The author would like to thank Steve Siegelbaum for support. Funding is provided by 5R01NS106983-02 as well as 1 F31 NS113466-01.

All methods described here have been approved by the Institutional Animal Care and Use Committee at Columbia University (AC-AAAU9451).
1. Prepare solutions
<ol>
	<li>Prepare sucrose cutting solution for slicing as described in Table 1. 
	NOTE: After preparing 1 L of sucrose solution, freeze a small amount (approximately 100&#8211;200 mL) in an ice tray. These frozen sucrose ice cubes will be blended into an icy slurry (see step 4.3).</li>
	<li>Prepare artificial cerebrospinal fluid (ACSF) for recording as described in Table 2.</li>
	<li>Prepare 1 M of NaCl for stimulation pipettes by dissolving 0.05844 g of NaCl in 1 mL of purified water that has been filtered to remove trace metals and other impurities. 
	NOTE: Sucrose cutting solution and ACSF should be prepared fresh for each experiment.</li>
</ol>
2. Prepare agar ramp
<ol>
	<li>Prepare 4% agar (e.g., 2 g of agar in 50 mL of water) by dissolving agar powder into purified water that has been filtered to remove trace metals and other impurities. Heat the mixture in a microwave until it just begins to boil (approximately 30&#8211;60 s). Pour the molten agar into a mold and allow it to cool and solidify at 4 &#176;C in a fridge at an approximately 12&#176; incline. 
	NOTE: For a mold, one can use any glass or plastic container set at an angle, with enough molten agar poured in to form a ramp of 4 cm in length and approximately 0.8 cm in height.</li>
</ol>
3. Stage the slicing area
<ol>
	<li>Fill a 400 mL beaker containing a slice recovery chamber with approximately 250 mL of ACSF; place it in a water bath warmed to 32 &#176;C and begin bubbling with carbogen (95% oxygen gas/5% carbon dioxide gas). 
	NOTE: Fill the recovery beaker with just enough fluid to place the nylon mesh of the holding chamber at the very surface of the solution, so that slices will be held at the interface of the warm solution mixture and the air (Figure 1). Place small pieces of lens paper on the nylon mesh. The slices will lay on top of the lens paper, which can later be used to transfer slices individually to the recording chamber (see step 6.6).</li>
	<li>Place a flask with the sucrose solution into an ice bucket to chill and begin bubbling with carbogen. 
	NOTE: The recovery beaker and sucrose solution should be warmed and chilled, respectively, with carbogen bubbling for at least 30 mins before the mouse is placed in the isoflurane chamber.</li>
	<li>Pull out the tray of premade, frozen sucrose solution ice cubes from the freezer and place on the bench to partially thaw.</li>
	<li>Place one half of a clean double-edged razor blade into the microtome and calibrate if necessary. Set aside the other half of the razor blade for use during the dissection.</li>
	<li>Run a carbogen line and temperature probe into the slicing chamber and surround with ice to chill the chamber.</li>
	<li>Prepare a clean bench or table covered with two disposable absorbent pads. Lay out all dissection tools and three pieces of lab tape on the left pad, where the transcardial perfusion will be performed. 
	NOTE: Dissection tools include small Bonn scissors, dissector scissors, large spatula/spoon, microspatula, scalpel with new blade, spatula, sharp/blunt surgical scissors, large tissue forceps, and small tissue forceps (see Table of Materials).</li>
	<li>On the other absorbent pad to the right of the perfusion area, place a circular piece of filter paper into the lid of a 100 mm diameter glass Petri dish. Place the bottom of the Petri dish next to the lid and place a carbogen line into both pieces of the dish.</li>
	<li>Use a razor blade or scalpel to cut a small ramp of agar (approximately 4 cm along the angled surface, 0.8 cm in height, 2 cm in width). Cut off a piece of the ramp from the tall end and reverse it to create a backing block of agar. Use cyanoacrylate adhesive to affix the agar ramp and backing block to the slicing platform and set aside. 
	NOTE: The backing block of agar helps stabilize the brain during slicing and provides a surface on which to dissect away unnecessary tissues from each slice (Figure 1).</li>
</ol>
4. Transcardial perfusion
<ol>
	<li>Suspend a 60 mL capacity syringe as a reservoir for sucrose cutting solution approximately 18 inches above the benchtop (e.g., using a three-pronged swivel clamp on a vertical post). Attach tubing to the bottom of the reservoir, run the tubing through a roller clamp, and connect the other end of the tube to a clean 20 G needle.</li>
	<li>Fill the 60 mL reservoir with approximately 30 mL of chilled sucrose solution and direct a carbogen line into the sucrose reservoir to continually bubble the perfusate. 
	NOTE: Ensure that the sucrose is flowing through the tubing and needle without any trapped bubbles. The flow rate should be just fast enough to have a steady, continual stream of dripping sucrose from the needle.</li>
	<li>Using a blender, crush and blend the frozen sucrose into an icy slurry and use the large spatula/spoon to distribute the slurry.
	<ol>
		<li>Place a small amount of sucrose slurry around the edges of the glass Petri dish lid. Add a small amount of sucrose slurry to the Petri dish bottom.</li>
		<li>Add roughly 20&#8211;30 mL of sucrose slurry to the perfusion fluid reservoir, mixing it well with the 30 mL of chilled liquid sucrose until the reservoir contains a mixture of very cold, predominantly liquid sucrose with some remnant frozen solution.</li>
	</ol>
	</li>
	<li>Place the mouse into a chamber connected to an isoflurane vaporizer. With oxygen flowing at approximately 2 L/min, turn the dial of the vaporizer to deliver isoflurane at 5% concentration and start a timer. 
	NOTE: Keep this timer going throughout the course of slicing to gauge how quickly the slices are obtained. The procedure should be done as quickly as possible, such that slicing is complete, and tissue is recovering in the interface chamber within 15&#8211;20 min of the animal entering the isoflurane chamber. Watch the mouse to ensure a state of deep anesthesia is achieved. After a minimum of 5 min, the mouse should be deeply anesthetized and unresponsive to toe pinches.</li>
	<li>Immediately before removing the mouse from the isoflurane chamber, fill the Petri dish lid with chilled sucrose solution to a depth of approximately 3&#8211;5 mm and fill the Petri dish bottom with chilled sucrose solution to a depth of approximately 1.0 cm.</li>
	<li>Quickly transfer the mouse to the left absorbent pad and use the 3 pieces of tape to secure the forelimbs and tail. Perform a hindlimb toe pinch to ensure that the mouse is unresponsive before any incision is performed.&#160;Using the large tissue forceps and surgical scissors, tent the skin and make a lengthwise incision from the bottom of the sternum to the top of the chest. Using the forceps pull up on the sternum and use the scissors to cut through the diaphragm. 
	NOTE: The initial positioning of the mouse and first several incisions to sever the diaphragm should be performed as quickly as possible to ensure the mouse does not regain consciousness.&#160;To ensure an adequate depth of anesthesia throughout the procedure, a nose cone can be placed on the mouse to deliver isoflurane during the perfusion.</li>
	<li>Use the scissors to cut through the rib cage on each side, cutting in one large motion towards the point where the forelimb meets the body. With the forceps, swing the front of the rib cage away and up toward the head, and then fully remove it with a horizontal cut using the large scissors. Hold the heart in place using the large forceps and insert the 20 G perfusion needle into the left ventricle. Once the needle is positioned correctly, the left side of the heart should quickly pale as chilled sucrose solution fills the ventricle.</li>
	<li>Use the small dissector scissors to cut into the right atrium and allow the blood to flow out of the circulatory system. If the incisions are performed correctly, there should be minimal damage to the heart, and it should continue pumping throughout the perfusion. 
	NOTE: Rolled-up pieces of tissue paper can be used to wick away blood and sucrose solution from the heart and maintain visibility of the perfusion needle during the procedure.</li>
	<li>Ensure that the perfusion needle stays in place and does not fall out of the left ventricle. With a proper flow rate and placement, the liver will start to pale to a light tan/beige color with 20&#8211;30 s.</li>
	<li>After 30&#8211;45 s, use the large scissors to decapitate the mouse. Holding the head in the left hand, push or peel the skin away from the skull. In a well-perfused animal, the skull should be very pale, and the brain should appear a very light pink color (approaching white) through the skull without clearly visible blood vessels.</li>
</ol>
5. Extract the brain and cut slices
<ol>
	<li>Using the small Bonn scissors, make two lateral cuts through the skull towards the midline at the front of the skull, near the eyes. Make two additional cuts on either side of the base of the skull.</li>
	<li>Transfer the head to the bottom of the glass Petri dish where it should be mostly immersed in chilled sucrose solution. Use the small Bonn scissors to cut down the midline along the length of the skull, pulling up with the scissors to minimize damage to the underlying brain tissue. Using the small tissue forceps, firmly grasp each side of the skull and swing it up and away from the brain, like opening a book.</li>
	<li>Use the fingers of the left hand to hold the flaps of skull open and insert the microspatula under the brain near the olfactory bulbs. Flip the brain out of the skull into the sucrose and use the microspatula to sever the brain stem. Wash the brain to remove any residual blood, fur, or other tissues.</li>
	<li>Use the large spatula/spoon to transfer the brain to the lid of the glass Petri dish. With the other half of the double-edged razor blade previously set aside, make two coronal cuts to first remove the cerebellum and then the most anterior portion of the brain, including the olfactory bulbs (Figure 1).</li>
	<li>Apply cyanoacrylate adhesive to the agar ramp. Very briefly place the brain onto a piece of dry filter paper using the large tissue forceps and then immediately transfer it to the agar ramp, placing the ventral surface of the brain onto the adhesive. 
	NOTE: For slices of the intermediate hippocampus, the brain should be oriented with the anterior facing up the slope of the ramp, and the posterior closer to the blade. For slices of the ventral hippocampus, orient the brain with the anterior end pointed down the slope of the ramp, with the posterior end at the top of the ramp, further away from the blade. In either case, the brain should be positioned at the top of the ramp, such that the coronally-cut surface of the brain contacts the agar backing block (Figure 1).</li>
	<li>Place the slicing platform with the agar and brain into the slicing chamber of the microtome and completely immerse with chilled sucrose solution. Use the large spatula/spoon to transfer some sucrose slurry to the chamber, stirring to melt any frozen sucrose and rapidly bring down the temperature of the mixture to ~1&#8211;2 &#176;C. 
	NOTE: Watch the temperature gauge throughout the slicing. If the sucrose solution warms above 3 &#176;C, add more slurry and mix to bring the temperature back down.</li>
	<li>Cut slices at a thickness of 450 &#956;m with the microtome speed set to 0.07 mm/s. As each slice is freed, use the small tissue forceps and a sharp scalpel to first separate the two hemispheres, and then to cut away tissue until the slice consists primarily of the hippocampus and parahippocampal regions (Figure 1).</li>
	<li>Use a plastic transfer pipette to transfer the slices individually to the interface recovery chamber, ensuring that they are positioned at the interface of the ACSF and the air with only a thin meniscus of ACSF covering the slices. Tightly close the lid of the chamber and allow slices to recover at 32 &#176;C for 30 min.</li>
	<li>After the initial recovery at 32 &#176;C, bring the interface recovery chamber out of the water bath and place on a stirrer set to a slow speed such that the magnetic stir bar promotes circulation of ACSF within the chamber. Allow it to gradually cool to room temperature as the slices recover for an additional 90 min. Ensure that the recovery chamber is continually bubbled with carbogen and do not allow large bubbles to become trapped underneath the slices.</li>
</ol>
6. Perform local field potential (LFP) recordings of spontaneous activity
<ol>
	<li>Prepare for LFP recordings by turning on all necessary equipment, including the computer running the acquisition software, the monitor connected to the computer, the stimulators, the micromanipulator, the temperature controller, the microscope light source, the microscope-attached camera, the microelectrode amplifier, the digitizer, and the peristaltic pump. If using a central vacuum system, open the wall valve to prepare the vacuum line that will remove ACSF from the recording chamber.</li>
	<li>Fill the heated reservoir with ACSF, then place one end of the tubing into the 400 mL beaker containing the bubbling ACSF. Turn on the peristaltic pump to direct ACSF from the 400 mL beaker to the heated reservoir, and from the reservoir onward to the recording chamber. Tap or pinch the tubing to release any trapped bubbles.</li>
	<li>Adjust the peristaltic pump to ensure that the ACSF flow rate through the recording chamber is fast (~ 8&#8211;10 mL/min). Use a temperature probe to ensure that the ACSF is 32 &#176;C in the center of the recording chamber. 
	NOTE: If ACSF is delivered to the recording chamber using a peristaltic pump, a high flow rate can result in a significant fluctuation in flow rate. Consistent flow rates can be achieved using a simple pulsation dampener consisting of a series of empty syringes integrated into the tubing (Figure 2).</li>
	<li>Prepare stimulation and recording pipettes from borosilicate glass capillaries using a heated filament puller. The puller protocol should be configured to yield pipettes with a resistance of 2&#8211;3 MOhm for stimulation or local field potential recording electrodes.</li>
	<li>Fill stimulation pipettes with 1 M NaCl and LFP pipettes with ACSF.</li>
	<li>Briefly clamp the tubing and turn off the pump to pause flow of ACSF. Transfer a slice to the recording chamber using fine forceps to grasp a corner of the lens paper the slice is resting on-the slice will stick to the lens paper. Place the lens paper and slice into the recording chamber with the slice facing down, then &#8220;peel&#8221; away the lens paper leaving the slice submerged in the recording chamber. Secure the slice using a harp. 
	NOTE: Harps can be purchased pre-made or made in-lab using a U-shaped piece of stainless steel or platinum and fine nylon filament.</li>
	<li>Place a NaCl-filled stimulation pipette in the manual micromanipulator and slowly advance the tip of the pipette into the surface of the slice (e.g., in the stratum radiatum layer) at an angle of approximately 30&#8211;45&#176;. Once the tip of the pipette enters the tissue, advance the pipette forward slowly and refrain from large movements in the lateral or vertical direction that could unnecessarily damage axons within the slice. Place the tip of the pipette at least 50&#8211;100 &#956;m deep into the slice to avoid cells near the surface that were damaged during slicing.</li>
	<li>Place an ACSF-filled LFP pipette into the pipette holder attached to the motorized micromanipulator. Apply a very light positive pressure using either mouth pressure or a 1 mL syringe connected via a stopcock valve and a short length of tubing to the pipette holder. 
	NOTE: Position LFP pipettes within the slice in order to record the signals of interest: in the case of sharp wave ripples, one LFP pipette should be placed in the stratum pyramidale (SP) and a second pipette in the stratum radiatum (SR). This configuration allows for simultaneous recordings of the negative sharp-wave in the SR and the high-frequency ripple oscillation in the SP (Figure 3).</li>
	<li>Using the micromanipulator slowly advance the tip of the LFP pipette into the region of interest (e.g., the CA1 pyramidal cell layer) at an angle of approximately 30&#8211;45&#176;. Place the tip of the pipette at least 50&#8211;100 &#956;m deep into the slice to avoid cells near the surface that were damaged during slicing. While advancing the LFP pipette, continually deliver a small voltage test pulse using the acquisition software and watch for a sudden increase in electrode resistance. This may indicate that the pipette has been clogged or pressed up against a cell.</li>
	<li>Once the LFP pipette is positioned in the region of interest, carefully release the positive pressure by opening the valve on the tubing attached to the pipette holder.</li>
	<li>To record the LFP in a second location simultaneously, repeat steps 6.8&#8211;6.10 with a second pipette and micromanipulator.</li>
	<li>Use the microelectrode amplifier in the current clamp configuration to record spontaneous activity in the local field potential after using the bridge balance function in the acquisition software to correct for series resistance of the pipette. Spontaneous SWRs will appear as positive deflections in the extracellular potential of the SP layer (Figure 3).</li>
	<li>In order to record evoked field potentials, use the stimulators connected to the digitizer to deliver a short (200 us) square voltage pulse through the NaCl-filled stimulation pipette. Adjust the stimulation voltage dial to evoke a range of response&#160;amplitudes.</li>
</ol>

The author has nothing to disclose.

There are several steps in this slicing protocol designed to promote tissue health and favor the emergence of spontaneous naturalistic network activity: the mouse is transcardially perfused with chilled sucrose cutting solution; horizontal-entorhinal cortex (HEC) slices are cut at a thickness of 450 &#956;m from the intermediate or ventral hippocampus; slices recover at the interface of warmed ACSF and humidified, carbogen-rich air; during recordings slices are superfused with ACSF warmed to 32 &#176;C and delivered at a...

Electrophysiological measurement from living hippocampal slices in vitro is a powerful experimental approach with numerous advantages. The experimenter can use a microscope, micromanipulators, and a recording system to directly visualize and collect measurements from individual neurons in the tissue. Tissue slices are also very accessible to photostimulation or drug delivery for optogenetic, chemogenetic, or pharmacological experiments.
The hippocampal network generates highly synchronous population activity in vivo, visible as oscillations in the extracellular local field potential1,2,3,4,5. Brain slice methods have been leveraged to gain insight into the cellular and circuit mechanisms underlying these neuronal network oscillations. Foundational work from Maier et al. demonstrated that sharp wave-ripple complexes (SWRs) can emerge spontaneously in slices of the ventral hippocampus6,7. Subsequent studies from multiple investigators have gradually elucidated many aspects of SWRs, including the role of neuromodulators in regulating the network state of the hippocampus8,9,10 and the synaptic mechanisms that drive the in vitro reactivation of neuronal ensembles previously active during behavior in vivo11. Brain slice experiments have also provided insight into the gamma range oscillation (30&#8211;100 Hz), a distinct hippocampal network state believed to support memory encoding and recall12,13. Finally, recognizing the central role of the hippocampus and associated structures in the pathophysiology of temporal lobe epilepsy14,15, researchers have used hippocampal slice preparations to investigate the generation and propagation of epileptiform activity. Carter et al. demonstrated that combined hippocampal-entorhinal cortex slices prepared from chronically epileptic animals can spontaneously generate epileptiform discharges in vitro16. Subsequently, Karl&#243;cai et al. explored the mechanisms underlying epileptiform discharges in hippocampal slices by using modified artificial cerebrospinal fluid (ACSF) with altered ion concentrations (reduced Mg2+ or elevated K+) or added drugs (4AP or gabazine)17.
Investigators have developed numerous hippocampal slice approaches that differ in key ways: (1) the region of the hippocampus contained in the slice (dorsal, intermediate, or ventral); (2) the presence or absence of extrahippocampal tissues such as the entorhinal cortex; (3) the orientation used to cut slices (coronal, sagittal, horizontal, or oblique); and (4) the conditions under which the tissue is maintained after slicing (submerged fully in ACSF or held at the interface of ACSF and humidified, carbogen-rich air).
The choice of which slicing approach to use should be determined by the experimental objective. For example, transverse or coronal slices of the dorsal hippocampus maintained under submerged conditions have been used very effectively for the investigation of intrahippocampal circuitry and synaptic plasticity18,19,20. However, such preparations do not spontaneously generate network oscillations as readily as slices from the ventral hippocampus21,22,23. Although a state of persistent SWR activity can be induced by tetanic stimulation in transverse slices from the dorsal and ventral hippocampus24, spontaneous SWRs are more readily observed in ventral slices7,25.
An inherent physiological and anatomical distinction between the dorsal and ventral hippocampus is supported by studies performed both in vivo and in vitro26. Recordings in rats revealed strongly coherent theta rhythms throughout the dorsal and intermediate hippocampus, yet poor coherence between the ventral region and the rest of the hippocampus27. SWRs in vivo propagate readily between the dorsal and intermediate hippocampus, while SWRs that originate in the ventral hippocampus often remain local28. The associational projections originating from CA3 pyramidal neurons that reside in the dorsal and intermediate hippocampus project long distances along the longitudinal axis of the hippocampus. CA3 projections originating from ventral regions remain relatively local, and thus are less likely to be severed during the slicing process29,30. Ventral slices may, therefore, better preserve the recurrent network necessary to generate population synchrony. The propensity of ventral slices to generate spontaneous network activities in vitro may also reflect higher intrinsic excitability of pyramidal neurons or weaker GABAergic inhibition in the ventral hippocampus as compared to more dorsal regions31. Indeed, ventral hippocampal slices are more susceptible to epileptiform activity32,33. Thus, many studies of spontaneous physiological8,9,11,24 or pathological16,34,35,36 network oscillations have traditionally used a horizontal slicing approach, sometimes with a slight angle in the fronto-occipital direction, which yields tissue slices parallel to the transverse plane of the ventral hippocampus.
Network connectivity is unavoidably impacted by the slicing procedure as many cells in the slice will be severed. The angle and thickness of the slice and the tissue retained in the preparation should be considered to optimize connectivity in the circuits of interest. Many studies have utilized horizontal combined hippocampal-entorhinal cortex slices (HEC) to explore interactions between the two structures in the context of physiological or pathological network oscillations. Roth et al. performed dual recordings from the CA1 subfield of the hippocampus and layer V of the medial entorhinal cortex to demonstrate propagation of SWR activity through the HEC slice37. Many studies of epileptiform activity have used the HEC slice preparation to investigate how epileptiform discharges propagate through the corticohippocampal network16,35,36,38. It is important to note that preservation of the intact corticohippocampal loop is not a prerequisite for spontaneous SWRs, epileptiform discharges, or gamma oscillations; network oscillations can be generated in transverse slices of the dorsal or ventral hippocampus with no attached parahippocampal tissues21,22,23, 25,39,40,41. A more important factor for the spontaneous generation of network oscillations in hippocampal slices may be the thickness of each slice, as a thicker slice (400&#8211;550 &#956;m) will preserve more connectivity in the CA2/CA3 recurrent network21,22,25.
Although angled horizontal HEC slices (cut with an approximately 12&#176; angle in the fronto-occipital direction) have been used to study the functional connectivity of the corticohippocampal loop11,16,34,35,42, such angled preparations are not required for spontaneous network activity43,44,45. However, the use of an angled slicing plane does allow the investigator to selectively make slices that best preserve the transversely-oriented lamellae of either the ventral or intermediate hippocampus, depending on whether a downward or an upward angle is applied (Figure 1). This approach is conceptually similar to that used by Papatheodoropoulos et al., 2002, who dissected each hippocampus free and then used a tissue chopper to create transverse slices along the entire dorsal-ventral axis21. In the light of the aforementioned functional distinctions between the ventral and dorsal-intermediate hippocampus, investigators should consider the anatomical origin of slices when designing experiments or interpreting results. Using an agar ramp during the slicing procedure is a simple way to preferentially produce slices from either the intermediate or ventral hippocampus.
Hippocampal slices can be maintained in either a submerged chamber (with the tissue fully immersed in ACSF), or an interface-style chamber (e.g., Oslo or Haas chamber, with slices covered only by a thin film of flowing media). Interface maintenance enhances oxygenation of the tissue, which promotes neuronal survival and allows for sustained high levels of interneuronal activity. Traditionally, submerged recording conditions use a slower ACSF flow rate that does not provide adequate tissue oxygenation for stable expression of network-level oscillations. In submerged hippocampal slices carbachol-induced gamma oscillations are only observed transiently46,47, while they can be stably maintained in interface recording chambers10,48,49. As such, many studies of complex spontaneous activity in vitro have relied on interface recording chambers to investigate sharp-wave ripple complexes6,7,8,9,10,25,37, gamma oscillations10,13, and epileptiform activity16,38,45,47.
In a submerged-style recording chamber, an immersion microscope objective can be used to visualize individual cells and selectively target healthy-looking cells for recordings. The submerged preparation also allows fine control over the cellular milieu, as submersion facilitates rapid diffusion of drugs or other compounds to the tissue. Thus, a modified methodology in which stable network oscillations are maintained under submerged conditions represents a powerful experimental approach. This approach is exemplified by the work of H&#225;jos et al., in which hippocampal slices recover in a simplified interface-style holding chamber for several hours before transfer to a modified submerged recording chamber with a high flow rate of ACSF (~6 mL/min) to enhance oxygen supply to the tissue12,48,49. Under these conditions, high levels of interneuron activity and stable spontaneous network oscillations can be maintained in a submerged recording chamber. This modified approach allows the investigators to perform visually guided whole-cell patch clamp recordings and characterize the contribution of morphologically identified cell types to carbachol-induced gamma oscillations12. SWRs can also occur spontaneously in submerged hippocampal slices with a fast flow rate of ACSF11,48,49. Maier et al. demonstrated that hippocampal slices that recovered in an interface chamber before transfer to a submerged recording chamber reliably exhibited spontaneous SWRs, whereas slices that recovered submerged in a beaker before transfer to a submerged recording chamber showed smaller evoked field responses, lower levels of spontaneous synaptic currents, and only very rarely exhibited spontaneous SWRs43. Schlingloff et al. used this improved methodology to demonstrate the role of parvalbumin-expressing basket cells in the generation of spontaneous SWRs44.
The following protocol presents a slicing method through which spontaneously active neurons in horizontal hippocampal slices can be recovered under interface conditions and subsequently maintained in a submerged recording chamber suitable for pharmacological or optogenetic manipulations and visually guided recordings.

<table><tbody><tr><td>3D printer</td><td>Lulzbot</td><td>LulzBot TAZ 6</td><td></td></tr><tr><td>Acute brain slice incubation holder</td><td>NIH 3D Print Exchange</td><td>3DPX-001623</td><td>Designed by ChiaMing Lee, available at https://3dprint.nih.gov/discover/3dpx-001623</td></tr><tr><td>Adenosine 5&amp;prime;-triphosphate magnesium salt</td><td>Sigma Aldrich</td><td>A9187-500MG</td><td></td></tr><tr><td>Ag-Cl ground pellets</td><td>Warner</td><td>64-1309, (E205)</td><td></td></tr><tr><td>agar</td><td>Becton, Dickinson</td><td>214530-500g</td><td></td></tr><tr><td>ascorbic acid</td><td>Alfa Aesar</td><td>36237</td><td></td></tr><tr><td>beaker (250 mL)</td><td>Kimax</td><td>14000-250</td><td></td></tr><tr><td>beaker (400 mL)</td><td>Kimax</td><td>14000-400</td><td></td></tr><tr><td>biocytin</td><td>Sigma Aldrich</td><td>B4261</td><td></td></tr><tr><td>blender</td><td>Oster</td><td>BRLY07-B00-NP0</td><td></td></tr><tr><td>Bonn scissors, small</td><td>becton, Dickinson</td><td>14184-09</td><td></td></tr><tr><td>borosilicate glass capillaries with filament (O.D. 1.5 mm, I.D. 0.86 mm, length 10 cm)</td><td>Sutter Instruments</td><td>BF150-86-10HP</td><td>Fire polished capillaries are preferable.</td></tr><tr><td>calcium chloride solution (1 M)</td><td>G-Biosciences</td><td>R040</td><td></td></tr><tr><td>camera</td><td>Olympus</td><td>OLY-150</td><td></td></tr><tr><td>compressed carbogen gas (95% oxygen / 5% carbon dioxide)</td><td>Airgas</td><td>X02OX95C2003102</td><td></td></tr><tr><td>compressed oxygen</td><td>Airgas</td><td>OX 200</td><td></td></tr><tr><td>constant voltage isolated stimulator</td><td>Digitimer Ltd.</td><td>DS2A-Mk.II</td><td></td></tr><tr><td>coverslips (22x50 mm)</td><td>VWR</td><td>16004-314</td><td></td></tr><tr><td>cyanoacrylate adhesive</td><td>Krazy Glue</td><td>KG925</td><td>Ideally use the brush-on form for precision</td></tr><tr><td>data acquisition software</td><td>Axograph</td><td>N/A</td><td>Any equivalent software (e.g. pClamp) would work.</td></tr><tr><td>Dell Precision T1500 Tower Workstation Desktop</td><td>Dell</td><td>N/A</td><td>Catalog number will depend on specific computer - any computer will work as long as it can run electrophysiology acquisition software.</td></tr><tr><td>Digidata 1440A</td><td>Molecular Devices</td><td>1-2950-0367</td><td></td></tr><tr><td>digital timer</td><td>VWR</td><td>62344-641</td><td>4-channel Traceable timer</td></tr><tr><td>disposable absorbant pads</td><td>VWR</td><td>56616-018</td><td></td></tr><tr><td>dissector scissors</td><td>Fine Science Tools</td><td>14082-09</td><td></td></tr><tr><td>double-edge razor blades</td><td>Personna</td><td>BP9020</td><td></td></tr><tr><td>dual automatic temperature controller</td><td>Warner Instrument Corporation</td><td>TC-344B</td><td></td></tr><tr><td>dual-surface or laminar-flow optimized recording chamber</td><td>N/A</td><td>N/A</td><td>The chamber presented in this protocol is custom made. A commercial equivalent would be the RC-27L from Warner Instruments.</td></tr><tr><td>equipment rack</td><td>Automate Scientific</td><td>FR-EQ70&quot;</td><td>A rack is not strictly necessary but useful for organizing electrophysiology</td></tr><tr><td>Ethylene glycol-bis(2-aminoethyiether)- N,N,N&#039;,N&#039;-teetraacetic acid (EGTA)</td><td>Sigma Aldrich</td><td>324626-25GM</td><td></td></tr><tr><td>filter paper</td><td>Whatman</td><td>1004 070</td><td></td></tr><tr><td>fine scale</td><td>Mettler Toledo</td><td>XS204DR</td><td></td></tr><tr><td>Flaming/Brown micropipette puller</td><td>Sutter Instruments</td><td>P-97</td><td></td></tr><tr><td>glass petri dish (100 x 15 mm)</td><td>Corning</td><td>3160-101</td><td></td></tr><tr><td>glucose</td><td>Fisher Scientific</td><td>D16-1</td><td></td></tr><tr><td>Guanosine 5&amp;prime;-triphosphate sodium salt hydrate</td><td>Sigma Aldrich</td><td>G8877-250MG</td><td></td></tr><tr><td>ice buckets</td><td>Sigma Aldrich</td><td>BAM168072002-1EA</td><td></td></tr><tr><td>isoflurane vaporizer</td><td>General Anesthetic Services</td><td>Tec 3</td><td></td></tr><tr><td>lab tape</td><td>Fisher Scientific</td><td>15-901-10R</td><td></td></tr><tr><td>lens paper</td><td>Fisher Scientific</td><td>11-996</td><td></td></tr><tr><td>light source</td><td>Olympus</td><td>TH4-100</td><td></td></tr><tr><td>magnesium chloride solution (1 M)</td><td>Quality Biological</td><td>351-033-721EA</td><td></td></tr><tr><td>magnetic stir bars</td><td>Fisher Scientific</td><td>14-513-56</td><td>Catalog number will be dependent on the size of the stir bar.</td></tr><tr><td>micromanipulator</td><td>Luigs &amp;amp; Neumann</td><td>SM-5</td><td></td></tr><tr><td>micromanipulator (manual)</td><td>Scientifica</td><td>LBM-2000-00</td><td></td></tr><tr><td>microscope</td><td>Olympus</td><td>BX51WI</td><td></td></tr><tr><td>microspatula</td><td>Fine Science Tools</td><td>10089-11</td><td></td></tr><tr><td>monitor</td><td>Dell</td><td>2007FPb</td><td></td></tr><tr><td>MultiClamp 700B Microelectrode Amplifier</td><td>Molecular Devices</td><td>MULTICLAMP 700B</td><td>The MultiClamp 700B should include headstages, pipette holders, and a model cell.</td></tr><tr><td>N-(2-Hydroxyethyl)piperazine-N&amp;prime;-(2-ethanesulfonic acid), (HEPES)</td><td>Sigma Aldrich</td><td>H3375-25G</td><td></td></tr><tr><td>needle (20 gauge, 1.5 in length)</td><td>Becton, Dickinson</td><td>305176</td><td></td></tr><tr><td>nylon filament</td><td>YLI Wonder Invisible Thread</td><td>212-15-004</td><td>size 0.004. This cat. # is from Amazon.com</td></tr><tr><td>nylon mesh</td><td>Warner Instruments Corporation</td><td>64-0198</td><td></td></tr><tr><td>perstaltic pump</td><td>Harvard Apparatus</td><td>70-2027</td><td></td></tr><tr><td>Phosphocreatine di(tris) salt</td><td>Sigma Aldrich</td><td>P1937-1G</td><td></td></tr><tr><td>pipette holders</td><td>Molecular Devices</td><td>1-HL-U</td><td></td></tr><tr><td>platinum wire</td><td>World Precision</td><td>PT0203</td><td></td></tr><tr><td>polylactic acid (PLA) filament</td><td>Ultimaker</td><td>RAL 9010</td><td></td></tr><tr><td>potassium chloride</td><td>Sigma Aldrich</td><td>P3911-500G</td><td></td></tr><tr><td>potassium gluconate</td><td>Sigma Aldrich</td><td>1550001-200MG</td><td></td></tr><tr><td>potassium hydroxide</td><td>Sigma Aldrich</td><td>60377-1KG</td><td></td></tr><tr><td>razor blades</td><td>VWR</td><td>55411-050</td><td></td></tr><tr><td>roller clamp</td><td>World Precision Instruments</td><td>14041</td><td></td></tr><tr><td>scale</td><td>Mettler Toledo</td><td>PM2000</td><td></td></tr><tr><td>scalpel handle</td><td>Fine Science Tools</td><td>10004-13</td><td></td></tr><tr><td>slice harp</td><td>Warner</td><td>SHD-26GH/2</td><td></td></tr><tr><td>sodium bicarbonate</td><td>Fisher Chemical</td><td>S233-500</td><td></td></tr><tr><td>sodium chloride</td><td>Sigma Aldrich</td><td>S9888-1KG</td><td></td></tr><tr><td>sodium phosphate monobasic anhydrous</td><td>Fisher Chemical</td><td>S369-500</td><td></td></tr><tr><td>sodium pyruvate</td><td>Fisher Chemical</td><td>BP356-100</td><td></td></tr><tr><td>spatula</td><td>VWR</td><td>82027-520</td><td></td></tr><tr><td>spatula/spoon, large</td><td>VWR</td><td>470149-442</td><td></td></tr><tr><td>sterile scalpel blades</td><td>Feather</td><td>72044-10</td><td></td></tr><tr><td>stirrer / hot plate</td><td>Corning</td><td>6795-220</td><td></td></tr><tr><td>stopcock valves, 1-way</td><td>World Precision Instruments</td><td>14054</td><td></td></tr><tr><td>stopcock valves, 3-way</td><td>World Precision Instruments</td><td>14036</td><td></td></tr><tr><td>sucrose</td><td>Acros Organics</td><td>AC177142500</td><td></td></tr><tr><td>support for swivel clamps</td><td>Fisher Scientific</td><td>14-679Q</td><td></td></tr><tr><td>surgical scissors, sharp/blunt</td><td>Fine Science Tools</td><td>14001-12</td><td></td></tr><tr><td>syringe (1 mL)</td><td>Becton, Dickinson</td><td>309659</td><td></td></tr><tr><td>syringe (60 mL with Luer-Lok tip)</td><td>Becton, Dickinson</td><td>309653</td><td></td></tr><tr><td>three-pronged clamp</td><td>Fisher Scientific</td><td>05-769-8Q</td><td></td></tr><tr><td>tissue forceps, large</td><td>Fine Science Tools</td><td>11021-15</td><td></td></tr><tr><td>tissue forceps, small</td><td>Fine Science Tools</td><td>11023-10</td><td></td></tr><tr><td>transfer pipettes</td><td>Fisher Scientific</td><td>13-711-7M</td><td></td></tr><tr><td>tubing</td><td>Tygon</td><td>E-3603</td><td>ID 1/16 inch, OD 3/16 inch</td></tr><tr><td>tubing</td><td>Tygon</td><td>R-3603</td><td>ID 1/8 inch, OD 1/4 inch</td></tr><tr><td>vacuum grease</td><td>Dow Corning</td><td>14-635-5D</td><td></td></tr><tr><td>vibrating blade microtome</td><td>Leica</td><td>VT 1200S</td><td></td></tr><tr><td>vibration-dampening table with faraday cage</td><td>Micro-G / TMC-ametek</td><td>2536-516-4-30PE</td><td></td></tr><tr><td>volumetric flask (1 L)</td><td>Kimax</td><td>KIM-28014-1000</td><td></td></tr><tr><td>volumetric flask (2 L)</td><td>PYREX</td><td>65640-2000</td><td></td></tr><tr><td>warm water bath</td><td>VWR</td><td>1209&lt;br/&gt; &amp;nbsp;</td><td></td></tr></tbody></table>

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.

Presented here are representative recordings from HEC slices prepared as described in this protocol. Following recovery in an interface holding chamber (Figure 1C), slices are transferred individually to a submerged recording chamber (Figure 2B). The recording chamber is supplied with carbogen-saturated ACSF using a peristaltic pump (Figure 2A). The pump first draws ACSF from a holding beaker into a heated reservoir. Carbogen lines ...

Watch this Scientific Journal Video about Acute Mouse Brain Slicing to Investigate Spontaneous Hippocampal Network Activity at JoVE.com

Acute Mouse Brain Slicing to Investigate Spontaneous Hippocampal Network Activity

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 ...

acute-mouse-brain-slicing-to-investigate-spontaneous-hippocampal

Research

JoVE Journal

Neuroscience

8.9K Views.  Columbia University. 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.

Video: Acute Mouse Brain Slicing to Investigate Spontaneous Hippocampal Network Activity

Preparation of Acute Hippocampal Slices from Rats and Transgenic Mice for the Study of Synaptic Alterations during Aging and Amyloid Pathology

The rodent hippocampal slice preparation is perhaps the most broadly used tool for investigating mammalian synaptic function and plasticity. The hippocampus can be extracted quickly and easily from rats and mice and slices remain viable for hours in oxygenated artificial cerebrospinal fluid. Moreover, basic electrophysisologic techniques are easily applied to the investigation of synaptic function in hippocampal slices and have provided some of the best biomarkers for cognitive impairments. The hippocampal slice is especially popular for the study of synaptic plasticity mechanisms involved in learning and memory. Changes in the induction of long-term potentiation and depression (LTP and LTD) of synaptic efficacy in hippocampal slices (or lack thereof) are frequently used to describe the neurologic phenotype of cognitively-impaired animals and/or to evaluate the mechanism of action of nootropic compounds. This article outlines the procedures we use for preparing hippocampal slices from rats and transgenic mice for the study of synaptic alterations associated with brain aging and Alzheimer's disease (AD)1-3. Use of aged rats and AD model mice can present a unique set of challenges to researchers accustomed to using younger rats and/or mice in their research. Aged rats have thicker skulls and tougher connective tissue than younger rats and mice, which can delay brain extraction and/or dissection and consequently negate or exaggerate real age-differences in synaptic function and plasticity. Aging and amyloid pathology may also exacerbate hippocampal damage sustained during the dissection procedure, again complicating any inferences drawn from physiologic assessment. Here, we discuss the steps taken during the dissection procedure to minimize these problems. Examples of synaptic responses acquired in "healthy" and "unhealthy" slices from rats and mice are provided, as well as representative synaptic plasticity experiments. The possible impact of other methodological factors on synaptic function in these animal models (e.g. recording solution components, stimulation parameters) are also discussed. While the focus of this article is on the use of aged rats and transgenic mice, novices to slice physiology should find enough detail here to get started on their own studies, using a variety of rodent models.

This article outlines procedures for preparing hippocampal slices from rats and transgenic mice for the study of synaptic alterations associated with brain aging and age-related neurodegenerative diseases, such as Alzheimer&#x2019;s disease.

The rodent hippocampal slice preparation is perhaps the most broadly used tool for investigating mammalian synaptic function and plasticity. The ...

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 ...

Neural Activity Propagation in an Unfolded Hippocampal Preparation with a Penetrating Micro-electrode Array

This protocol describes a method for preparing a new in vitro flat hippocampus preparation combined with a micro-machined array to map neural activity in the hippocampus. The transverse hippocampal slice preparation is the most common tissue preparation to study hippocampus electrophysiology. A longitudinal hippocampal slice was also developed in order to investigate longitudinal connections in the hippocampus. The intact mouse hippocampus can also be maintained in vitro because its thickness allows adequate oxygen diffusion. However, these three preparations do not provide direct access to neural propagation since some of the tissue is either missing or folded. The unfolded intact hippocampus provides both transverse and longitudinal connections in a flat configuration for direct access to the tissue to analyze the full extent of signal propagation in the hippocampus in vitro. In order to effectively monitor the neural activity from the cell layer, a custom made penetrating micro-electrode array (PMEA) was fabricated and applied to the unfolded hippocampus. The PMEA with 64 electrodes of 200 &#181;m in height could record neural activity deep inside the mouse hippocampus. The unique combination of an unfolded hippocampal preparation and the PMEA provides a new in-vitro tool to study the speed and direction of propagation of neural activity in the two-dimensional CA1-CA3 regions of the hippocampus with a high signal to noise ratio.

We have developed an in vitro unfolded hippocampus which preserves CA1-CA3 array of neurons. Combined with the penetrating micro-electrode array, neural activity can be monitored in both the longitudinal and transverse orientations. This method provides advantages over hippocampal slice preparations as the propagation in the entire hippocampus can be recorded simultaneously.

This protocol describes a method for preparing a new in vitro flat hippocampus preparation combined with a micro-machined array to map neural activity in ...

Evaluation of Synapse Density in Hippocampal Rodent Brain Slices

In the brain, synapses are specialized junctions between neurons, determining the strength and spread of neuronal signaling. The number of synapses is tightly regulated during development and neuronal maturation. Importantly, deficits in synapse number can lead to cognitive dysfunction. Therefore, the evaluation of synapse number is an integral part of neurobiology. However, as synapses are small and highly compact in the intact brain, the assessment of absolute number is challenging. This protocol describes a method to easily identify and evaluate synapses in hippocampal rodent slices using immunofluorescence microscopy. It includes a three-step procedure to evaluate synapses in high-quality confocal microscopy images by analyzing the co-localization of pre- and postsynaptic proteins in hippocampal slices. It also explains how the analysis is performed and gives representative examples from both excitatory and inhibitory synapses. This protocol provides a solid foundation for the analysis of synapses and can be applied to any research investigating the structure and function of the brain.

A protocol for accurately identifying and analyzing synapses in hippocampal slices using immunofluorescence is outlined in this article.

In the brain, synapses are specialized junctions between neurons, determining the strength and spread of neuronal signaling. The number of synapses is ...

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) ...

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 ...

Horizontal Hippocampal Slices of the Mouse Brain

The hippocampus is a highly organized structure in the brain that is a part of the limbic system and is involved in memory formation and consolidation as well as the manifestation of severe brain disorders, including Alzheimer&#8217;s disease and epilepsy. The hippocampus receives a high degree of intra- and inter-connectivity, securing a proper communication with internal and external brain structures. This connectivity is accomplished via different informational flows in the form of fiber pathways. Brain slices are a frequently used methodology when exploring neurophysiological functions of the hippocampus. Hippocampal brain slices can be used for several different applications, including electrophysiological recordings, light microscopic measurements as well as several molecular biological and histochemical techniques. Therefore, brain slices represent an ideal model system to assess protein functions, to investigate pathophysiological processes involved in neurological disorders as well as for drug discovery purposes.
There exist several different ways of slice preparations. Brain slice preparations with a vibratome allow a better preservation of the tissue structure and guarantee a sufficient oxygen supply during slicing, which present advantages over the traditional use of a tissue chopper. Moreover, different cutting planes can be applied for vibratome brain slice preparations. Here, a detailed protocol for a successful preparation of vibratome-cut horizontal hippocampal slices of mouse brains is provided. In contrast to other slice preparations, horizontal slicing allows to keep the fibers of the hippocampal input path (perforant path) in a fully intact state within a slice, which facilitates the investigation of entorhinal-hippocampal interactions. Here, we provide a thorough protocol for the dissection, extraction, and acute horizontal slicing of the murine brain, and discuss challenges and potential pitfalls of this technique. Finally, we will show some examples for the use of brain slices in further applications.

This article aims to describe a systematic protocol to obtain horizontal hippocampal brain slices in mice. The objective of this methodology is to preserve the integrity of hippocampal fiber pathways, such as the perforant path and the mossy fiber tract to assess dentate gyrus related neurological processes.

The hippocampus is a highly organized structure in the brain that is a part of the limbic system and is involved in memory formation and consolidation as ...

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 ...

Craniotomy Procedure for Visualizing Neuronal Activities in Hippocampus of Behaving Mice

Imaging neuronal activities at single-cell resolution in awake behaving animals is a very powerful approach for the investigation of neural circuit functions in systems neuroscience. However, high absorbance and scattering of light in mammalian tissue limit intravital imaging mostly to superficial brain regions, leaving deep-brain areas, such as the hippocampus, out of reach for optical microscopy. In this video, we show the preparation and implantation of the custom-made imaging window to enable chronic in vivo imaging of the dorsal hippocampal CA1 region in head-fixed behaving mice. The custom-made window is supplemented with an infusion cannula that allows targeted delivery of viral vectors and drugs to the imaging area. By combining this preparation with wide-field imaging, we performed a long-term recording of neuronal activity using a fluorescent calcium indicator from large subsets of neurons in behaving mice over several weeks. We also demonstrated the applicability of this preparation for voltage imaging with single-spike resolution. High-performance genetically encoded indicators of neuronal activity and scientific CMOS cameras allowed the recurrent visualization of subcellular morphological details of single neurons at high temporal resolution. We also discuss the advantages and potential limitations of the described method and its compatibility with other imaging techniques.

This article demonstrates the preparation of a custom-made imaging window supplemented with infusion cannula and its implantation onto the CA1 region of the hippocampus in mice.

Imaging neuronal activities at single-cell resolution in awake behaving animals is a very powerful approach for the investigation of neural circuit ...

Recording of Local Field Potential in Mouse Hippocampal-Entorhinal Cortex Slices

Source: Whitebirch, A. C. <a href="https://app.jove.com/v/61704">Acute Mouse Brain Slicing to Investigate Spontaneous Hippocampal Network Activity.</a> J. Vis. Exp. (2020)
The video demonstrates a procedure to record the local field potential (LFP) in mouse hippocampal-entorhinal cortex slices. Electrical stimulation is provided at the CA3 stratum radiatum of the hippocampus, which causes changes in the postsynaptic potential at the CA1 stratum pyramidale. The combined change in the membrane potential of CA1 neurons, termed the local field potential, is recorded.

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1. Transcardial perfusion

	Suspend ...