiological techniques are readily applied to investigate how synchronous activity and synaptic function of specific neuronal types influence hippocampal dynamics.
Our procedure has provided the first opportunity to systematically explore the dynamics of local field potential oscillations across the whole septo-temporal (dorso-ventral) axis of the hippocampus in vitro (see Refs 22,23,24,25,26,27,28). At the network and physiological levels, the intact preparation preserves: 1) The functional synaptic interactions between cells required to reliably support intrinsic network oscillations with the frequency range and profile of theta waves in vivo, 2) The sharp change in relative phase of theta oscillation at the level of stratum pyramidale, 3) The spatial distribution and coherence of theta oscillation in the hippocampus and its interaction with local gamma frequency rhythms, 4) The amplitude-phase relationship of theta/gamma waves and 5) The specific temporal association of principal cell and interneuron firing with field potential theta oscillations.
Whereas the absence of external input to the hippocampus may appear as a limitation to the technique, it also allows the study of the internal dynamics of the hippocampus in a manner that cannot be done in vivo. In addition, external inputs can be mimicked by optogenetic stimulation of specific fiber terminals and afferent pathways. Intact preparations offer new possibilities to study how network activities propagate and interact in large-scale networks involving the hippocampus and other connected structures. As such, one main application of the intact preparation technique is the investigation of functional network connectivity within or between brain regions. Using the in vitro intact hippocampus should therefore not only provide further information on cellular and network properties of the hippocampus, but also shed light on how these interact to shape processing, integration and generation of information within the brain.
Rhythmic network oscillations engaging the coordinated electro-chemical signaling of large neuronal ensembles act as a central mechanism for encoding, storing and transferring information within and across brain areas. Hippocampal oscillations are thought to be crucial for processing of spatiotemporal information, encoding, and memory. Conversely, hippocampal dysfunction is thought to underlie disorders such as Alzheimer&#39;s disease and schizophrenia, which are associated with altered oscillation patterns29. As such, the study of self-generated hippocampal oscillations using intact hippocampal preparations has become a renewed means for elucidating basic properties of nervous system function and dysfunction.

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.

<table border="0" cellpadding="0" cellspacing="0" width="820">
	<tbody>
		<tr height="21">
			<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>
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		<tr height="21">
			<td height="21" style="height:21px;">Sucrose</td>
			<td>Sigma Aldrich</td>
			<td>S9378</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Sodium Bicarbonate</td>
			<td>Sigma Aldrich</td>
			<td>S5761</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">NaH2PO4 - sodium phosphate monobasic</td>
			<td>Sigma Aldrich</td>
			<td>S8282</td>
			<td></td>
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			<td height="21" style="height:21px;">Magnesium sulfate</td>
			<td>Sigma Aldrich</td>
			<td>M7506</td>
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			<td height="21" style="height:21px;">Potassium Chloride</td>
			<td>Sigma Aldrich</td>
			<td>P3911</td>
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		<tr height="21">
			<td height="21" style="height:21px;">D-(+)-Glucose</td>
			<td>Sigma Aldrich</td>
			<td>G7528</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Calcium chloride dihydrate</td>
			<td>Sigma Aldrich</td>
			<td>C5080</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Sodium Ascorbate</td>
			<td>Sigma Aldrich</td>
			<td>A7631-25G</td>
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			<td height="30" style="height:30px;width:319px;">Name</td>
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			<td>Comments</td>
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			<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 different regions and layers of the isolated hippocampus (Figure 2). Representative results from simultaneous whole-cell patch clamp and extracellular recordings are presented to characterize the firing and synaptic properties of specific cell types during spontaneous hippocampal theta oscillations (Figure 3), as well as during optogenetic manipulation of rhythmic activity (Figure 4).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/55851/55851fig1.jpg" /> 
Figure 1: Dissection Procedure for the Isolated Intact Hippocampus Preparation. 
(a) General view of the dissection setup. Top right: carbogenated ice-cold sucrose solution flask (1); bottom left: ice-filled plastic tray (2) holding the dissection dish covered with lens paper (3); the cold holding chamber containing sucrose solution (4); and a set of surgical tools (5). (b) View of the mouse brain before hemisection on the dissection dish. (c) Recovery of hemisected brain in the cold holding chamber and zoomed view (inset) of the left brain hemisphere before inserting the small end of the coated spatula under the septum. (d) Coated spatula placed under the isolated hippocampus, along the CA1/SUB region, with the remaining brain tissue is pulled out from underneath. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55851/55851fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/55851/55851fig2.jpg" /> 
Figure 2: Configuration of the Setup for Recording in vitro Theta Oscillations from the Intact Hippocampal Preparation in the Submerged Recording Chamber. 
(a) The isolated hippocampus is shown with a layout of hippocampal regions and multiple electrodes placed in four different recording sites distributed septotemporally (indicated by white asterisks). In the view of the recording chamber platform shown above (inset i), the inlet and outlet for fast-perfusion flow are indicated by numbers (1, 2). In the enlarged image of the hippocampus shown below (inset ii), a single electrode is placed in the midsepetotemporal CA1 and fibers of the alveus are readily visible, running diagonally toward the subiculum. S: septal, T: temporal, f/fx: fimbria-fornix. (b) Schematic representation of the organization of CA1 layers with representative LFP traces recorded simultaneously from stratum oriens (grey) and stratum radiatum (black). Note the inverted phase of signals between the two layers. Alv: stratum alveus, PR: stratum pyramidale, SR: stratum radiatum. SLM: Stratum Lacunosum Moleculare. (c) Example LFP trace showing spontaneous theta oscillation recorded from the CA1/SUB area (20 sec segment) and 2 sec expanded segments (below) from unfiltered signal; band-pass filtered for theta frequencies (0.5 - 12 Hz); slow gamma (25 - 55 Hz); and fast gamma (125 - 250 Hz). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55851/55851fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/55851/55851fig3.jpg" /> 
Figure 3: Cell-type Specific Activity of Pyramidal Cell and PV Interneuron during Spontaneous Theta Oscillations in the Intact Hippocampal Preparation from a PV-TOM Mouse. 
(a) Synaptic activity recorded from a pyramidal cell during theta oscillations. Current-clamp traces (left panel) show spontaneous but not rhythmic firing at rest, and inhibitory postsynaptic potentials (iPSPs) that were not clearly synchronized with the slowly emerging LFP oscillation. Voltage-clamp recordings (right panel) show that the corresponding inhibitory postsynaptic currents (iPSCs) have their reversal potential around -70 mV. (b) Synaptic activity recorded from a fast-spiking fluorescent PV interneuron during theta oscillations. In current-clamp (left), this cell was spontaneously firing at rest and was strongly driven by rhythmic excitatory postsynaptic potentials (ePSPs) that were phase-locked with the stable LFP oscillation. In voltage-clamp recordings (right), the excitatory postsynaptic current reversal potential was approximately 0 mV. Schematic drawings on top show the experimental setup together with the placement of patch and field electrodes in the CA1/SUB and high (40X) magnification images of the recorded pyramidal cell and fluorescent PV interneuron. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55851/55851fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/55851/55851fig4.jpg" /> 
Figure 4: Local Field Potential and Simultaneous Patch Clamp Recordings from Single CA1/SUB Pyramidal and PV Neurons during Spontaneous and Optogenetically-driven Theta Oscillations in the Isolated Hippocampus. 
(a) Diagram showing the LFP and patch electrode recording sites with placement of the light-fiber guide above the CA1/SUB region expressing a blue-light sensitive opsin (ChETA coupled with the fluorophore eYFP). (b) Characterization of a pyramidal neuron showing typical Regular-Spiking (RS) properties (top) and high magnification (40X) image of the recorded cell (bottom). (c) Sample current-clamp recording of the same cell at depolarized membrane potential (black) together with the LFP signal (grey) and showing synchronized rhythmic IPSPs during spontaneous oscillation (left) and during theta-frequency (6 Hz) light stimulation (right). The pattern of light stimulation (blue shading) is depicted on top of the voltage traces. (d) Spectrogram and power spectra of LFP waveform before, during and after a 6 Hz light simulation (baseline, stim, post). (e) Low magnification bright-field and fluorescence images of the isolated hippocampal preparation showing eYFP fluorescence (in green) localized in the CA1/SUB region. (f) Current steps characterization showing Fast-Spiking (FS) behaviour of a recorded PV-TOM interneuron (40X fluorescence image below). (g) Membrane potential recording showing large EPSPs and rhythmic firing of the recorded PV cell synchronized with the LFP signal during spontaneous field oscillation (left) and during light stimulation (right) at 3 Hz. (h) Field Triggered Average (FTA) of PV cell spikes over the CA1/SUB LFP signal. The discharge of the PV cell over multiple trials (centered on the LFP peaks) was converted to a FTA of spikes recorded during spontaneous oscillation (baseline) and during light stimulation (stim). Middle and bottom graphs show the raster plots of spikes and spike probability histograms (average probability is shown in red). Topmost graphs show plots of the average LFP signal which increased in power during light stimulation in parallel with highly synchronized firing of the PV cell phase-locked to the peak of the LFP oscillation. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55851/55851fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="4">SUCROSE SOLUTION (1X) FOR ISOLATED HIPPOCAMPUS</td>
		</tr>
		<tr>
			<td colspan="2">(Stock solution)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Compound</td>
			<td>MW</td>
			<td>Final Conc. (mM)</td>
			<td>Amount for 1 L (g)</td>
		</tr>
		<tr>
			<td>sucrose</td>
			<td>342.3</td>
			<td>252</td>
			<td>86.26 g</td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>84.01</td>
			<td>24</td>
			<td>2.020 g</td>
		</tr>
		<tr>
			<td>glucose</td>
			<td>180.2</td>
			<td>10</td>
			<td>1.800 g</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>74.55</td>
			<td>3</td>
			<td>0.223 g</td>
		</tr>
		<tr>
			<td>MgSO4</td>
			<td>120.4</td>
			<td>2</td>
			<td>0.241 g</td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>120</td>
			<td>1.25</td>
			<td>0.150 g</td>
		</tr>
		<tr>
			<td>CaCl2.2H2O [1 M] stock</td>
			<td>147</td>
			<td>1.2</td>
			<td>120 &#956;L / 0.1 L *</td>
		</tr>
		<tr>
			<td colspan="4">* add 360 &#956;L CaCl2 [1 M] for 0.3 L oxygenated sucrose solution&#160;</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td colspan="2">pH = 7.4 when oxygenated, Osm 310 - 320&#160;</td>
		</tr>
	</tbody>
</table>
Table 1.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="4">STANDARD ACSF SOLUTION (5X) FOR PERFUSION</td>
		</tr>
		<tr>
			<td colspan="2">(Stock solution)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Compound</td>
			<td>MW</td>
			<td>Final Conc. (mM)</td>
			<td>Amount for 2 L&#160; 5X</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>58.44</td>
			<td>126</td>
			<td>73.6</td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>84.01</td>
			<td>24</td>
			<td>20.2</td>
		</tr>
		<tr>
			<td>glucose</td>
			<td>180.2</td>
			<td>10</td>
			<td>18</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>74.55</td>
			<td>&#9830; 4.5</td>
			<td>3.355</td>
		</tr>
		<tr>
			<td>MgSO4</td>
			<td>120.4</td>
			<td>2</td>
			<td>2.41</td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>120</td>
			<td>1.25</td>
			<td>1.5</td>
		</tr>
		<tr>
			<td>Ascorbate</td>
			<td>176.1</td>
			<td>0.4</td>
			<td>0.705</td>
		</tr>
		<tr>
			<td>CaCl2.2H2O [1 M] stock</td>
			<td>147</td>
			<td>2</td>
			<td>2 mL / L *</td>
		</tr>
		<tr>
			<td colspan="4">* add 2 mL CaCl2 [1 M] for 1 L aCSF (1x) oxygenated solution&#160;</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td colspan="2">pH = 7.4 when oxygenated, Osm 310 - 320&#160;</td>
		</tr>
		<tr>
			<td colspan="4">&#9830; A slightly elevated [K+]o is used for this aCSF solution (compared to normal aCSF 2.5 mM KCl) to increase excitability of hippocampal networks and facilitate the emergence of theta oscillations.</td>
		</tr>
	</tbody>
</table>
Table 2.

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 ...

tuning-hippocampal-theta-band-vitro-methodologies-for-recording-from

Nanyang Technological University

Research

JoVE Journal

Neuroscience

9.7K Views.  McGill University. 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.

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

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 ...

Multi-unit Recording Methods to Characterize Neural Activity in the Locust (Schistocerca Americana) Olfactory Circuits

Detection and interpretation of olfactory cues are critical for the survival of many organisms. Remarkably, species across phyla have strikingly similar olfactory systems suggesting that the biological approach to chemical sensing has been optimized over evolutionary time1. In the insect olfactory system, odorants are transduced by olfactory receptor neurons (ORN) in the antenna, which convert chemical stimuli into trains of action potentials. Sensory input from the ORNs is then relayed to the antennal lobe (AL; a structure analogous to the vertebrate olfactory bulb). In the AL, neural representations for odors take the form of spatiotemporal firing patterns distributed across ensembles of principal neurons (PNs; also referred to as projection neurons)2,3. The AL output is subsequently processed by Kenyon cells (KCs) in the downstream mushroom body (MB), a structure associated with olfactory memory and learning4,5. Here, we present electrophysiological recording techniques to monitor odor-evoked neural responses in these olfactory circuits.
First, we present a single sensillum recording method to study odor-evoked responses at the level of populations of ORNs6,7. We discuss the use of saline filled sharpened glass pipettes as electrodes to extracellularly monitor ORN responses. Next, we present a method to extracellularly monitor PN responses using a commercial 16-channel electrode3. A similar approach using a custom-made 8-channel twisted wire tetrode is demonstrated for Kenyon cell recordings8. We provide details of our experimental setup and present representative recording traces for each of these techniques.

Multi-unit Recording Methods to Characterize Neural Activity in the Locust Schistocerca Americana Olfactory Circuits

We demonstrate variations of the extracellular multi-unit recording technique to characterize odor-evoked responses in the first three stages of the invertebrate olfactory pathway. These techniques can easily be adapted to examine ensemble activity in other neural systems as well.

Detection and interpretation of olfactory cues are critical for the survival  of many organisms. Remarkably, species across phyla have strikingly similar ...

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 ...

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 ...

Personalized Needles for Microinjections in the Rodent Brain

Microinjections have been used for a long time for the delivery of drugs or toxins within specific brain areas and, more recently, they have been used to deliver gene or cell therapy products. Unfortunately, current microinjection techniques use steel or glass needles that are suboptimal for multiple reasons: in particular, steel needles may cause tissue damage, and glass needles may bend when lowered deeply into the brain, missing the target region. In this article, we describe a protocol to prepare and use quartz needles that combine a number of useful features. These needles do not produce detectable tissue damage and, being very rigid, ensure reliable delivery in the desired brain region even when using deep coordinates. Moreover, it is possible to personalize the design of the needle by making multiple holes of the desired diameter. Multiple holes facilitate the injection of large amounts of solution within a larger area, whereas large holes facilitate the injection of cells. In addition, these quartz needles can be cleaned and re-used, such that the procedure becomes cost-effective.

We describe here a protocol for microinjection in the rodent brain that uses quartz needles. These needles do not produce detectable tissue damage and ensure reliable delivery even in deep regions. Moreover, they can be adapted to research needs by personalized designs and can be re-used.

Microinjections have been used for a long time for the delivery of drugs or toxins within specific brain areas and, more recently, they have been used to ...

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 ...