Minimizing Hypoxia in Hippocampal Slices from Adult and Aging Mice

Summary

This is a protocol for acute slice preparation from adult and aging mouse hippocampi that takes advantage of transcardial perfusion and slice cutting with ice-cold NMDG-aCSF to reduce hypoxic damage to the tissue. The resulting slices stay healthy over many hours, and are suitable for long-term patch-clamp and field-recordings.

Abstract

Acute hippocampal slices have enabled generations of neuroscientists to explore synaptic, neuronal, and circuit properties in detail and with high fidelity. Exploration of LTP and LTD mechanisms, single neuron dendritic computation, and experience-dependent changes in circuitry, would not have been possible without this classical preparation. However, with a few exceptions, most basic research using acute hippocampal slices has been performed using slices from rodents of relatively young ages, ~P20-P40, even though synaptic and intrinsic excitability mechanisms have a long developmental tail that reaches past P60. The main appeal of using young hippocampal slices is preservation of neuronal health aided by higher tolerance to hypoxic damage. However, there is a need to understand neuronal function at more mature stages of development, further accentuated by the development of various animal models of neurodegenerative diseases that require an aging brain preparation. Here we describe a modification to an acute hippocampal slice preparation that reliably delivers healthy slices from adult and aging mouse hippocampi. The protocol’s critical steps are transcardial perfusion and cutting with ice-cold sodium-free NMDG-aSCF. Together, these steps attenuate the hypoxia-induced drop in ATP upon decapitation, as well as cytotoxic edema caused by passive sodium fluxes. We demonstrate how to cut transversal slices of hippocampus plus cortex using a vibrating microtome. Acute hippocampal slices obtained in this way are reliably healthy over many hours of recording, and are appropriate for both field-recordings and targeted patch-clamp recordings, including targeting of fluorescently labeled neurons.

Introduction

The advent of mammalian acute brain slice preparations facilitated experiments at the cellular and synaptic level that were previously possible only in invertebrate preparations like Aplysia¹. Development of acute hippocampal slices was of particular significance, as it is a structure responsible for working memory and context formation, and has a specialized tri-synaptic circuitry that is amenable to easy physiological manipulation. However, the vast majority of acute brain slices are still prepared from relatively young mice and rats, as it is easier to preserve healthy neurons and circuits, and the slices remain viable for longer periods of time^2,3,4. Here, we introduce modifications to standard slicing protocols that result in increased viability of acute hippocampal slices from adult and aging mice.

The major impediment to the long-term ex vivo viability of mammalian brain parenchyma is the initial hypoxic damage that occurs rapidly once blood flow to the brain stops following decapitation. Loss of oxygen results in fast metabolic consumption of major energy resources in the brain with the loss of phospho-creatine (P-creatine) being the most rapid, followed by glucose, adenosine triphosphate (ATP), and glycogen⁴. Preservation of ATP is of particular importance for the long-term health of brain slices, as ATP is needed to maintain the membrane potential via the Na-K ATPase, and consequently the neural activity^5,6. The ATP level in the adult rodent brain is ~2.5 mM, and it drops precipitously within 20 s of decapitation to reach a basal steady state (~0.5 mM) at around 1 min post-decapitation^4,7,8. In young animals, it takes longer to observe the same drop in ATP (~2 min); with phenobarbital anesthesia it is further slowed to 4 min⁴. These considerations show that preventing loss of ATP and other energy resources is a necessary strategy to prevent hypoxic damage to the brain and in turn to maintain the health of brain slices over longer periods of time, especially in adult animals.

Low temperatures slow down the metabolism. Consequently, it has been demonstrated that modest hypothermia protects brain energy reserves: in young animals, lowering body temperature by six degrees, from 37 °C to 31 °C, preserves ATP levels to around 80% of normal levels over 4 h of controlled hypoxia⁹. P-creatine levels are similarly preserved, as well as the overall phosphorylation potential⁹. This suggests that lowering body temperature prior to decapitation could be neuroprotective, as near-normal levels of ATP could be maintained through the slice cutting and slice recovery periods.

To the degree that an ATP drop cannot be completely prevented upon decapitation, a partially impaired function of the Na-K ATPase is expected, followed by depolarization via passive sodium influx. As the passive sodium influx is followed by water entry into cells, it causes cytotoxic edema and eventually pyknosis. In adult rats, replacing Na+ ions with sucrose in slice-cutting solutions has been a successful strategy to alleviate the burden of cytotoxic edema^10,11. More recently, methylated organic cations that decrease sodium channel permeability¹² have shown to offer more effective protection than sucrose, especially in slices from adult mice, with N-methyl-D-glucamine (NMDG) being most widely applicable across different ages and brain regions^13,14,15,16.

Numerous brain-slicing protocols involve using cold temperatures only during the slice-cutting step, sometimes in combination with Na⁺ ion replacement strategy^16,17. In young animals, these protocols appear to offer sufficient neuroprotection since the brains can be extracted quickly after decapitation because the skull is still thin and easy to remove³. However, this strategy does not produce healthy slices from adult animals. Over time, a number of laboratories studying adult rodents have introduced transcardial perfusion with an ice-cold solution to decrease the animal’s body temperature, and therefore hypoxic damage to the brain, prior to decapitation. This procedure was successfully applied to produce cerebellar slices¹⁸, midbrain slices¹⁹, neocortical slices^11,20, perirhinal cortex²¹, rat hippocampus^10,22,23, olfactory bulb²⁴, ventral striatum²⁵, visual cortex²⁶.

In spite of the advantages offered by transcardial perfusion and Na⁺ ion replacement in preparing slices from rat and in some brain regions in mice, mouse hippocampus remains one of the most challenging areas to protect from hypoxia^13,20. To date, one of the most common approaches to slicing hippocampus from aging mice and mouse models of neurodegeneration involves the classical fast slicing of the isolated hippocampi²⁷. In the protocol described here, we minimize the loss of ATP in the adult brain by introducing hypothermia prior to decapitation by transcardially perfusing the animal with ice-cold Na⁺- free NMDG-based artificial cerebrospinal fluid (NMDG-aCSF). Slices are then cut in ice-cold Na⁺-free NMDG-aCSF. With this enhanced protocol we obtain acute hippocampal slices from adult and aging mice that are healthy for up to 10 h after slicing and are appropriate for long-term field-recordings and patch-clamp studies.

Protocol

The protocol is carried out in accordance with the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health and approved by the Stanford University Institutional Animal Care and Use Committee. Methods are also in accordance with the Policies of the Society for Neuroscience on the Use of Animals and Humans in Neuroscience Research.

NOTE: All mice were maintained in a pathogen-free environment. Wild-type mice on mixed C57Bl/ 6 x SV/ 129J genetic background were used here, unless otherwise noted.

1. Setup

1. Prepare 1 L of 1x aCSF (in mM): 125 NaCl, 26 NaHCO₃, 2.3 KCl, 1.26 KH₂PO₄, 1.3 Mg₂SO₄·7H₂O, 2.5 CaCl₂, 25 glucose (Table 1). Use this solution for the recovery chamber and subsequent recordings.
   NOTE: Store aCSF as a 10x stock solution that contains NaCl, NaHCO₃, KCl, and KH₂PO₄. Store Mg₂SO₄·7H₂O and CaCl₂ as 1 M stock solutions. Prepare the working solution on the day of experiment from the above stock solutions, and add glucose before adjusting the final volume with 18 MΩ water.
   NOTE: Depending on the brain region or mouse strain, chilled aCSF can be also used for transcardial perfusion with success^11,15,26.
2. Prepare 300 mL of NMDG-aCSF (in mM): 135 NMDG, 1 KCl, 1.2 KH₂PO₄, 1.5 MgCl₂, 0.5 CaCl₂, 20 choline bicarbonate, 10 glucose (Table 1). This volume of NMDG-aCSF is sufficient for both transcardial perfusion and cutting steps.
   NOTE: Store NMDG-aCSF as a 3x stock solution at 4 °C. Prepare working solution on the day of the experiment; add choline bicarbonate and glucose before adjusting the final volume with 18 MΩ water. Bubble the solution with 95%O₂/5%CO₂ to buffer it, and saturate with oxygen.
   NOTE: NMDG stock starts as highly alkaline, and requires concentrated hydrochloric acid (HCl) to adjust pH to ~7.4. Addition of hydrochloric acid should be slow once below pH 8, as it is easy to acidify the solution to ~pH 3 with excess HCl. This adjustment should be done prior to adding divalent cations. This NMDG-aCSF recipe is sodium-free. Previously published NMDG recipes use 30 mM NaHCO₃ for buffering, which results in 30 mM sodium present in NMDG-aCSF^13,20.
3. Set the vibrating microtome cutting tray and mounting disk to -20 °C.
4. Prepare the recovery chamber.
   1. Fill the recovery chamber to just above the slice-holding mesh, keep it on the bench at room temperature, and bubble.
      NOTE: The slice recovery chamber used here is very similar to the classical chambers described previously by Edwards and Konnerth³. aCSF is kept in a glass beaker (400 mL) that holds a round acrylic frame with black nylon mesh glued to the bottom. The acrylic frame is suspended in the middle of the beaker via an acrylic hook resting over the edge of the beaker. The glass bubbler is inserted all the way to the bottom. The design allows for oxygenation of both sides of slices. Bubbling also provides constant mixing of aCSF in the recovery chamber. The black mesh provides a high contrast against the off-white slices, which are then easier to see.
5. Prepare the transcardial perfusion and cutting solution.
   1. Chill the entire 300 mL of NMDG-aCSF in a freezer, until ice crystals start to form on the surface and the walls of the bottle. DO NOT over-freeze!
   2. Place the bottle with chilled NMDG-aCSF on ice and bubble. The solution should be between 0‒2 °C.
      NOTE: Slushy NMDG- aCSF implies having most of the solution as liquid, while a small fraction of slushy ice present will keep the solution near 0 °C throughout perfusion and cutting. Care should be taken to keep away ice crystals from the brain during cutting.
6. Prepare the tissue-mounting disk.
   1. Take the disk out of the freezer, wipe it dry if needed.
   2. Cut out a block of 5% agar from the previously prepared agar plate, and glue it in the center of the disk using a thin layer of cyanoacrylate glue. The agar piece should be about the size of a mouse brain. Place the disk with glued agar on ice, and cover with paper towels until it is ready to use.
      1. For 5% agar plate, dissolve 5 g of agar in 100 mL of 18 MΩ water by microwaving, and pour into a clean petri dish. Keep at 4 °C.
7. Take the cutting tray out of the freezer, place it into the microtome, surround it with ice, and load the blade.

2. Transcardial perfusion and brain extraction

1. Overdose mice via an intraperitoneal injection of anesthetic cocktail. Check for the anesthesia depth by checking the pain reflex (toe pinch); a mouse should not exhibit the reflex once it reaches deep anesthesia.
   NOTE: Rodent anesthesia cocktail recipe: Ketamine HCl (66 mg/mL), xylazine HCl (6.6 mg/mL), acepromazine maleate (0.1 mg/mL), 18 MΩ water to final volume. Dose: 0.4 mg/g body weight, 0.04 mg/g body weight, 6 x 10^-4 mg/g body weight for ketamine, xylazine, and acepromazine, respectively.
2. Set-up the peristaltic pump for transcardial perfusion. Insert one side of the pump tubing into the bottle with iced NMDG-aCSF. Fit the other side of the tubing with the 27 G needle that will be inserted into the left ventricle.
3. Set the pump speed at approximately 3.5 mL/min. At this speed, the outflow of NMDG-aCSF is a fast drip, not continuous flow.
   NOTE: Perfusion by gravity is a good substitute for the peristaltic pump perfusion, as long as the same approximate flow-rate can be achieved. A perfusion at high flow-rate will result in burst blood vessels in the brain. A telltale sign of an overly fast perfusion and increased pressure in blood vessels is solution coming out of the nose of the animal.
4. Perfusion
   1. Place the properly anesthetized mouse on its back on a diaper. Using paper tape, tape down its front and hind legs so that the chest and abdomen are exposed.
   2. Cut out a large patch of the skin atop of the chest, going from below the sternum to the throat; this should provide a large working area. Grab the sternum with forceps, lift gently, and start cutting through the rib cage on both sides until the chest cavity is exposed.
   3. Cut through the diaphragm in order to expose the chest cavity. The flap of the rib cage should be left attached via a thin piece of muscle. It should be possible to set it on a side without having it fall back onto the exposed chest cavity. Check that the heart is still beating. Ensure that most of the liver is visible.
   4. Insert the 27 G needle into the left ventricle; the mouse’s left ventricle looks lighter in color than the right. Locate the dark red-colored right atrium. Cut through the right atrium with small scissors; the blood should start flowing out.
   5. Start the pump that has been preset to the correct flow speed. If everything was done correctly, the liver should start to change color from red to brown soon after the start of perfusion. Monitor the liver color to determine the length of the perfusion; the liver should turn pale brown.
   6. Run the pump for few more minutes. If using rectal thermometer, the body temperature should fall down to 28‒29 °C, and animal’s nose should be cold to the touch.
5. Brain extraction.
   NOTE: For this step, have the following dissection tools ready: decapitation scissors, small scissors with straight or angled blade, scalpel and #10 blade, single edge blade, #3 forceps, spatula with one side bent to 90°, a spatula, a “60°” tool (Figure 1A,B), and a small soft brush.
   1. Decapitate the mouse with large decapitation scissors. Using a scalpel with #10 blade, cut open the skin on top of the skull. With small angled scissors, cut the skull at the midline. Next, using the #3 forceps, pry away the right and left halves of the skull, being careful to take the dura away with it. The brain should be exposed now.
      NOTE: If the dura detaches from the skull, it will stay over the brain and has to be removed separately. The edges of the remaining dura can tighten and slice deep through the brain, potentially damaging the brain region of interest.
   2. Remove the brain by scooping it out with a small spatula. Drop the brain into the NMDG-aCSF solution placed in a separate small beaker on ice. Leave it there for up to a minute.
      NOTE: In addition to hypothermia, exposure to the ice-cold saline firms up the brain, which is necessary for even cutting. The entire procedure from decapitation to the brain extraction should be under 30 s.

3. Slicing

1. Take the brain out of the NMDG-aCSF and place it on a piece of filter paper.
2. Cut and remove a 60° wedge from the rostral end of the forebrain using a “60°” tool centered at the midline. Cut surfaces will be used for mounting to achieve the proper angle for transversal hippocampal slices as discussed below (Figure 1A,B).
   NOTE: Two single-edged blades held together via a home-made holder make an easy-to-use tool for making the 60° cut (Figure 1A).
3. Separate hemispheres down the midline with the scalpel.
4. Glue the hemispheres onto the mounting disk as follows. Take the mounting disk that has been on ice until now. If needed, wipe it dry again. Glue each hemisphere in front of the agar block, cut side down.
5. Ensure that the ventral side of each hemisphere is touching the agar block. The agar block provides additional support during cutting and is essential for even slices. Dorsal sides of each hemisphere should be facing the blade. When glued on the cut side, each hemisphere is oriented relative to the blade in a way that results in transverse slices of dorsal hippocampus in situ (Figure 1B).
6. Submerge the disk with hemispheres into a cutting chamber containing ice-cold carbogenated NMDG-aCSF.
7. Cut 400 µm sections. Cutting should be done in less than 10 min. Different microtomes will require different settings to achieve this time. A total of 8‒10 slices from the dorsal hippocampus region will be obtained.

4. Recovery

1. Transfer slices to a recovery chamber containing carbogenated aCSF at room temperature using disposable transfer pipettes with cut-off tips (Figure 1C).
   NOTE: It is highly recommended to fortify recovery chamber-aSCF with 5 mM Na-ascorbate and 3 mM Na-pyruvate. Pyruvate is an energy substrate shown to boost the production of ATP in slices²⁸, while Na-ascorbate is a free-radical quencher²⁹.
2. Incubate at room temperature (22‒24 °C) for approximately 2 h before recording (up to 4 h).
   NOTE: A longer incubation at room temperature results in healthier slices for longer periods of time, relative to the standard incubation at 37 °C for 30 min followed by room temperature incubation. Rewarming at 37 °C can introduce cytotoxic edema¹³.

Results

We applied the above protocol to generate hippocampus slices from CamKIIa-Cre+;WT mice on a mixed genetic background C57Bl/ 6 x SV/ 129J, at P > 120. Large numbers of pyramidal cells in the CA1 field (Figure 2A) and subiculum (Figure 2B) appear in low contrast when observed under infrared differential contrast microscopy (IR-DIC), a hallmark of healthy cells in a slice preparation. With this preparation, a high rate of giga-ohm seals (>90%) is routi...

Discussion

The protocol described here demonstrates that hippocampal slices obtained from adult and aging mice can remain healthy and viable for many hours after cutting. The slices prepared using this protocol are appropriate for patch-clamp recordings, as well as long-lasting field-recordings in the CA1 regions.

There are two critical steps in this protocol. First step is the transcardial perfusion step with an ice-cold solution. Fast clearance of blood is signaled by rapid change of liver color. ...

Materials

Name	Company	Catalog Number	Comments
“60 degree” tool	made in-house
#10 scalpel blade	Bard-Parker (Aspen Surgical)	371110
1M CaCl2	Fluka Analytical	21114
95%O2/5%CO2	Praxiar or another local supplier
Acepromazine maleate (AceproJect)	Henry Schein	5700850
Agar	Fisher	BP1423-500
Beakers, measuring cylinders, reagent bottles
Brushes size 00-2	Ted Pella		Crafts stores are another source of soft brushes, with larger selection and better quality than Ted Pella.
CCD camera	Olympus	XM10
Choline bicarbonate	Pfalz & Bauer	C21240
Cyanoacrilate glue	Krazy glue		Singles
Decapitation scissors	FST	14130-17
Feather blades	Feather	FA-10
Filter paper #2	Whatman		Either rounds or pieces cut from a bigger sheet work well.
Forceps	A. Dumont & Fils	Inox 3c
Glass bubblers (Robu glass borosillicate microfilter candles) - porosity 3	Robuglas.com	18103 or 18113	Glass bubblers are more expensive than bubbling stones used in aquaria. However, they are easy to clean and sterilize, and can last a long time.
Glucose	Sigma-Aldrich	G8270
HCl	Fisher	A144SI-212
Ice buckets
KCl	Sigma-Aldrich	P4504
Ketamine HCl (KetaVed)	VEDCO	NDC 50989-996-06
KH2PO4	Sigma-Aldrich	P0662
Leica Tissue slicer VT1000S			The cutting settings are 1 mm horizontal blade amplitude, frequency dial at 9, and speed setting at 2
Magnetic stirrers and stir bars
Mg2SO4 x 7H2O	Sigma-Aldrich	230391
MgCl2	Sigma-Aldrich	M9272
MilliQ water machine	Millipore		Source for 18 Mohm water
Na-ascorbate	Sigma-Aldrich	A4035
Na-pyruvate	Sigma-Aldrich	P8574
NaCl	Sigma-Aldrich	S3014
NaHCO3	EMD	SX0320-1
Needle 27G1/2
NMDG	Sigma-Aldrich	M2004
Paper tape
Peristaltic pump	Cole-Parmer	#7553-70
Peristaltic pump head	Cole-Parmer	Masterflex #7518-00
Personna blades		Personna double edge	Amazon
pH meter
Recovery chamber	in-house made
Scalpel blade handle size 3	Bard-Parker (Aspen Surgical)	371030
Scissors angled blade	FST	14081-09
Single edge industrial razor blade #9	VWR	55411
Spatulas
Transfer pipettes	Samco Scientific	225
Upright microscope	Olympus BX51WI
Xylazine HCl (XylaMed)	VetOne	510650