Acute Mouse Brain Slicing to Investigate Spontaneous Hippocampal Network Activity

Summary

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.

Abstract

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 μm 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.

Introduction

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 potential^1,2,3,4,5. 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 hippocampus^6,7. Subsequent studies from multiple investigators have gradually elucidated many aspects of SWRs, including the role of neuromodulators in regulating the network state of the hippocampus^8,9,10 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 (30–100 Hz), a distinct hippocampal network state believed to support memory encoding and recall^12,13. Finally, recognizing the central role of the hippocampus and associated structures in the pathophysiology of temporal lobe epilepsy^14,15, 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, Karlócai 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 plasticity^18,19,20. However, such preparations do not spontaneously generate network oscillations as readily as slices from the ventral hippocampus^21,22,23. 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 slices^7,25.

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 process^29,30. 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 activity^32,33. Thus, many studies of spontaneous physiological^8,9,11,24 or pathological^16,34,35,36 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 network^16,35,36,38. 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 tissues^{21,22,23, 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 (400–550 μm) will preserve more connectivity in the CA2/CA3 recurrent network^21,22,25.

Although angled horizontal HEC slices (cut with an approximately 12° angle in the fronto-occipital direction) have been used to study the functional connectivity of the corticohippocampal loop^{11,16,34,35,42}, such angled preparations are not required for spontaneous network activity^43,44,45. 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 transiently^46,47, while they can be stably maintained in interface recording chambers^10,48,49. As such, many studies of complex spontaneous activity in vitro have relied on interface recording chambers to investigate sharp-wave ripple complexes^{6,7,8,9,10,25,37}, gamma oscillations^10,13, and epileptiform activity^16,38,45,47.

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 Hájos 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 tissue^12,48,49. 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 ACSF^11,48,49. 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.

Protocol

All methods described here have been approved by the Institutional Animal Care and Use Committee at Columbia University (AC-AAAU9451).

1. Prepare solutions

1. Prepare sucrose cutting solution for slicing as described in Table 1.
   NOTE: After preparing 1 L of sucrose solution, freeze a small amount (approximately 100–200 mL) in an ice tray. These frozen sucrose ice cubes will be blended into an icy slurry (see step 4.3).
2. Prepare artificial cerebrospinal fluid (ACSF) for recording as described in Table 2.
3. 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.

2. Prepare agar ramp

1. 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 30–60 s). Pour the molten agar into a mold and allow it to cool and solidify at 4 °C in a fridge at an approximately 12° 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.

3. Stage the slicing area

1. 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 °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).
2. 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.
3. Pull out the tray of premade, frozen sucrose solution ice cubes from the freezer and place on the bench to partially thaw.
4. 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.
5. Run a carbogen line and temperature probe into the slicing chamber and surround with ice to chill the chamber.
6. 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).
7. 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.
8. 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).

4. Transcardial perfusion

1. 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.
2. 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.
3. Using a blender, crush and blend the frozen sucrose into an icy slurry and use the large spatula/spoon to distribute the slurry.
   1. 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.
   2. Add roughly 20–30 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.
4. 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 15–20 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.
5. Immediately before removing the mouse from the isoflurane chamber, fill the Petri dish lid with chilled sucrose solution to a depth of approximately 3–5 mm and fill the Petri dish bottom with chilled sucrose solution to a depth of approximately 1.0 cm.
6. 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. 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. 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.
7. 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.
8. 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.
9. 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 20–30 s.
10. After 30–45 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.

5. Extract the brain and cut slices

1. 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.
2. 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.
3. 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.
4. 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).
5. 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).
6. 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 ~1–2 °C.
   NOTE: Watch the temperature gauge throughout the slicing. If the sucrose solution warms above 3 °C, add more slurry and mix to bring the temperature back down.
7. Cut slices at a thickness of 450 μm 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).
8. 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 °C for 30 min.
9. After the initial recovery at 32 °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.

6. Perform local field potential (LFP) recordings of spontaneous activity

1. 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.
2. 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.
3. Adjust the peristaltic pump to ensure that the ACSF flow rate through the recording chamber is fast (~ 8–10 mL/min). Use a temperature probe to ensure that the ACSF is 32 °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).
4. 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 2–3 MOhm for stimulation or local field potential recording electrodes.
5. Fill stimulation pipettes with 1 M NaCl and LFP pipettes with ACSF.
6. 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 “peel” 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.
7. 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 30–45°. 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 50–100 μm deep into the slice to avoid cells near the surface that were damaged during slicing.
8. 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).
9. 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 30–45°. Place the tip of the pipette at least 50–100 μm 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.
10. 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.
11. To record the LFP in a second location simultaneously, repeat steps 6.8–6.10 with a second pipette and micromanipulator.
12. 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).
13. 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 amplitudes.

Results

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 ...

Discussion

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 μm 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 °C and delivered at a...

Materials

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