We would like to thank the members of the Montgomery and Madison labs for helpful discussion. We acknowledge the funding received from the following sources in this research: NFNZ, AMRF, Marsden Fund, HRC, and NIH.

Animal Ethics Statement:
The protocols described in this manuscript follow the animal care guidelines established by The University of Auckland and Stanford University. P7 rat pups were euthanized by rapid decapitation. Hippocampal dissection is then immediately performed as described below.
1. Organotypic Hippocampal Slice Culture
<ol>
	<li>Preparation
	<ol>
		<li>Prepare dissection Medium (used only for dissecting the brain). Combine 200 ml Minimum Essential Medium, 2 ml penicillin-streptomycin solution (10,000 units of each, in 0.85% NaCl), 5 ml HEPES buffer solution, 2 ml 1 M Tris stock solution (pH 7.2), and filter sterilize with 0.22 &#956;m filter. Carry out hippocampal dissection in ice-cold dissection medium.</li>
		<li>Cool the dissection medium. Place the dissection media in the freezer approximately 1 hr prior to beginning the dissection until the liquid is very cold. Do not allow large ice crystals to form. Store on ice until required.</li>
		<li>Prepare culture medium (used for everything except dissection of hippocampi). Combine 100 ml Minimum Essential Medium, (1x concentration, liquid) w/Hank&#8217;s salts, w/ L-glutamine, 2 ml penicillin-streptomycin solution (liquid, 10,000 units of each, in 0.85% NaCl ), 2.5 ml HEPES 1 M buffer solution, 50 ml Hank&#8217;s Balanced Salt Solution, 50 ml Horse Serum (defined, heat inactivated), and filter sterilize with 0.22 &#956;m filter.</li>
		<li>Prepare the culture dishes. Place 1 ml of culture media per 35 mm culture dish, and add a membrane insert to each dish. Put up to seven of these dishes into one 150 mm Petri dish (referred to hereafter as a &#8216;plate&#8217;). Place the plate in a CO2 incubator for at least an hour before the dissection begins so that the culture medium in the dishes attains the proper temperature and pH.</li>
	</ol>
	</li>
	<li>Dissection of the Hippocampi from Rat Pups at Postnatal Day 7 (P7)
	<ol>
		<li>Pre-sterilize all dissection tools under UV light before the procedure.</li>
		<li>After rapid decapitation, remove the brain and place into chilled medium in one dish, and then remove it to a piece of moistened filter paper for the dissection. Tease the cortex away from the midbrain using blunt smooth plastic-coated miniature spatulas, exposing the hippocampus. Cut the fornix, and then gently work the spatula underneath the hippocampus to flip it out (Figure 1A). 
		NOTE: Successful slice cultures can be prepared using animals up to P10.</li>
		<li>Trim the isolated hippocampus away from the rest of the brain. Transfer the hippocampi into a new dish containing chilled dissection media using a moistened soft paintbrush (e.g., white sable #4).</li>
		<li>Clean the underside (i.e. the flatter side) of the hippocampus of choroid plexus during dissection, as these spongy, meninge-like tissues make it difficult to separate hippocampal slices from one another later on. 
		NOTE: Leave these tissues in place if they cannot be teased away gently.</li>
		<li>Perform the entire dissection as quickly as possible without damaging the hippocampi. 
		NOTE: A careful dissection is more important than a fast one, as long as the experimental conditions are chilled. Slice the hippocampi from three rats simultaneously. The slice health is not affected as long as the total time from start to when the plated slices go into the incubator is under 30 min.</li>
	</ol>
	</li>
	<li>Slicing
	<ol>
		<li>Slice the hippocampi transversely into 400 &#956;m cross-sections using a manual tissue slicer. The method by which the slicing is conducted is not important, so long as it is not overly damaging to the tissue and produces slices of consistent thickness with readily visible laminar architecture (i.e. Ammon&#8217;s Horn is easily seen to be preserved in the tissue).</li>
		<li>Lie the hippocampi on the stage of the tissue chopper on top of a triple thickness of #2 filter paper. The hippocampi are sliced entirely without removing any slices individually (i.e. the entire hippocampus is left on the stage of the chopper until all cuts have been made; rather like loaves of sliced bread at this point; Figure 1B).</li>
		<li>Transfer the hippocampi into dissection medium using a soft paintbrush laid next to each hippocampus (white sable #2). Gently push each hippocampus to the side to break the adhesion between the hippocampus and the underlying filter paper. Roll the brush underneath each hippocampus to pick it up off the stage. Place all hippocampi into the same 35 mm Petri dish of chilled dissection medium. Separate the hippocampi into individual slices by manually agitating the dish in alternating clockwise and counterclockwise motions.</li>
		<li>Inspect the slices under a dissecting microscope and discard any which are damaged, small, or which do not possess clearly visible cell body layers. 
		NOTE: It will often be the case that not all the slices are successfully separated from their siblings in this manner. In this case, they can be separated by turning them on edge and prodding them with the tips of fine forceps. &#8220;Good&#8221; slices are then transferred to another clean Petri dish containing chilled dissection medium using a cut off and fire-polished Pasteur pipette (Figures 1C-E).</li>
	</ol>
	</li>
	<li>Storage
	<ol>
		<li>Place the individual slices onto the membrane filter inserts. Using a Pasteur pipette cut off and fire polished to give a larger bore, individually transfer slices to the culture inserts. 
		NOTE: The size of the bore created when cutting and fire-polishing the pipette is important. Too small and the slices will not easily leave the pipette for the membrane. Too large and excess fluid is placed on the surface of the membrane along with the slice. The best bore diameter is typically about half the diameter of the barrel of the pipette. When cutting, this diameter can be chosen by cutting at the proper place on the taper of the pipette.</li>
		<li>Slice Transfer
		<ol>
			<li>Fill the pipette with medium first, and then suck up a single slice with small suction forces. This keeps the slice near the opening of the pipette and prevents too much medium expulsion into the insert to place the slice.</li>
			<li>Apply a slight pressure on the bulb to form a small hanging droplet, allow the slice to settle into that droplet, and then touch that droplet to the membrane and let the slice &#8220;fall&#8221; onto the membrane. Generally three slices are placed onto each membrane insert. 
			NOTE: If the fluid droplet is too large, the droplets from the three slices tend to merge on the membrane surface, and the slices will thus all gather to the center of the membrane, making it more difficult to separate them for electrophysiological recording later.</li>
		</ol>
		</li>
		<li>Fluid Removal
		<ol>
			<li>Aspirate any excess medium from the top of the culture plate insert for all slices in that plate (3 slices per insert = 21 slices per plate), so that slices are not immersed in or surrounded by a pool of dissection medium.</li>
			<li>Perform this is with an uncut, but fire-polished Pasteur pipette. Allow the slice to settle and adhere to the membrane for a minute before pipetting. Take care not to suck the slice up into the pipette. Carry out fluid removal for the inserts that are plated first, to allow sufficient time for the slices to settle.</li>
		</ol>
		</li>
		<li>Slice Storage
		<ol>
			<li>Store slice cultures in a 5.0% CO2 incubator at 37 &#176;C. Observe the final arrangement of the cultures in Figure 1E. The individual slices rest upon a culture plate insert which has a porous membrane for a bottom, and this insert fits inside a Petri dish filled with a small amount of medium; by this means, the slices are not immersed in the medium, but have access to it only through the membrane at the bottom of the culture plate insert.</li>
			<li>Optionally, remove the slices may for study either by removing the culture plate insert or by cutting out a section of insert membrane containing a slice.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Maintenance
	<ol>
		<li>Exchange the medium in the Petri dishes the day after making cultures. Transfer the culture plate insert to a new petri dish containing 1 ml of fresh culture medium that is equilibrated in the incubator for at least an hour prior to transfer.</li>
		<li>Change the medium again in the same manner on the third day after making cultures. On the third day, transfer the cultures to an incubator set at 34 &#176;C. Change the medium twice each week (every 3-4 days).</li>
		<li>Identify healthy cultures. Select healthy cultures for paired whole cell recordings. 
		NOTE: Healthy cultures have a well-defined edge and a clearly defined pyramidal cell layer. Cultures with dark (likely necrotic) areas present or a vacuolated appearance with flattened borders are rejected. Employing these criteria, usually two-thirds of slice cultures are sufficiently healthy for recording.</li>
		<li>Utilize these cultures within a fairly small window of time. Do not used tissues that are cultured for more than two weeks since the neurons begin to display slightly epileptiform behavior which slowly worsens with time. The exact upper limit of neuronal survival is not known but it is possible to record from functioning pyramidal cells as late as 16&#160;weeks in culture.</li>
	</ol>
	</li>
</ol>
2. Paired Whole Cell Recordings
<ol>
	<li>ACSF and Intracellular Solutions
	<ol>
		<li>Prepare presynaptic electrode solution by mixing (in mM) 120 potassium gluconate, 40 HEPES, 5 MgCl2, 2 NaATP, and 0.3 NaGTP (pH 7.2 with KOH; osmolarity: 290 mOsm). Whilst the same electrode solution is used for postsynaptic neurons, cesium gluconate (120 mM) is typically used as the major salt in the postsynaptic electrode, plus 5 mM QX314. 
		NOTE: This enables stable voltage clamping at positive potentials to record postsynaptic NMDAR-mediated currents8-10,13. It also prevents the potassium-induced hyperexcitability of the presynaptic neuron by the internal solution flowing from the postsynaptic recording electrode before a giga ohm seal is made.</li>
		<li>Prepare postsynaptic electrode solution composing (in mM) 120 cesium gluconate, 40 HEPES, 5 MgCl2, 5 QX314, 2 NaATP, and 0.3 NaGTP (pH 7.2 with CsOH; osmolarity: 290 mOsm). Pull glass microelectrodes at 5-10 M&#937; resistance and fill with filtered internal solution.</li>
		<li>Prepare artificial cerebrospinal fluid (ACSF) composing (in mM) 119 NaCl, 2.4 KCl, 1.3 MgSO4, 2.4 CaCl2, 1 Na2HPO4, 26.2 NaHCO3, 11 glucose, pH 7.4, saturated with 95% O2, 5% CO2. NOTE: If AMPAR-mediated responses need to be blocked for examination of NMDAR EPSCs, include 10 &#956;M CNQX or NBQX in the ACSF.</li>
	</ol>
	</li>
	<li>Obtain Paired Whole Cell Recordings
	<ol>
		<li>To examine synaptic transmission between two individual neurons, designate one neuron as the presynaptic neuron and hold in current clamp to induce action potentials and initiate synaptic transmission.</li>
		<li>Designate the second neuron, as the postsynaptic neuron, and hold it in current or voltage clamp depending on the information required by the researcher. Obtain detailed examination of the glutamatergic currents, with AMPAR-mediated EPSCs examined at -65 mV and NMDAR-mediated EPSCs examined at +30 mV6-16 by holding the postsynaptic neuron in voltage clamp</li>
		<li>Establish a paired recording. To establish a paired recording, the presynaptic whole cell recording is always obtained first, enabling multiple sequential postsynaptic neurons to be obtained until one that is synaptically connected is obtained. If performing synaptic plasticity experiments, it is especially critical to obtain the presynaptic neuron first as the induction of LTP in the postsynaptic neuron must be initiated within 10 min of obtaining the whole cell recordings to prevent washout of cytoplasmic factors required for LTP8,19.</li>
		<li>Avoid movement-induced cell loss. A major challenge when performing paired whole cell recordings is avoiding vibration and movement-induced disruption of the first whole cell recording whilst obtaining the second recording.
		<ol>
			<li>Obtain the first whole cell recording and then move the microscope lens 10-200 &#956;m in the x-axis so that the established recording is located at the edge of the area visible on the monitor (Figure 2A).</li>
			<li>Raise the lens of the microscope ~5-10 mm while ensuring that the ACSF still maintains contact with the lens. Mount the second electrode and guide into the fluid meniscus (Figure 2B).</li>
			<li>Move the electrode in the x-y plane until it is directly under the light path through the lens but still significantly above the established recording electrode. Once the electrode is visible under the microscope, ensure the tip is at the far edge of the monitor away from the first electrode.</li>
			<li>Move the second electrode down in sequence with the microscope focus until both electrodes are in the same focal plane. It is important that the level of positive pressure is strong enough to avoid tip blockage but not too strong so as to disrupt the first whole cell recording. This translates to the positive pressure inducing movement in 2-3 neurons from the electrode when it first enters the slice.</li>
			<li>Locate a postsynaptic partner in a similar focal plane to the first presynaptic recording (Figure 2C). Recording is generally Obtain paired recordings between CA3 pyramidal neurons the second whole cell from a 0-200 mm neuron of the first recording (Figures 2C,D). 
			NOTE: Synaptic transmission between neuronal pairs are stable over time periods of up to 3-4 hr when using these standard whole cell recording techniques.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Synaptic plasticity: Once Upon obtaining a successful synaptically-connected pyramidal cell pair (Figures 3A,B), examine the characteristics of synaptic transmission and plasticity.
	<ol>
		<li>LTP induction.
		<ol>
			<li>Induce LTP by pairing presynaptic action potentials (1 Hz) with postsynaptic depolarization to -10 to 0 mV for 1 min (Figure 3C)7-9,12-14. Initiate pairing within 10 min of break-in to the postsynaptic neuron. 
			NOTE: LTP can also be induced with both neurons held in current clamp by pairing presynaptic and postsynaptic action potentials at 1 Hz for 1 min, with postsynaptic action potentials elicited 10 msec following injection of current into the presynaptic neuron8.</li>
		</ol>
		</li>
		<li>Further induce LTD by low frequency stimulation (LFS) at 1 Hz combined with a slight depolarization of the postsynaptic neuron to -55 mV for 5-10 min (Figure 3C)9.</li>
	</ol>
	</li>
</ol>

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V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Presynaptic+FMR1+genotype+influences+the+degree+of+synaptic+connectivity+in+a+mosaic+mouse+model+of+fragile+X+syndrome.">Presynaptic FMR1 genotype influences the degree of synaptic connectivity in a mosaic mouse model of fragile X syndrome.</a> Journal of Neuroscience. 27 (15), 4014-4018 (2007).</li><li>Hanson, J. E., Blank, M., Valenzuela, R. A., Garner, C. C., Madison, D. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+functional+nature+of+synaptic+circuitry+is+altered+in+area+CA3+of+the+hippocampus+in+a+mouse+model+of+Down's+syndrome.">The functional nature of synaptic circuitry is altered in area CA3 of the hippocampus in a mouse model of Down's syndrome.</a> Journal of Physiology. 579, 53-67 (2007).</li></ol>

The authors have nothing to disclose

Here we have described the requirements for establishing successful paired whole cell recordings in organotypic hippocampal slice cultures. Paired recordings can also be performed in multiple preparations, including acute slices and dissociated culture systems26,27. While the focus here has been on the induction of longer forms of synaptic plasticity (namely LTP and LTD), it is important to highlight that paired whole cell recordings in organotypic, acute slice and dissociated cell preparations have provided i...

Glutamate receptors mediate the majority of excitatory synaptic transmission at central nervous system synapses. The two major subtypes of ionotropic glutamate receptors localized at the spine head of the postsynaptic membrane are N-methyl-D-aspartate (NMDA) and &#945;-amino-3-hydroxy-5-methylisoxazole-4-proprionic acid (AMPA) receptors. At resting membrane potentials, AMPA receptors carry most of the postsynaptic current during synaptic transmission. In the hippocampus, the NMDA receptor plays a key role in triggering changes in the number of AMPA receptors in the postsynaptic membrane: by acting as a &#8221;coincidence detector&#8221;1 to initiate changes in synaptic strength1, the NMDA receptor participates in the synaptic mechanisms that are thought to underpin learning and memory at a subcellular level. In response to depolarization of the postsynaptic neuron in parallel with presynaptic transmitter release, calcium enters via the NMDA receptor to initiate AMPA receptor insertion or removal2. These receptor dynamics underlie synapse plasticity: an increase in synaptic strength is long-term potentiation2,3 (LTP), while a decrease in synaptic strength is long-term depression4 (LTD). Therefore AMPA receptor movement is thought to be responsible for synaptic plasticity expression, while NMDA receptors are thought to control its induction.
Determining the precise mechanisms underlying synaptic transmission and plasticity requires studying small populations of synapses, ideally single synapses. While some synapses are highly suited for study at this level, e.g.,&#160;the Calyx of Held5, for most synaptic populations this is extremely difficult due to the small and diffuse nature of the synaptic connections. Two major electrophysiology techniques have been developed to examine single synaptic connections: The first is minimal stimulation, where one presynaptic fiber is presumed stimulated extracellularly. The second technique is paired recordings, where two simultaneous whole cell recordings from synaptically connected neurons is performed. A major advantage of minimal stimulation is that it is rapid and relatively simple to perform, involving placement of an extracellular stimulating electrode into the axonal tract while simultaneously recording from a postsynaptic neuron. The primary concern when using this technique is that reliable stimulation of a single cell can rarely be guaranteed trial after trial.
Over the past fifteen years we have routinely used paired whole-cell recordings from two synaptically connected pyramidal neurons6-17. The major advantage of this technique is that only one presynaptic neuron is consistently and reliably stimulated. It also allows not only electrophysiological characterization but also pharmacological manipulation of the presynaptic neuron6,18. However, the probability of synaptic connectivity between neurons is low, making connected pairs difficult to obtain19. The use of organotypic brain slice cultures circumvents this obstacle as synaptic connectivity can re-establish in vitro and moreover the nature of the resulting connectivity is similar to that in native brain tissue20. In addition, organotypic cultures express LTP, LTD7-10,12-15,21 and additional forms of short-term synaptic plasticity including paired-pulse facilitation (PPF) and depression (PPD)6,22,23, enabling plasticity mechanisms to be studied in pairs of neurons. Here we describe the detailed methodology involved in successfully attaining paired recordings in this in vitro system. This information can readily be adapted to other experimental systems, including acute slices and other brain regions.

<table><tbody><tr><td>Minimum Essential Medium</td><td></td><td></td><td>Stable motorized micromanipulators&amp;nbsp;</td></tr><tr><td>Penicillin-Streptomycin solution&amp;nbsp;</td><td></td><td></td><td>Shallow tissue bath</td></tr><tr><td>HEPES buffer solution</td><td></td><td></td><td>DIC camera</td></tr><tr><td>1 M Tris stock solution</td><td></td><td></td><td>Amplifier</td></tr><tr><td>Hank&amp;rsquo;s Balanced Salt Solution</td><td></td><td></td><td>Computer</td></tr><tr><td>Horse Serum&amp;nbsp;</td><td></td><td></td><td>Vibration isolation table</td></tr><tr><td>Plastic-coated miniature spatulas</td><td></td><td></td><td>Upright microscope</td></tr><tr><td>Soft paintbrush</td><td></td><td></td><td>Data acquistion and analysis software</td></tr><tr><td>Manual tissue chopper</td><td></td><td></td><td>Electrode puller</td></tr><tr><td>#2 Filter paper</td><td></td><td></td><td>Faraday cage</td></tr><tr><td>#5 Forceps</td><td></td><td></td><td></td></tr><tr><td>Membrane inserts</td><td></td><td></td><td></td></tr><tr><td>CO&lt;sub&gt;2&lt;/sub&gt; incubator</td><td></td><td></td><td></td></tr><tr><td>Dissection hood</td><td></td><td></td><td></td></tr><tr><td>Class II hood</td><td></td><td></td><td></td></tr></tbody></table>

paired whole cell recordings in organotypic hippocampal slices

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct electrophysiological characterization of the synaptic connections between individual neurons, but also pharmacological manipulation of either the presynaptic or the postsynaptic neuron. When carried out in organotypic hippocampal slice cultures, the probability that two neurons are synaptically connected is significantly increased. This preparation readily enables identification of cell types, and the neurons maintain their morphology and properties of synaptic function similar to that in native brain tissue. A major advantage of paired whole cell recordings is the highly precise information it can provide on the properties of synaptic transmission and plasticity that are not possible with other more crude techniques utilizing extracellular axonal stimulation. Paired whole cell recordings are often perceived as too challenging to perform. While there are challenging aspects to this technique, paired recordings can be performed by anyone trained in whole cell patch clamping provided specific hardware and methodological criteria are followed. The probability of attaining synaptically connected paired recordings significantly increases with healthy organotypic slices and stable micromanipulation allowing independent attainment of pre- and postsynaptic whole cell recordings. While CA3-CA3 pyramidal cell pairs are most widely used in the organotypic slice hippocampal preparation, this technique has also been successful in CA3-CA1 pairs and can be adapted to any neurons that are synaptically connected in the same slice preparation. In this manuscript we provide the detailed methodology and requirements for establishing this technique in any laboratory equipped for electrophysiology.

Synaptic connectivity is evident by stimulating the presynaptic neuron to fire an action potential by passing a depolarizing current pulse (typically 20-50 pA for 20 msec) via the recording electrode. The postsynaptic current trace is then examined for the presence of a monosynaptic EPSC evoked at short (&#60;5 msec) and consistent latencies after the peak of the presynaptic action potential (Figure 3A). In most experiments multiple postsynaptic neurons are tested before a synaptically-connected pair can...

Watch this Scientific Journal Video about Paired Whole Cell Recordings in Organotypic Hippocampal Slices at JoVE.com

Paired Whole Cell Recordings in Organotypic Hippocampal Slices

Pair recordings are simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling precise electrophysiological and pharmacological characterization of the synapses between individual neurons. Here we describe the detailed methodology and requirements for establishing this technique in organotypic hippocampal slice cultures in any laboratory equipped for electrophysiology.

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct ...

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14.6K Views.  University of Auckland. Pair recordings are simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling precise electrophysiological and pharmacological characterization of the synapses between individual neurons. Here we describe the detailed methodology and requirements for establishing this technique in organotypic hippocampal slice cultures in any laboratory equipped for electrophysiology.

Video: Paired Whole Cell Recordings in Organotypic Hippocampal Slices

Whole Cell Recording from an Organotypic Slice Preparation of Neocortex

We have been studying the expression and functional roles of voltage-gated potassium channels in pyramidal neurons from rat neocortex. Because of the lack of specific pharmacological agents for these channels, we have taken a genetic approach to manipulating channel expression. We use an organotypic culture preparation (16) in order to maintain cell morphology and the laminar pattern of cortex. We typically isolate acute neocortical slices at postnatal days 8-10 and maintain the slices in culture for 3-7 days. This allows us to study neurons at a similar age to those in our work with acute slices and minimizes the development of exuberant excitatory connections in the slice. We record from visually-identified pyramidal neurons in layers II/III or V using infrared illumination (IR-) and differential interference contrast microscopy (DIC) with whole cell patch clamp in current- or voltage-clamp. We use biolistic (Gene gun) transfection of wild type or mutant potassium channel DNA to manipulate expression of the channels to study their function. The transfected cells are easily identified by epifluorescence microscopy after co-transfection with cDNA for green fluorescent protein (GFP). We compare recordings of transfected cells to adjacent, untransfected neurons in the same layer from the same slice.

This is a protocol to prepare and maintain a neocortical slice preparation in organotypic culture for the purpose of making electrical recordings from pyramidal neurons.

We have been studying the expression and functional roles of voltage-gated potassium channels in pyramidal neurons from rat neocortex.  Because of the ...

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 ...

Simultaneous Whole-cell Recordings from Photoreceptors and Second-order Neurons in an Amphibian Retinal Slice Preparation

One of the central tasks in retinal neuroscience is to understand the circuitry of retinal neurons and how those connections are responsible for shaping the signals transmitted to the brain. Photons are detected in the retina by rod and cone photoreceptors, which convert that energy into an electrical signal, transmitting it to other retinal neurons, where it is processed and communicated to central targets in the brain via the optic nerve. Important early insights into retinal circuitry and visual processing came from the histological studies of Cajal1,2 and, later, from electrophysiological recordings of the spiking activity of retinal ganglion cells - the output cells of the retina3,4.
A detailed understanding of visual processing in the retina requires an understanding of the signaling at each step in the pathway from photoreceptor to retinal ganglion cell. However, many retinal cell types are buried deep in the tissue and therefore relatively inaccessible for electrophysiological recording. This limitation can be overcome by working with vertical slices, in which cells residing within each of the retinal layers are clearly visible and accessible for electrophysiological recording.
Here, we describe a method for making vertical sections of retinas from larval tiger salamanders (Ambystoma tigrinum). While this preparation was originally developed for recordings with sharp microelectrodes5,6, we describe a method for dual whole-cell voltage clamp recordings from photoreceptors and second-order horizontal and bipolar cells in which we manipulate the photoreceptor's membrane potential while simultaneously recording post-synaptic responses in horizontal or bipolar cells. The photoreceptors of the tiger salamander are considerably larger than those of mammalian species, making this an ideal preparation in which to undertake this technically challenging experimental approach. These experiments are described with an eye toward probing the signaling properties of the synaptic ribbon - a specialized synaptic structure found in a only a handful of neurons, including rod and cone photoreceptors, that is well suited for maintaining a high rate of tonic neurotransmitter release7,8 - and how it contributes to the unique signaling properties of this first retinal synapse.

We describe the preparation of thin retinal slices from aquatic tiger salamanders (Ambystoma tigrinum) and explain how we use these slices to study synaptic processing in the retina by obtaining dual whole-cell voltage clamp recordings from photoreceptors and second-order horizontal and bipolar cells.

One of the central tasks in retinal neuroscience  is to understand the circuitry of retinal neurons and how those connections are  responsible for shaping ...

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 ...

Whole-cell Patch-clamp Recordings in Brain Slices

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the neuron. In this configuration, the micropipette is in tight contact with the cell membrane, which prevents current leakage and thereby provides more accurate ionic current measurements than the previously used intracellular sharp electrode recording method. Classically, whole-cell recording can be performed on neurons in various types of preparations, including cell culture models, dissociated neurons, neurons in brain slices, and in intact anesthetized or awake animals. In summary, this technique has immensely contributed to the understanding of passive and active biophysical properties of excitable cells. A major advantage of this technique is that it provides information on how specific manipulations (e.g., pharmacological, experimenter-induced plasticity) may alter specific neuronal functions or channels in real-time. Additionally, significant opening of the plasma membrane allows the internal pipette solution to freely diffuse into the cytoplasm, providing means for introducing drugs, e.g., agonists or antagonists of specific intracellular proteins, and manipulating these targets without altering their functions in neighboring cells. This article will focus on whole-cell recording performed on neurons in brain slices,&#160;a preparation that has the advantage of recording neurons in relatively well preserved brain circuits, i.e., in a physiologically relevant context. In particular,&#160;when combined with appropriate pharmacology, this technique is a powerful tool allowing identification of specific neuroadaptations that occurred following any type of experiences, such as learning, exposure to drugs of abuse, and stress. In summary, whole-cell patch-clamp recordings in brain slices provide means to measure in ex vivo preparation long-lasting changes in neuronal functions that have developed in intact awake animals.

This protocol describes basic procedural steps for performing whole-cell patch-clamp recordings. This technique allows the study of&#160;the electrical behavior of neurons, and when performed in brain slices, allows the assessment of various neuronal functions from neurons that are still integrated in relatively well preserved brain circuits.

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the ...

A Model of Epileptogenesis in Rhinal Cortex-Hippocampus Organotypic Slice Cultures

Organotypic slice cultures have been widely used to model brain disorders and are considered excellent platforms for evaluating a drug&#8217;s neuroprotective and therapeutic potential. Organotypic slices are prepared from explanted tissue and represent a complex multicellular ex vivo environment. They preserve the three-dimensional cytoarchitecture and local environment of brain cells, maintain the neuronal connectivity and the neuron-glia reciprocal interaction. Hippocampal organotypic slices are considered suitable to explore the basic mechanisms of epileptogenesis, but clinical research and animal models of epilepsy have suggested that the rhinal cortex, composed of perirhinal and entorhinal cortices, play a relevant role in seizure generation.
Here, we describe the preparation of rhinal cortex-hippocampus organotypic slices. Recordings of spontaneous activity from the CA3 area under perfusion with complete growth medium, at physiological temperature and in the absence of pharmacological manipulations, showed that these slices depict evolving epileptic-like events throughout time in culture. Increased cell death, through propidium iodide uptake assay, and gliosis, assessed with fluorescence-coupled immunohistochemistry, was also observed. The experimental approach presented highlights the value of rhinal cortex-hippocampus organotypic slice cultures as a platform to study the dynamics and progression of epileptogenesis and to screen potential therapeutic targets for this brain pathology.

Here, we describe the preparation of rhinal cortex-hippocampus organotypic slices. Under a gradual and controlled deprivation of serum, these slices depict evolving epileptic-like events and can be considered an ex vivo model of epileptogenesis. This system represents an excellent tool for monitoring the dynamics of spontaneous activity, as well as for assessing the progression of neuroinflammatory features throughout the course of epileptogenesis.

Organotypic slice cultures have been widely used to model brain disorders and are considered excellent platforms for evaluating a drug&#8217;s ...

Preparing Acute Brain Slices from the Dorsal Pole of the Hippocampus from Adult Rodents

Whole-cell patch-clamp recordings from acute rodent brain slices are a mainstay of modern neurophysiological research, allowing precise measurement of cellular and synaptic properties. Nevertheless, there is an ever increasing need to perform correlated analyses between different experimental modes in addition to slice electrophysiology, for example: immunohistochemistry, molecular biology, in vivo imaging or electrophysiological recording; to answer evermore complex questions of brain function. However, making meaningful conclusions from these various experimental approaches is not straightforward, as even within relatively well described brain structures, a high degree of sub-regional variation of cellular function exists. Nowhere is this better exemplified than in the CA1 of the hippocampus, which has well-defined dorso-ventral properties, based on cellular and molecular properties. Nevertheless, many published studies examine protein expression patterns or behaviorally correlated in vivo activity in the dorsal extent of the hippocampus; and explain findings mechanistically with cellular electrophysiology from the ventro-medial region. This is further confounded by the fact that many acute slice electrophysiological experiments are performed in juvenile animals, when other experimental modes are performed in more mature animals. To address these issues, this method incorporates transcardial perfusion of mature (&#62;60 day old rodents) with artificial cerebrospinal fluid followed by preparation of modified coronal slices including the septal pole of the dorsal hippocampus to record from CA1 pyramidal cells. This process leads to the generation of healthy acute slices of dorsal hippocampus allowing for slice-based cellular electrophysiological interrogation matched to other measures.

The purpose of this protocol is to describe a method to produce slices of the dorsal hippocampus for electrophysiological examination. This procedure employs perfusion with chilled ACSF prior to slice preparation with a near-coronal slicing angle which allows for preservation of healthy principal neurons.

Whole-cell patch-clamp recordings from acute rodent brain slices are a mainstay of modern neurophysiological research, allowing precise measurement of ...

Acute Mouse Brain Slicing to Investigate Spontaneous Hippocampal Network Activity

Acute rodent brain slicing offers a tractable experimental approach to gain insight into the organization and function of neural circuits with single-cell resolution using electrophysiology, microscopy, and pharmacology. However, a major consideration in the design of in vitro experiments is the extent to which different slice preparations recapitulate naturalistic patterns of neural activity as observed in vivo. In the intact brain, the hippocampal network generates highly synchronized population activity reflective of the behavioral state of the animal, as exemplified by the sharp-wave ripple complexes (SWRs) that occur during waking consummatory states or non-REM sleep. SWRs and other forms of network activity can emerge spontaneously in isolated hippocampal slices under appropriate conditions. In order to apply the powerful brain slice toolkit to the investigation of hippocampal network activity, it is necessary to utilize an approach that optimizes tissue health and the preservation of functional connectivity within the hippocampal network. Mice are transcardially perfused with cold sucrose-based artificial cerebrospinal fluid. Horizontal slices containing the hippocampus are cut at a thickness of 450 &#956;m to preserve synaptic connectivity. Slices recover in an interface-style chamber and are transferred to a submerged chamber for recordings. The recording chamber is designed for dual surface superfusion of artificial cerebrospinal fluid at a high flow rate to improve oxygenation of the slice. This protocol yields healthy tissue suitable for the investigation of complex and spontaneous network activity in vitro.

This protocol describes the preparation of horizontal hippocampal-entorhinal cortex (HEC) slices from mice exhibiting spontaneous sharp-wave ripple activity. Slices are incubated in a simplified interface holding chamber and recordings are performed under submerged conditions with fast-flowing artificial cerebrospinal fluid to promote tissue oxygenation and the spontaneous emergence of network-level activity.

Acute rodent brain slicing offers a tractable experimental approach to gain insight into the organization and function of neural circuits with single-cell ...

Preparation of Acute Slices from Dorsal Hippocampus for Whole-Cell Recording and Neuronal Reconstruction in the Dentate Gyrus of Adult Mice

Although the general architecture of the hippocampus is similar along its longitudinal axis, recent studies have revealed prominent differences in molecular, anatomical and functional criteria suggesting a division into different sub-circuits along its rostro-caudal extent. Owing to differential connectivity and function the most fundamental distinction is made between the dorsal and the ventral hippocampus, which are preferentially involved in spatial and emotional processing, respectively. Accordingly, in vivo work regarding spatial memory formation has focused on the dorsal hippocampus.
In contrast, electro-physiological in vitro recordings have been preferentially performed on intermediate-ventral hippocampus, largely motivated by factors like slice viability and circuit integrity. To allow for direct correlation of in vivo data on spatial processing with in vitro data we have adapted previous sectioning methods to obtain highly viable transverse brain slices from the dorsal-intermediate hippocampus for long-term recordings of principal cells and interneurons in the dentate gyrus. As spatial behavior is routinely analyzed in adult mice, we have combined this transversal slicing procedure with the use of protective solutions to enhance viability of brain tissue from mature animals. We use this approach for mice of about 3 months of age. The method offers a good alternative to the coronal preparation which is frequently used for in vitro studies on dorsal hippocampus. We compare these two preparations in terms of quality of recordings and preservation of morphological features of recorded neurons.

We present a protocol to prepare acute-slices from the dorsal-intermediate hippocampus of mice. We compare this transversal preparation with coronal slicing in terms of quality of recordings and preservation of morphological features of recorded neurons.

Although the general architecture of the hippocampus is similar along its longitudinal axis, recent studies have revealed prominent differences in ...

Photostimulation and Whole-Cell Patch Clamp Recordings of Neurons in Mouse Hippocampal Slices

Source: Richevaux, L., et al. <a href="https://app.jove.com/v/59534">In Vivo Intracerebral Stereotaxic Injections for Optogenetic Stimulation of Long-Range Inputs in Mouse Brain Slices.</a> J. Vis. Exp. (2019)
The video demonstrates a protocol to study synaptic interactions and neuronal responses in a transfected mouse hippocampal brain slice. It involves illuminating long-range sensory neurons to trigger action potentials and recording the resulting excitatory post-synaptic potentials in the recorded neuron using the whole-cell patch-clamp technique.

All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.
1. Solutions for acute slice ...