Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

Procedures involving transgenic mice have been approved by the Institutional Animal Care and Use Committee (IACUC) at the Allen Institute for Brain Science. Both male and female C57BL/6 mice (weight range 10-30 g) were used in these experiments. Some of the representative results describe data collected from living human brain slices. Neocortical tissue specimens were obtained during neurosurgeries for tumor removal. It was necessary to remove the overlying neocortical tissue to gain access to the diseased tissue. Informed patient consent was obtained in all cases for the use of neurosurgical tissue for research purposes under a protocol approved by the institutional review board of Swedish Medical Center.

1. Preparation of Media and Reagents (Table 1)

NOTE: Solutions should be made up in purified water that is free of trace metals and other impurities. It is recommended that solutions be made freshly on the day of the experiment, although unused solutions may be stored at 4 °C for up to 1 week, if desired. 1 L of each formulation above is sufficient for 1–2 slicing procedures. All aCSF solutions must be saturated with carbogen (95% O₂/5% CO₂) prior to use to ensure stable pH buffering and adequate oxygenation. The pH of all solutions should be adjusted to 7.3-7.4 and osmolality measured and adjusted to 300-310 mOsmol/kg.

1. Prepare NMDG-HEPES aCSF (in mM): 92 NMDG, 2.5 KCl, 1.25 NaH₂PO₄, 30 NaHCO₃, 20 HEPES, 25 glucose, 2 thiourea, 5 Na-ascorbate, 3 Na-pyruvate, 0.5 CaCl₂·2H₂O, and 10 MgSO₄·7H₂O. Titrate pH to 7.3–7.4 with 17 mL +/- 0.5 mL of 5 M hydrochloric acid.
   NOTE: This titration step should ideally be performed prior to the addition of divalent cations to avoid precipitation; however, the precipitation can be reversed upon adjustment of the pH to the physiological range.
2. Prepare HEPES holding aCSF (in mM): 92 NaCl, 2.5 KCl, 1.25 NaH₂PO₄, 30 NaHCO₃, 20 HEPES, 25 glucose, 2 thiourea, 5 Na-ascorbate, 3 Na-pyruvate, 2 CaCl₂·2H₂O, and 2 MgSO₄·7H₂O. Titrate pH to 7.3–7.4 with several drops of concentrated 10 N NaOH.
3. Prepare recording aCSF (in mM): 124 NaCl, 2.5 KCl, 1.25 NaH₂PO₄, 24 NaHCO₃, 12.5 glucose, 5 HEPES, 2 CaCl₂·2H₂O, and 2 MgSO₄·7H₂O. Titrate pH to 7.3-7.4 with a few drops of concentrated 10 N NaOH.
4. Prepare Na⁺ spike-in solution (2 M): 580 mg of NaCl dissolved in 5 mL of freshly prepared NMDG-HEPES aCSF. This is enough solution for one brain slice prep.
5. Prepare 2% agarose to be used for tissue embedding. Dissolve 2 g of agarose type 1B (see Table of Materials) in 100 mL of 1x PBS and microwave until just boiling. Swirl to mix, then pour the mixture into a sterile 10 cm Petri dishes and allow to solidify. Store the agarose plate in a sealed plastic bag at 4 °C until use.
6. Prepare injectable anesthetic working stock solution. Mix 2.5 g of 2,2,2-Tribromoethanol with 5 mL of 2-methyl-2-butanol and then gradually dissolve in 200 mL PBS, pH 7.0–7.3. Filter the stock solution with a 0.22 µm filter before use and store at 4 °C protected from light.
   NOTE: Consult respective animal use committee guidelines and rules for determining the expiration and disposal procedure for the anesthetic working stock solution.

2. Setup of the Slicing Station

1. Set up the slicing station with the tissue slicer machine and surgical instruments (see Table of Materials). To calibrate the slicer machine, attach a zirconium ceramic injector blade to the blade arm using fast-adhesive glue, then insert the specimen holder and align the leading edge of the blade to the specimen holder rim leaving a tiny gap to ensure the blade does not scrape the metal.
   NOTE: If the blade edge is not physically damaged it can be reused for many weeks or even months without replacement.Various tissue slicer models are commercially available, many of which can provide excellent performance when optimally calibrated. The ideal instrument should have minimal z-axis deflection, either directly measured and tuned or empirically observed.
2. Set up a 250 mL beaker filled with 200 mL of NMDG-HEPES aCSF and pre-chill on ice with constant carbogenation (applied via a gas diffuser stone immersed in the media) for >10 min.
   NOTE: This solution will be used for transcardial perfusion and for filling the slicing machine reservoir during sectioning.
3. Set up the initial brain slice recovery chamber filled with 150 mL of NMDG-HEPES aCSF (maintain constant carbogenation) and place the chamber in a heated water bath maintained at 32–34 °C.
   NOTE: A slice chamber after the design of Edwards and Konnerth (1992)⁵² is recommended for this step. These chambers can be made with readily available laboratory items (250 mL beaker, nylon mesh netting, 50 mL conical tube, 35 mm plastic round dish). Care must be taken to ensure that the slice netting remains free of air bubbles, particularly those continuously produced by the carbogen gas bubble stones as these can cause slices to float up and become damaged. The netting should be submerged roughly 1 cm under the liquid surface.
4. Set up a brain slice holding chamber; a design with multiple independent wells in a larger reservoir is recommended (see Table of Materials). Fill the reservoir with 450 mL of HEPES aCSF and warm to room temperature under constant carbogenation until use.
   NOTE: The brain slices will be transferred from the initial recovery chamber to this chamber for long-term storage before electrophysiological recording. Care must be taken to ensure that the slice netting remains free of air bubbles at all times.
5. Prepare molten agarose for tissue embedding. Use the open end of a 50 mL conical vial like a cookie cutter to cut out a block of 2% agarose from the previously prepared dish. Loosely cap the conical vial, then microwave for 10–30 s until the agarose is just melted. Do not overheat.
6. Pour the molten agarose into 1.5 mL tubes. Maintain the agarose in the molten state using a thermomixer set to 42 °C with vigorous shaking. Carefully ensure that the molten agarose does not solidify prematurely.
7. Place the accessory chilling block for the slicer on ice to pre-cool at this time.

3. Transcardial Perfusion

NOTE: The transcardial perfusion procedure is an important step when working with adult animals and is important to achieve rapid cooling of the brain and slowed metabolism via brain infusion of low Na⁺, low Ca²⁺/high Mg²⁺ aCSF solution. The transcardial perfusion also serves to clear red blood cells from brain vasculature, which reduces autofluorescence that might interfere with visualization and targeting of fluorescently labeled cell populations in transgenic lines. It is not advisable to omit transcardial perfusion.

1. Deeply anesthetize mice by intraperitoneal injection of anesthetic working stock solution (250 mg/kg: 0.2 mL of 1.25% anesthetic working stock solution per 10 g body weight, see the Table of Materials). After ~2–3 min, verify sufficient depth of anesthesia by assessing toe pinch reflex. If required, inject an additional volume of anesthetic working stock and reassess toe pinch reflex after another 2-3 min.
2. Load a 30 mL syringe with 25 mL of carbogenated NMDG-HEPES aCSF from the pre-chilled 250 mL beaker (2–4 °C is optimal, as opposed to slushy or frozen solution). Attach a 25 5/8 gauge needle.
3. With the mouse on its back, pin down the forepaws and hind paws for stability. A 15 cm glass Petri dish filled with hardened silicone works well as the base.
4. Using a scalpel, make a lateral incision to open the thoracic cavity at the level of the diaphragm. Use fine scissors to cut through the rib cage on either side taking care to avoid clipping the heart and lungs.
5. Pin back the center portion of the rib cage to expose the heart. Insert the needle of the 30 mL syringe into the left ventricle and cut the right atrium with fine scissors to allow blood to exit the heart.
6. Depress the syringe plunger using manual constant pressure and perfuse the animal with the chilled NMDG-HEPES aCSF at a rate of ~10 mL/min.
   NOTE: If the perfusion is successful the liver will change in color from deep red to pale yellow, and in some cases clear fluids can be observed exiting the nostrils towards the end of the procedure.

4. Brain Dissection and Slicing

1. Decapitate the animal. Use a scalpel to open the skin on the head and expose the skull cap.
2. Use fine super-cut scissors to cut away the skin over the skull cap and make small incisions laterally on either side at the caudal/ventral base of the skull. Make additional shallow cuts starting at the caudal/dorsal aspect of the skull moving in the rostral direction up the dorsal midline taking care not to damage the underlying brain. Make a final 'T' cut perpendicular to the midline at the level of the olfactory bulbs.
   NOTE: Care must be taken to ensure no damage is done to the brain region(s) of interest. In particular, at no time should there be any compressive force applied to the brain itself.
3. Use the round-tip forceps to grasp the skull starting at the rostral-medial aspect and peel back towards the caudal-lateral direction. Repeat for both sides to crack open and remove the dorsal halves of the skull cap to expose the brain. Gently scoop out the intact brain into the beaker of pre-chilled NMDG-HEPES aCSF. Allow the brain to uniformly cool for ~1 min.
4. Use the large spatula to lift the brain out of the beaker and onto the Petri dish covered with filter paper. Trim and block the brain according to the preferred angle of slicing and desired brain region of interest. Work quickly to avoid prolonged oxygen deprivation during handling.
   NOTE: Many slicing angles are possible. The exact blocking method and slicing angle will depend on the exact brain region, cell type, and circuit to be studied.
5. Affix the brain block to the specimen holder using adhesive glue. Retract the inner piece of the specimen holder enough to withdraw the brain block fully inside. Pour the molten agarose directly into the holder until the brain block is fully covered in agarose. Clamp the pre-cooled accessory chilling block around the specimen holder for ~10 s until the agarose has solidified.
6. Insert the specimen holder into the receptacle on the slicer machine and verify proper alignment. Fill the reservoir with remaining pre-chilled, oxygenated NMDG-HEPES aCSF from the 250 mL beaker and move a bubble stone into the reservoir for the duration of slicing to ensure adequate oxygenation.
7. Adjust the micrometer to begin advancing the agarose-embedded brain specimen. Start the slicer and empirically adjust the advance speed and oscillation frequency to the desired level.
   NOTE: Both settings should be in the low range. For best results, a single pass of the blade arm should take approximately 20 s and the oscillation should produce a very smooth and gentle humming noise with no overt buzzing.
8. Continue advancing and slicing the tissue in 300 µm increments (or other preferred thickness) until the brain region of interest is fully sectioned; the total time for the slicing procedure should be less than 15 min.

5. Optimized NMDG Protective Recovery Procedure

1. Initial NMDG recovery step (critical step): Upon completion of the sectioning procedure, collect up all of the slices using a cut-off plastic Pasteur pipe t and transfer them into a pre-warmed (34 °C) initial recovery chamber filled with 150 mL of NMDG-HEPES aCSF. Transfer all slices in short succession and start a timer as soon as all slices are moved into the recovery chamber.
2. Consult Table 2 to determine the optimal Na⁺ spike-in schedule according to mouse age.
   NOTE: This procedure is a practical method to achieve a controlled rate of reintroduction of Na⁺ into the brain slice chamber and is optimized for a specific brain slice chamber geometry and reservoir volume and type (see Table of Materials).
3. Carry out the stepwise Na⁺ spike-in procedure by adding the indicated volumes of Na⁺ spike-in solution at the indicated times. Add the Na⁺ spike-in solution directly into the bubbler chimney of the initial recovery chamber to facilitate rapid mixing.
4. Transfer all slices to the HEPES aCSF long-term holding chamber maintained at room temperature. Allow slices to recover for an additional 1 h in the HEPES holding chamber prior to initiating patch clamp recording experiments.

6. Patch Clamp Recording

NOTE: The following basic procedures merely provide some practical considerations and are not intended to represent detailed protocols for patch clamp recordings, as these can be found elsewhere^53,54. A patch clamp electrophysiology rig is required for this application. This will generally be composed of an upright microscope equipped with infrared differential interference contrast (IR-DIC) optics and a fluorescence illumination system, a patch clamp amplifier and data digitizer, motorized micromanipulator and microscope platform, vibration isolation table, Faraday cage, and solution heating and perfusion system. The sample chamber and platform should be designed for submerged slice recording. For multi-neuron patch clamp recordings, a rig equipped with multiple amplifiers, head stages, and high quality micromanipulators is required. In addition, for best results, a rig equipped with a 900 nm IR band pass filter and matching optical components is highly recommended to ensure adequate visualization of cells located >50 µm deep in the brain slices. Proper alignment for Kӧhler illumination is also important for clear visualization.

1. Prepare intracellular pipette solution (in mM): 130 K-Gluconate, 4 KCl, 10 HEPES, 0.3 EGTA, 10 phosphocreatine-Na₂, 4 MgATP, 0.3 Na₂-GTP, and 13.4 biocytin. Adjust the pH to 7.35 with 1 M KOH and the osmolality to 285–290 mOsmol/kg using sucrose as needed.
2. Prepare patch clamp electrodes from thick-walled borosilicate glass capillaries; the ideal patch clamp electrode has a relatively short and stubby taper with a ~3–6 MOhm filled tip resistance in the bath.
3. Ensure that the silver electrode wires are properly chlorided in order to ensure stable recordings. Do this (typical) by submerging the last 3-4 mm of the silver wire in liquid bleach for approximately 30 min or until the wire turns black.
4. Establish solution perfusion using a peristaltic pump set to 3–4 mL/min. Circulate carbogenated recording aCSF through the recording chamber taking care to match inflow and outflow to avoid overflow.
5. Transfer a single brain slice into the submersion recording chamber and secure in place using a U-shaped slice anchor with nylon cross strings. Identify the target brain region using a 4X air objective before switching to a higher power objective (e.g., high numerical aperture 40X or 60X water immersion objectives).
6. Visually identify a healthy target cell. The appearance of the neuronal membrane at the soma, as visualized by IR-DIC microscopy, is used to judge the suitability of a candidate cell for patch clamp recording.
   NOTE: Healthy neurons typically exhibit the following features: neither shrunken nor swollen somata, soft contrast of membrane edges, and smooth membrane appearance. In addition, the majority of healthy cells are located >30 µm deep in the slice, as superficial neurons are likely to be damaged with severed dendritic processes. Neurons that exhibit a crinkled appearance, clearly visible nuclei or 'fried egg' appearance, or very dark or high contrast membrane edges should be avoided.
7. Back-fill the patch pipette with ~5 µL of intracellular solution and load it onto the electrode holder. Move the pipette into the recording bath above the brain slice and apply light positive pressure to clear any obstructions in the tip. Zero the pipette offset and monitor the tip resistance using a membrane test function.
8. Move the pipette tip into contact with the target neuron's cell body; a small dimple should form on the membrane surface due to the light positive pressure.
9. As soon as a membrane dimple is observed, swiftly remove the positive pressure and apply gentle suction to facilitate seal formation. Once the pipette resistance increases to >100 MOhm, turn on a holding command to a level that matches the anticipated resting membrane potential for the targeted cell type (-70 mV is a good starting point).
10. Once the tip resistance reaches ≥1 gigaohm, attempt to break into the cell by rupturing the membrane underneath the patch pipette using brief bouts of sharp suction; the 'zap' feature may be utilized to facilitate break-in as needed.
    NOTE: A holding potential of -70 mV is suggested for voltage clamp experiments on cortical neurons. Healthy neurons will have a leak current no more negative than -100 pA for the duration of the experiment, but this is in part dependent on the cell type. A neuron would be excluded from analysis if the resting membrane potential was more depolarized than -50 mV, or if the access resistance changed by more than 20%.
11. Once a stable whole-cell patch clamp recording is obtained, target additional neurons for recording by repeating steps 6.6–6.10. Take care to avoid mechanical disturbances that would result in losing the first recording.
    1. Select additional neurons within 100 µm from the first neuron to ensure a reasonable probability of finding synaptically-coupled neurons.
       NOTE: It may be helpful in some cases to identify multiple candidate neurons up front and to pre-load and pre-position all pipettes nearby to the various targeted cells before establishing the first patch clamp recording.