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 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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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 ...

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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

Automatic Detection of Highly Organized Theta Oscillations in the Murine EEG

Theta activity is generated in the septohippocampal system and can be recorded using deep intrahippocampal electrodes and implantable electroencephalography (EEG) radiotelemetry or tether system approaches. Pharmacologically, hippocampal theta is heterogeneous (see dualistic theory) and can be differentiated into type I and type II theta. These individual EEG subtypes are related to specific cognitive and behavioral states, such as arousal, exploration, learning and memory, higher integrative functions, etc. In neurodegenerative diseases such as Alzheimer&#39;s, structural and functional alterations of the septohippocampal system can result in impaired theta activity/oscillations. A standard quantitative analysis of the hippocampal EEG includes a Fast-Fourier-Transformation (FFT)-based frequency analysis. However, this procedure does not provide details about theta activity in general and highly-organized theta oscillations in particular. In order to obtain detailed information on highly-organized theta oscillations in the hippocampus, we have developed a new analytical approach. This approach allows for time- and cost-effective quantification of the duration of highly-organized theta oscillations and their frequency characteristics.

Theta activity in the hippocampus is related to specific cognitive and behavioral stages. Here, we describe an analytical method to detect highly-organized theta oscillations within the hippocampus using a time-frequency (i.e., wavelet analysis)-based approach.

Theta activity is generated in the septohippocampal system and can be recorded using deep intrahippocampal electrodes and implantable ...

Behavior

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 ...

Improved Preparation and Preservation of Hippocampal Mouse Slices for a Very Stable and Reproducible Recording of Long-term Potentiation

Long-term potentiation (LTP) is a type of synaptic plasticity characterized by an increase in synaptic strength and believed to be involved in memory encoding. LTP elicited in the CA1 region of acute hippocampal slices has been extensively studied. However the molecular mechanisms underlying the maintenance phase of this phenomenon are still poorly understood. This could be partly due to the various experimental conditions used by different laboratories. Indeed, the maintenance phase of LTP is strongly dependent on external parameters like oxygenation, temperature and humidity. It is also dependent on internal parameters like orientation of the slicing plane and slice viability after dissection.
The optimization of all these parameters enables the induction of a very reproducible and very stable long-term potentiation. This methodology offers the possibility to further explore the molecular mechanisms involved in the stable increase in synaptic strength in hippocampal slices. It also highlights the importance of experimental conditions in in vitro investigation of neurophysiological phenomena.

This paper presents a complete methodology to prepare and preserve in vitro acute hippocampal slices from adult mice. This protocol allows the recording of very stable long-lasting long-term potentiation (LTP) for more than 8 hours with a success rate of 95%.

Long-term potentiation (LTP) is a type of synaptic plasticity characterized by an increase in synaptic strength and believed to be involved in memory ...

Whole-cell Patch-clamp Recordings from Morphologically- and Neurochemically-identified Hippocampal Interneurons

GABAergic inhibitory interneurons play a central role within neuronal circuits of the brain. Interneurons comprise a small subset of the neuronal population (10-20%), but show a high level of physiological, morphological, and neurochemical heterogeneity, reflecting their diverse functions. Therefore, investigation of interneurons provides important insights into the organization principles and function of neuronal circuits. This, however, requires an integrated physiological and neuroanatomical approach for the selection and identification of individual interneuron types. Whole-cell patch-clamp recording from acute brain slices of transgenic animals, expressing fluorescent proteins under the promoters of interneuron-specific markers, provides an efficient method to target and electrophysiologically characterize intrinsic and synaptic properties of specific interneuron types. Combined with intracellular dye labeling, this approach can be extended with post-hoc morphological and immunocytochemical analysis, enabling systematic identification of recorded neurons. These methods can be tailored to suit a broad range of scientific questions regarding functional properties of diverse types of cortical neurons.

Cortical networks are controlled by a small, but diverse set of inhibitory interneurons. Functional investigation of interneurons therefore requires targeted recording and rigorous identification. Described here is a combined approach involving whole-cell recordings from single or synaptically-coupled pairs of neurons with intracellular labeling, post-hoc morphological and immunocytochemical analysis.

GABAergic inhibitory interneurons play a central role within neuronal circuits of the brain. Interneurons comprise a small subset of the neuronal ...

Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools to selectively manipulate brain rhythms. Here we describe an approach combining projection-specific optogenetics with extracellular electrophysiology for high-fidelity control of hippocampal theta oscillations (5-10 Hz) in behaving mice. The specificity of the optogenetic entrainment is achieved by targeting channelrhodopsin-2 (ChR2) to the GABAergic population of medial septal cells, crucially involved in the generation of hippocampal theta oscillations, and a local synchronized activation of a subset of inhibitory septal afferents in the hippocampus. The efficacy of the optogenetic rhythm control is verified by a simultaneous monitoring of the local field potential (LFP) across lamina of the CA1 area and/or of neuronal discharge. Using this readily implementable preparation we show efficacy of various optogenetic stimulation protocols for induction of theta oscillations and for the manipulation of their frequency and regularity. Finally, a combination of the theta rhythm control with projection-specific inhibition addresses the readout of particular aspects of the hippocampal synchronization by efferent regions.

We describe the use of optogenetics and electrophysiological recordings for selective manipulations of hippocampal theta oscillations (5-10 Hz) in behaving mice. The efficacy of the rhythm entrainment is monitored using local field potential. A combination of opto- and pharmacogenetic inhibition addresses the efferent readout of hippocampal synchronization.

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools ...

Recording and Modulation of Epileptiform Activity in Rodent Brain Slices Coupled to Microelectrode Arrays

Temporal lobe epilepsy (TLE) is the most common partial complex epileptic syndrome and the least responsive to medications. Deep brain stimulation (DBS) is a promising approach when pharmacological treatment fails or neurosurgery is not recommended. Acute brain slices coupled to microelectrode arrays (MEAs) represent a valuable tool to study neuronal network interactions and their modulation by electrical stimulation. As compared to conventional extracellular recording techniques, they provide the added advantages of a greater number of observation points and a known inter-electrode distance, which allow studying the propagation path and speed of electrophysiological signals. However, tissue oxygenation may be greatly impaired during MEA recording, requiring a high perfusion rate, which comes at the cost of decreased signal-to-noise ratio and higher oscillations in the experimental temperature. Electrical stimulation further stresses the brain tissue, making it difficult to pursue prolonged recording/stimulation epochs. Moreover, electrical modulation of brain slice activity needs to target specific structures/pathways within the brain slice, requiring that electrode mapping be easily and quickly performed live during the experiment. Here, we illustrate how to perform the recording and electrical modulation of 4-aminopyridine (4AP)-induced epileptiform activity in rodent brain slices using planar MEAs. We show that the brain tissue obtained from mice outperforms rat brain tissue and is thus better suited for MEA experiments. This protocol guarantees the generation and maintenance of a stable epileptiform pattern that faithfully reproduces the electrophysiological features observed with conventional field potential recording, persists for several hours, and outlasts sustained electrical stimulation for prolonged epochs. Tissue viability throughout the experiment is achieved thanks to the use of a small-volume custom recording chamber allowing for laminar flow and quick solution exchange even at low (1 mL/min) perfusion rates. Quick MEA mapping for real-time monitoring and selection of stimulating electrodes is performed by a custom graphic user interface (GUI).

We illustrate how to perform recording and electrical modulation of 4-aminopyridine-induced epileptiform activity in rodent brain slices using microelectrode arrays. A custom recording chamber maintains tissue viability throughout prolonged experimental sessions. Live electrode mapping and selection of stimulating pairs are performed by a custom graphical user interface.

Temporal lobe epilepsy (TLE) is the most common partial complex epileptic syndrome and the least responsive to medications. Deep brain stimulation (DBS) ...

Recording Spatially Restricted Oscillations in the Hippocampus of Behaving Mice

The local field potential (LFP) emerges from ion movements across neural membranes. Since the voltage recorded by LFP electrodes reflects the summed electrical field of a large volume of brain tissue, extracting information about local activity is challenging. Studying neuronal microcircuits, however, requires a reliable distinction between truly local events and volume-conducted signals originating in distant brain areas. Current source density (CSD) analysis offers a solution for this problem by providing information about current sinks and sources in the vicinity of the electrodes. In brain areas with laminar cytoarchitecture such as the hippocampus, one-dimensional CSD can be obtained by estimating the second spatial derivative of the LFP. Here, we describe a method to record multilaminar LFPs using linear silicon probes implanted into the dorsal hippocampus. CSD traces are computed along individual shanks of the probe. This protocol thus describes a procedure to resolve spatially restricted neuronal network oscillations in the hippocampus of freely moving mice.

This protocol describes the recording of local field potentials with multi-shank linear silicon probes. Conversion of the signals using current source density analysis allows the reconstruction of local electrical activity in the mouse hippocampus. With this technique, spatially restricted brain oscillations can be studied in freely moving mice.

The local field potential (LFP) emerges from ion movements across neural membranes. Since the voltage recorded by LFP electrodes reflects the summed ...

Selective Manipulation of Theta Oscillations in a Mouse Model

Source: Bender, F. et al., <a href="https://app.jove.com/v/57349">Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice.</a> J. Vis. Exp (2018)
This video demonstrates a protocol to modulate theta oscillations in a genetically engineered mouse model with light-sensitive ion channels in its GABAergic medial septum neurons. By delivering light pulses to the hippocampus, the protocol aims to synchronize neuronal activity and generate theta oscillations, which are recorded as local field potentials.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
PV-Cre ...

JoVE Encyclopedia of Experiments

Neurophysiology

Stimulation and Modulation Techniques

Optogenetic Control of Hippocampal Network Dynamics

Source: Manseau, F., et al. <a href="https://app.jove.com/v/55851">Tuning in the Hippocampal Theta Band In Vitro: Methodologies for Recording from the Isolated Rodent Septohippocampal Circuit.</a> J. Vis. Exp. (2017)
This video demonstrates the method to use optogenetics to modulate hippocampal network dynamics by activating genetically modified light-sensitive interneurons in a mouse brain slice, enhancing theta oscillation synchronization and studying neural circuit behavior.

All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.

	Optogenetic control in the ...

In Vitro Recording of Theta Oscillations in the Mouse Hippocampus

Source: Manseau, F., et.al. <a href="https://app.jove.com/v/55851">Tuning in the Hippocampal Theta Band In Vitro: Methodologies for Recording from the Isolated Rodent Septohippocampal Circuit.</a> J. Vis. Exp. (2017).
This video demonstrates the recording of rhythmic neuronal network activity, also known as theta oscillations, in an isolated hippocampus. Electrodes are inserted into various layers and spatial positions to detect and analyze theta oscillations. This shows the unique neuronal compositions and the connectivity of the neuronal network across different layers of the hippocampus.

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

Summary

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Chapters in this video

Automatic Detection of Highly Organized Theta Oscillations in the Murine EEG

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

Improved Preparation and Preservation of Hippocampal Mouse Slices for a Very Stable and Reproducible Recording of Long-term Potentiation

Whole-cell Patch-clamp Recordings from Morphologically- and Neurochemically-identified Hippocampal Interneurons

Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice

Recording and Modulation of Epileptiform Activity in Rodent Brain Slices Coupled to Microelectrode Arrays

Recording Spatially Restricted Oscillations in the Hippocampus of Behaving Mice

Selective Manipulation of Theta Oscillations in a Mouse Model

Optogenetic Control of Hippocampal Network Dynamics

In Vitro Recording of Theta Oscillations in the Mouse Hippocampus