This work was funded by the Allen Institute for Brain Science. The authors wish to thank the Allen Institute founders, Paul G. Allen and Jody Allen, for their vision, encouragement, and support. We also thank the Allen Institute technical support staff for performing animal care, husbandry, and genotyping.

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 &#176;C for up to 1 week, if desired. 1 L of each formulation above is sufficient for 1&#8211;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.
<ol>
	<li>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&#183;2H2O, and 10 MgSO4&#183;7H2O. Titrate pH to 7.3&#8211;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.</li>
	<li>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&#183;2H2O, and 2 MgSO4&#183;7H2O. Titrate pH to 7.3&#8211;7.4 with several drops of concentrated 10 N NaOH.</li>
	<li>Prepare recording aCSF (in mM): 124 NaCl, 2.5 KCl, 1.25 NaH2PO4, 24 NaHCO3, 12.5 glucose, 5 HEPES, 2 CaCl2&#183;2H2O, and 2 MgSO4&#183;7H2O. Titrate pH to 7.3-7.4 with a few drops of concentrated 10 N NaOH.</li>
	<li>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.</li>
	<li>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 &#176;C until use.</li>
	<li>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&#8211;7.3. Filter the stock solution with a 0.22 &#181;m filter before use and store at 4 &#176;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.</li>
</ol>
2. Setup of the Slicing Station
<ol>
	<li>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.</li>
	<li>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 &#62;10 min. 
	NOTE: This solution will be used for transcardial perfusion and for filling the slicing machine reservoir during sectioning.</li>
	<li>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&#8211;34 &#176;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.</li>
	<li>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.</li>
	<li>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&#8211;30 s until the agarose is just melted. Do not overheat.</li>
	<li>Pour the molten agarose into 1.5 mL tubes. Maintain the agarose in the molten state using a thermomixer set to 42 &#176;C with vigorous shaking. Carefully ensure that the molten agarose does not solidify prematurely.</li>
	<li>Place the accessory chilling block for the slicer on ice to pre-cool at this time.</li>
</ol>
3.&#160;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.
<ol>
	<li>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&#8211;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.</li>
	<li>Load a 30 mL syringe with 25 mL of carbogenated NMDG-HEPES aCSF from the pre-chilled 250 mL beaker (2&#8211;4 &#176;C is optimal, as opposed to slushy or frozen solution). Attach a 25 5/8 gauge needle.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>
4.&#160;Brain Dissection and Slicing
<ol>
	<li>Decapitate the animal. Use a scalpel to open the skin on the head and expose the skull cap.</li>
	<li>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 &#39;T&#39; 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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Continue advancing and slicing the tissue in 300 &#181;m 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.</li>
</ol>
5. Optimized NMDG Protective Recovery Procedure
<ol>
	<li>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 &#176;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.</li>
	<li>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).</li>
	<li>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.</li>
	<li>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.</li>
</ol>
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 elsewhere53,54. 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 &#62;50 &#181;m deep in the brain slices. Proper alignment for K&#1255;hler illumination is also important for clear visualization.
<ol>
	<li>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&#8211;290 mOsmol/kg using sucrose as needed.</li>
	<li>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&#8211;6 MOhm filled tip resistance in the bath.</li>
	<li>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.</li>
	<li>Establish solution perfusion using a peristaltic pump set to 3&#8211;4 mL/min. Circulate carbogenated recording aCSF through the recording chamber taking care to match inflow and outflow to avoid overflow.</li>
	<li>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).</li>
	<li>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 &#62;30 &#181;m 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 &#39;fried egg&#39; appearance, or very dark or high contrast membrane edges should be avoided.</li>
	<li>Back-fill the patch pipette with ~5 &#181;L 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.</li>
	<li>Move the pipette tip into contact with the target neuron&#39;s cell body; a small dimple should form on the membrane surface due to the light positive pressure.</li>
	<li>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 &#62;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).</li>
	<li>Once the tip resistance reaches &#8805;1 gigaohm, attempt to break into the cell by rupturing the membrane underneath the patch pipette using brief bouts of sharp suction; the &#39;zap&#39; 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%.</li>
	<li>Once a stable whole-cell patch clamp recording is obtained, target additional neurons for recording by repeating steps 6.6&#8211;6.10. Take care to avoid mechanical disturbances that would result in losing the first recording.
	<ol>
		<li>Select additional neurons within 100 &#181;m 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.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

Na+ Spike-in Improves Gigaohm Seal Formation and Patch Clamp Recording Success 
The initial version of the NMDG protective recovery method was specifically designed for adult and aging animals2,5. Some early adopters have also sought to apply this methodology to juvenile animal brain slicing (i.e., mice &#60;30 days old). However, it has been noted that in contrast to the outstanding visually-confirmed neuronal preservation with the NMDG protective recovery method in this age range, gigaohm seal formation can frequently stall out, leading to failed patch clamp recording attempts. One hypothesis is that NMDG cations are more readily trapped in juvenile brain slices relative to adult brain slices and can impede seal formation; however, gigaohm seals can readily form while juvenile brain slices are fully submerged in NMDG aCSF (data not shown), thus indicating that NMDG aCSF per se is not impeding gigaohm seal formation.
The rapid transition from low-to-high Na+ solution at the completion of the initial brain slice recovery step causes damage to neuronal membranes and perturbs the seal formation process. This is intuitive given that the transition from low-to-high Na+, cold-to-warm temperature, and dramatic elevation of the Ca2+ to Mg2+ ratio collectively lead to a massive resurgence of spontaneous synaptic activity. This inhibitory rebound phase in the brain slicing procedure is likely to mirror reperfusion injury following an ischemic insult. Thus, to further mitigate neuronal membrane damage in the initial recovery phase a gradual Na+ spike-in procedure has been incorporated in which the elevation of Na+ concentration in the NMDG protective recovery incubation chamber is slowly and reproducibly elevated with precise timing. As in the original protective recovery procedure, the temporal dissociation of Na+ elevation from temperature and Ca2+/Mg2+ ratio elevation is beneficial. But additionally, the Na+ spike-in procedure leads to small incremental increases in extracellular Na+ concentration over the early time points and large increases towards the late time points, thereby affording the brain tissue an opportunity to better accommodate to the rising Na+ levels. This procedure is an alternative to gradual solution exchange controlled by a perfusion pump or gravity drip lines which lead to constant increases in Na+ levels and require attention to both inflow and outflow to avoid overflow of the slice chamber. Notably, in this Na+ spike-in procedure the osmolality of the solution in the slice chamber gradually rises over a period of several minutes before the slices are returned to normal osmolality solution, but this did not adversely affect the slice health or patch clamp recording success. A high osmolality cutting solution has previously been used for midbrain slice preparations in order to better preserve dopamine neurons for patch clamp recordings57,58, thus demonstrating that this temporary hyperosmolality may be beneficial in some contexts.
By implementing an optimized procedure combining the NMDG protective recovery method and gradual Na+ spike-in step the utility of this brain slice methodology has been extended to cover juvenile through mature adult animal ages. This updated protocol is now suitable for a wide range of animal ages using a single optimal NMDG aCSF formulation and procedure. If necessary, the Na+ spike-in procedure can be applied with a progressively longer delay and/or slower time course to enhance the viability of brain slices from older animals, and we have provided a basic guide of recommended spike-in schedules according to animal age (see Table 2). While we have provided a basic framework suitable for a wide array of applications, additional advanced steps can be explored for further enhancing viability and longevity of brain slices from adult and aging animals. For example, glutathione restoration strategies are particularly effective in this respect and can be implemented as described elsewhere2,6.
Improving Throughput for Challenging Experiments 
The analysis of synaptic connectivity by patch clamp recording is a demanding application that requires excellent preservation of both neuronal structure and function in order to achieve a high reliability of success. As the number of neurons to be simultaneously recorded goes up linearly, the technical difficulty level goes up supra-linearly. There are numerous failure modes, and one of the most frequent causes of failures is the inability to form adequate gigaohm seals onto one or more of the targeted cells. This can dramatically slow progress, particularly when three or more neurons must be recorded simultaneously. Consistent with the finding of faster gigaohm seal formation time with the optimized NMDG protective recovery method, there was a marked improvement in the success rate and throughput of multi-neuron patch clamp recording experiments with both adult transgenic mouse brain slices and adult human neurosurgical brain slices. The improved efficiency is almost certainly attributable to both the more rapid and reliable gigaohm seal formation and the improved neuronal preservation of the slices with this protocol. Although this protocol focuses on the benefits explicitly for patch clamp recording applications, similar gains are anticipated for other challenging experimental applications where brain slice viability is paramount.

The acute brain slice preparation is an essential experimental model system in neuroscience. For roughly half of a century, this platform has enabled dynamic functional studies of the living brain across a wide variety of anatomical brain regions and animal species. Whether the intended application is biochemistry, functional imaging, morphology, or electrophysiology, it is of the utmost importance to ensure optimal integrity and viability of the sliced tissue. It is for this reason that the highly resilient juvenile rodent brain slice preparation (i.e., younger than postnatal day 30 for mice) has been the most preferred to date. The difficulty in obtaining sufficiently healthy brain slices from mature adult and aging animals has proven to be a formidable challenge for most and has imposed severe limitations for studying the functional architecture of the mature brain. This is particularly true for patch clamp recording, a technique that requires excellent morphological and functional preservation and is indispensable for characterizing detailed intrinsic and synaptic properties of identified single neurons. For the past several decades, the vast majority of patch clamp electrophysiologists have relied on a &#39;protective cutting&#39; method using sucrose-substituted low Na+ aCSF1 for preparing healthy brain slices from juvenile, and to a far lesser extent, young adult animals. This method is based on the premise that passive Na+ influx and subsequent water entry and cell swelling during the slice cutting step is the predominant insult that leads to poor survival of neurons, particularly for those neurons located in the superficial layers that are most likely to sustain direct trauma from the blade movement. However, the protective cutting method still leaves much to be desired for brain slice preparation from mature adult animals regardless of the particular aCSF formulation implemented.
A simple but effective solution to this problem has been described2,3,4,5,6 and termed the &#39;protective recovery&#39; brain slice method. The original version of this method uses an NMDG-substituted aCSF, as NMDG was identified as the most versatile and effective among various other candidate sodium ion substitutes (including sucrose, glycerol, choline, and Tris). The media formulation was further enhanced by addition of HEPES to resist brain slice edema and provide stronger pH buffering7, as well as the addition of supplements to counteract the detrimental effects of oxidative stress (Table 1). It was empirically determined that an initial recovery incubation step in low Na+, low Ca2+, and high Mg2+ NMDG aCSF immediately after adult brain tissue slicing was both necessary and sufficient for improved neuronal preservation over a wide range of brain regions, cell types, and animal ages3,5,6.
Notably, earlier incarnations of what is now dubbed the protective recovery method can be found in the literature1,8,9,10,11,12,13, although the full potential for mature adult and aging animal brain slice and patch clamp recording was not recognized or demonstrated in these earlier works. In addition, nuanced procedural variations continue to emerge in support of specific experimental applications4,14,15,16. The collective body of work of these numerous research groups imparts high confidence in the robustness of the protective recovery method for improved tissue preservation. The NMDG protective recovery method has now been widely adopted and implemented in numerous published research studies utilizing adult animal brain slice preparations. These acute slice studies span neocortical3,17,18, hippocampal15,19,20,21, striatal22,23,24, midbrain25,26,27,28,29, and hindbrain30,31,32,33,34 regions, and a variety of neurotransmitter and neuromodulator types including glutamatergic4,30, GABAergic18,20,31,35,36, dopaminergic24,29,37,38, cholinergic14,37,38,39, noradrenergic40, and serotonergic27,28 neurotransmission. The method is also well suited for optogenetic control of neuronal activity in slices derived from transgenic animals3,39 or following in vivo viral injections17,27,28,40,41,42,43, as well as functional Ca2+ imaging of neuronal activity2,44,45,46. Analyses of both short term plasticity4,47,48 and diverse forms of long-term plasticity16,35,48 have been reported. A recent study applied the NMDG protective recovery method to facilitate extensive and systematic probing of synaptic connectivity in the visual cortex in mature adult mouse brain slices using the octopatch recording configuration49&#160;&#8212; a powerful demonstration of the utility and robustness of this method. The protective recovery method has even been applied successfully in previously unforeseen experimental contexts, such as, improved preservation of vasculature and pericytes in adult cortical brain slices50, patch clamp recording from transplanted interneuron populations in 1&#8211;1.5 year old Alzheimer&#39;s Disease mouse models20, and an adult brain slice receptor trafficking assay51.
The following protocol describes step-by-step procedures for implementing an optimized NMDG protective recovery method of brain slice preparation to improve the viability of acute brain slices. The principles for improved neuronal preservation are discussed, as well as demonstration of the clear benefits of this methodology for complex multi-neuron patch clamp recording experiments in both young adult transgenic mouse brain slices and mature adult human neurosurgical brain slices. The following protocol has been validated for mice from 21 days old to more than one years old, as well as for human neurosurgical specimens derived from adult patients.

<table><tbody><tr><td>Compresstome VF-200</td><td>Precisionary Instruments</td><td>VF-200</td><td>Vibrating tissue slicer (recommended)</td></tr><tr><td>N-methyl-D-glucamine</td><td>Sigma Aldrich</td><td>M2004</td><td>aCSF constituent</td></tr><tr><td>Sodium Chloride</td><td>Sigma Aldrich</td><td>S3014</td><td>aCSF constituent</td></tr><tr><td>Potassium Chloride</td><td>Sigma Aldrich</td><td>P5405</td><td>aCSF constituent</td></tr><tr><td>Sodium Phosphate monobasic dihydrate</td><td>Sigma Aldrich</td><td>71505</td><td>aCSF constituent</td></tr><tr><td>Sodium Bicarbonate</td><td>Sigma Aldrich</td><td>S5761</td><td>aCSF constituent</td></tr><tr><td>HEPES</td><td>Sigma Aldrich</td><td>H4034</td><td>aCSF constituent</td></tr><tr><td>Glucose</td><td>Sigma Aldrich</td><td>G7021</td><td>aCSF constituent</td></tr><tr><td>Sodium Ascorbate</td><td>Sigma Aldrich</td><td>A4034</td><td>aCSF constituent</td></tr><tr><td>Thiourea</td><td>Sigma Aldrich</td><td>T8656</td><td>aCSF constituent</td></tr><tr><td>Sodium pyruvate</td><td>Sigma Aldrich</td><td>P5280</td><td>aCSF constituent</td></tr><tr><td>Calcium chloride dihydrate</td><td>Sigma Aldrich</td><td>C7902</td><td>aCSF constituent</td></tr><tr><td>Magnesium Sulfate heptahydrate</td><td>Sigma Aldrich</td><td>M1880</td><td>aCSF constituent</td></tr><tr><td>2,2,2-Tribromoethanol&amp;nbsp;</td><td>Sigma Aldrich</td><td>T48402</td><td>Anesthetic component 1</td></tr><tr><td>2-methyl-2-butanol&amp;nbsp;</td><td>Sigma Aldrich</td><td>240486</td><td>Anesthetic component 2</td></tr><tr><td>Curved blunt forceps</td><td>Fine Science Tools</td><td>11065-07</td><td>Brain dissection tools</td></tr><tr><td>Fine dissecting scissors (supercut)</td><td>Fine Science Tools</td><td>14058-09</td><td>Brain dissection tools</td></tr><tr><td>Large heavy duty scissors 7&#039;&#039;</td><td>Fine Science Tools</td><td>14000-18</td><td>Brain dissection tools</td></tr><tr><td>Metal spatula</td><td>Sigma Aldrich</td><td>Z511455-1PAK</td><td>Brain dissection tools</td></tr><tr><td>Razor blades</td><td>VWR</td><td>89031-954</td><td>Brain dissection tools</td></tr><tr><td>Brain Slice Keeper-4</td><td>Automate Scientific</td><td>S-BSK4</td><td>brain slice holding chamber</td></tr><tr><td>nylon netting&amp;nbsp;</td><td>Warner Instruments</td><td>64-0198</td><td>For building small slice recovery chambers</td></tr><tr><td>Pyrex glass beakers (250 mL)</td><td>VWR</td><td>89090-434</td><td>For building small slice recovery chambers</td></tr><tr><td>35 mm plastic dish, round</td><td>VWR</td><td>100488-376</td><td>For building small slice recovery chambers</td></tr><tr><td>Gas diffuser stones (10 &amp;micro;m)</td><td>Sigma Aldrich</td><td>59277</td><td>For constant carbogenation (fine bubbles)</td></tr><tr><td>Agarose Type I-B</td><td>Sigma Aldrich</td><td>A0576</td><td>For embedding brain specimens</td></tr><tr><td>Micro loader tips</td><td>Eppendorf</td><td>22491229</td><td>For filling patch clamp electrodes</td></tr><tr><td>Sylgard</td><td>VWR</td><td>102092-312</td><td>For making a custom dissecting platform</td></tr><tr><td>Hydrochloric acid&amp;nbsp;</td><td>Sigma Aldrich</td><td>H1758-100ML</td><td>For pH adjustment of media</td></tr><tr><td>Sodium Hydroxide&amp;nbsp;</td><td>Sigma Aldrich</td><td>221465-25G</td><td>For pH adjustment of media</td></tr><tr><td>Potassium Hydroxide</td><td>Sigma Aldrich</td><td>221473</td><td>For pH adjustment of media</td></tr><tr><td>Plastic transfer pipets 3 mL graduated</td><td>VWR</td><td>89497-676</td><td>For slice transfer</td></tr><tr><td>Zirconium ceramic injector blades</td><td>Cadence Specialty Blades</td><td>EF-INZ10</td><td>http://cadenceinc.com/</td></tr><tr><td>KG-33 borosilicate glass capillary w/filament</td><td>King Glass Company</td><td>custom quote</td><td>ID: 0.87mm, OD 1.50mm</td></tr><tr><td>Biocytin</td><td>Sigma Aldrich</td><td>B4261</td><td>Intern pipette solution</td></tr><tr><td>Phosphocreatine disodium</td><td>Sigma Aldrich</td><td>P7936</td><td>Intern pipette solution</td></tr><tr><td>Potassium Gluconate</td><td>Sigma Aldrich</td><td>G4500-100G</td><td>Intern pipette solution</td></tr><tr><td>EGTA</td><td>Sigma Aldrich</td><td>E3889</td><td>Intern pipette solution</td></tr><tr><td>Mg-ATP</td><td>Sigma Aldrich</td><td>A9187</td><td>Intern pipette solution</td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;-GTP</td><td>Sigma Aldrich</td><td>51120</td><td>Intern pipette solution</td></tr><tr><td>sucrose</td><td>Sigma Aldrich</td><td>S0389</td><td>Intern pipette solution</td></tr><tr><td>Heated water bath (2.5L)</td><td>VWR</td><td>13491-060</td><td>Miscellaneous</td></tr><tr><td>Filter paper rounds</td><td>VWR</td><td>28456-022</td><td>Miscellaneous</td></tr><tr><td>Cyanoacrylate glue</td><td>Amazon</td><td>B000BQRBO6</td><td>Miscellaneous</td></tr><tr><td>Glass petri dish</td><td>VWR</td><td>89000-326</td><td>Miscellaneous</td></tr><tr><td>10X Phosphate buffered saline</td><td>Sigma Aldrich</td><td>P5493</td><td>Miscellaneous</td></tr><tr><td>30 mL syringes</td><td>VWR</td><td>BD302832</td><td>Miscellaneous</td></tr><tr><td>1 mL syringes</td><td>VWR</td><td>BD-309628</td><td>Miscellaneous</td></tr><tr><td>25 5/8 gauge needles</td><td>VWR</td><td>89219-292</td><td>Miscellaneous</td></tr><tr><td>Thermomixer (w/1.5 mL tube block)</td><td>VWR</td><td>89232-908</td><td>To keep agarose molten&amp;nbsp;</td></tr></tbody></table>

preparation of acute brain slices using an optimized n-methyl-d-glucamine protective recovery method

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have validated the utility of this method for enhancing neuronal preservation and overall brain slice viability. The implementation of this technique by early adopters has facilitated detailed investigations into brain function using diverse experimental applications and spanning a wide range of animal ages, brain regions, and cell types. Steps are outlined for carrying out the protective recovery brain slice technique using an optimized NMDG artificial cerebrospinal fluid (aCSF) media formulation and enhanced procedure to reliably obtain healthy brain slices for patch clamp electrophysiology. With this updated approach, a substantial improvement is observed in the speed and reliability of gigaohm seal formation during targeted patch clamp recording experiments while maintaining excellent neuronal preservation, thereby facilitating challenging experimental applications. Representative results are provided from multi-neuron patch clamp recording experiments to assay synaptic connectivity in neocortical brain slices prepared from young adult transgenic mice and mature adult human neurosurgical specimens. Furthermore, the optimized NMDG protective recovery method of brain slicing is compatible with both juvenile and adult animals, thus resolving a limitation of the original methodology. In summary, a single media formulation and brain slicing procedure can be implemented across various species and ages to achieve excellent viability and tissue preservation.

Watch this Scientific Journal Video about Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method at JoVE.com

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

This protocol demonstrates the implementation of an optimized N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. A single media formulation is used to reliably obtain healthy brain slices from animals of any age and for diverse experimental applications.

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

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have ...

preparation-acute-brain-slices-using-an-optimized-n-methyl-d

Universidad San Sebastian

Research

JoVE Journal

Neuroscience

45.8K Views.  Allen Institute for Brain Science. This protocol demonstrates the implementation of an optimized N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. A single media formulation is used to reliably obtain healthy brain slices from animals of any age and for diverse experimental applications.

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

Strategies for Study of Neuroprotection from Cold-preconditioning

Neurological injury is a frequent cause of morbidity and mortality from general anesthesia and related surgical procedures that could be alleviated by development of effective, easy to administer and safe preconditioning treatments. We seek to define the neural immune signaling responsible for cold-preconditioning as means to identify novel targets for therapeutics development to protect brain before injury onset. Low-level pro-inflammatory mediator signaling changes over time are essential for cold-preconditioning neuroprotection. This signaling is consistent with the basic tenets of physiological conditioning hormesis, which require that irritative stimuli reach a threshold magnitude with sufficient time for adaptation to the stimuli for protection to become evident.
Accordingly, delineation of the immune signaling involved in cold-preconditioning neuroprotection requires that biological systems and experimental manipulations plus technical capacities are highly reproducible and sensitive. Our approach is to use hippocampal slice cultures as an in vitro model that closely reflects their in vivo counterparts with multi-synaptic neural networks influenced by mature and quiescent macroglia / microglia. This glial state is particularly important for microglia since they are the principal source of cytokines, which are operative in the femtomolar range. Also, slice cultures can be maintained in vitro for several weeks, which is sufficient time to evoke activating stimuli and assess adaptive responses. Finally, environmental conditions can be accurately controlled using slice cultures so that cytokine signaling of cold-preconditioning can be measured, mimicked, and modulated to dissect the critical node aspects. Cytokine signaling system analyses require the use of sensitive and reproducible multiplexed techniques. We use quantitative PCR for TNF-&alpha; to screen for microglial activation followed by quantitative real-time qPCR array screening to assess tissue-wide cytokine changes. The latter is a most sensitive and reproducible means to measure multiple cytokine system signaling changes simultaneously. Significant changes are confirmed with targeted qPCR and then protein detection. We probe for tissue-based cytokine protein changes using multiplexed microsphere flow cytometric assays using Luminex technology. Cell-specific cytokine production is determined with double-label immunohistochemistry. Taken together, this brain tissue preparation and style of use, coupled to the suggested investigative strategies, may be an optimal approach for identifying potential targets for the development of novel therapeutics that could mimic the advantages of cold-preconditioning.

We seek to define the neural immune signaling responsible for cold-preconditioning as means to identify novel targets for therapeutics development to protect brain before injury onset. We present strategies for such work that require biological systems, experimental manipulations plus technical capacities that are highly reproducible and sensitive.

Neurological injury is a frequent cause of morbidity and mortality from general anesthesia and related surgical procedures that could be alleviated by ...

Experimental Models to Study the Neuroprotection of Acidic Postconditioning Against Cerebral Ischemia

Stroke is one of the leading causes of mortality and disability worldwide, with limited therapeutic approaches. As an endogenous strategy for neuroprotection, postconditioning treatments have proven to be promising therapies against cerebral ischemia. However, complicated procedures and potential safety issues limit their clinical application. To overcome these disadvantages, we have developed acidic postconditioning (APC) as a therapy for experimental focal cerebral ischemia. APC refers to the mild acidosis treatment by inhaling CO2 during reperfusion following ischemia. Here we present two models to execute APC in vitro and in vivo, respectively. The oxygen-glucose deprivation (OGD) treatment of mice and the corticostriatal occlusion and middle cerebral artery occlusion (MCAO) of mice were employed to mimic cerebral ischemia. APC can be simply achieved by transferring brain slices to acidic buffer bubbled with 20% CO2, or by mice inhaling 20% CO2. APC showed significant protective effects against cerebral ischemia, as reflected by tissue viability and brain infarct volume.

Acidic postconditioning protects against cerebral ischemia. Here we present two models to execute APC. They are achieved respectively by transferring corticostriatal slices to acidic buffer after oxygen-glucose deprivation in vitro and by inhaling 20% CO2 after middle cerebral artery occlusion in vivo.

Stroke is one of the leading causes of mortality and disability worldwide, with limited therapeutic approaches. As an endogenous strategy for ...

Recording Synaptic Plasticity in Acute Hippocampal Slices Maintained in a Small-volume Recycling-, Perfusion-, and Submersion-type Chamber System

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful analysis of synaptic transmission modulation when performing field potential or intracellular recordings. This manuscript describes methodological aspects that might be helpful in improving experimental conditions for the maintenance of acute brain slices and for recording field excitatory postsynaptic potentials in a commercially available submersion chamber with an outflow-carbogenation unit. The outflow-carbogenation helps to stabilize the oxygen level in experiments that rely on the recycling of a small buffer reservoir to enhance the cost-efficiency of drug experiments. In addition, the manuscript presents representative experiments that examine the effects of different carbogenation modes and stimulation paradigms on the activity-dependent synaptic plasticity of synaptic transmission.

This protocol describes the stabilization of the oxygen level in a small volume of recycled buffer and methodological aspects of recording activity-dependent synaptic plasticity in submerged acute hippocampal slices.

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful ...

Two-photon Imaging of Microglial Processes' Attraction Toward ATP or Serotonin in Acute Brain Slices

Microglial cells are resident innate immune cells of the brain that constantly scan their environment with their long processes and, upon disruption of homeostasis, undergo rapid morphological changes. For example, a laser lesion induces in a few minutes an oriented growth of microglial processes, also called &#34;directional motility&#34;, toward the site of injury. A similar effect can be obtained by delivering locally ATP or serotonin (5-hydroxytryptamine [5-HT]). In this article, we describe a protocol to induce a directional growth of microglial processes toward a local application of ATP or 5-HT in acute brain slices of young and adult mice and to image this attraction over time by multiphoton microscopy. A simple method of quantification with free and open-source image analysis software is proposed. A challenge that still characterizes acute brain slices is the limited time, decreasing with age, during which the cells remain in a physiological state. This protocol, thus, highlights some technical improvements (medium, air-liquid interface chamber, imaging chamber with a double perfusion) aimed at optimizing the viability of microglial cells over several hours, especially in slices from adult mice.

Microglia, the resident immune cells of the brain, respond quickly with morphological changes to modifications of their environment. This protocol describes how to use two-photon microscopy to study the attraction of microglial processes toward serotonin or ATP in acute brain slices of mice.

Microglial cells are resident innate immune cells of the brain that constantly scan their environment with their long processes and, upon disruption of ...

Targeted Neuronal Injury for the Non-Invasive Disconnection of Brain Circuitry

Surgical intervention can be quite effective for treating certain types of medically intractable neurological diseases. This approach is particularly useful for disorders in which identifiable neuronal circuitry plays a key role, such as epilepsy and movement disorders. Currently available surgical modalities, while effective, generally involve an invasive surgical procedure, which can result in surgical injury to non-target tissues. Consequently, it would be of value to expand the range of surgical approaches to include a technique that is both non-invasive and neurotoxic.
Here, a method is presented for producing focal, neuronal lesions in the brain in a non-invasive manner. This approach utilizes low-intensity focused ultrasound together with intravenous microbubbles to transiently and focally open the Blood Brain Barrier (BBB). The period of transient BBB opening is then exploited to focally deliver a systemically administered neurotoxin to a targeted brain area. The neurotoxin quinolinic acid (QA) is normally BBB-impermeable, and is well-tolerated when administered intraperitoneally or intravenously. However, when QA gains direct access to brain tissue, it is toxic to the neurons. This method has been used in rats and mice to target specific brain regions. Immediately after MRgFUS, successful opening of the BBB is confirmed using contrast enhanced T1-weighted imaging. After the procedure, T2 imaging shows injury restricted to the targeted area of the brain and the loss of neurons in the targeted area can be confirmed post-mortem utilizing histological techniques. Notably, animals injected with saline rather than QA do demonstrate opening of the BBB, but dot not exhibit injury or neuronal loss. This method, termed Precise Intracerebral Non-invasive Guided surgery (PING) could provide a non-invasive approach for treating neurological disorders associated with disturbances in neural circuitry.

The goal of the protocol is to provide a method for producing non-invasive neuronal lesions in the brain. The method utilizes Magnetic Resonance-guided Focused Ultrasound (MRgFUS) to open the Blood Brain Barrier in a transient and focal manner, in order to deliver a circulating neurotoxin to the brain parenchyma.

Surgical intervention can be quite effective for treating certain types of medically intractable neurological diseases. This approach is particularly ...

Minimizing Hypoxia in Hippocampal Slices from Adult and Aging Mice

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&#8217;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.

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.

Acute hippocampal slices have enabled generations of neuroscientists to explore synaptic, neuronal, and circuit properties in detail and with high ...

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 ...

Assessing Changes in Synaptic Plasticity Using an Awake Closed-Head Injury Model of Mild Traumatic Brain Injury

Mild traumatic brain injuries (mTBIs) are a prevalent health issue in North America. There is increasing pressure to utilize ecologically valid models of closed-head mTBI in the preclinical setting to increase translatability to the clinical population. The awake closed-headed injury (ACHI) model uses a modified controlled cortical impactor to deliver closed-headed injury, inducing clinically relevant behavioral deficits without the need for a craniotomy or the use of an anesthetic.
This technique does not normally induce fatalities, skull fractures, or brain bleeds, and is more consistent with being a mild injury. Indeed, the mild nature of the ACHI procedure makes it ideal for studies investigating repetitive mTBI (r-mTBI). Growing evidence indicates that r-mTBI can result in a cumulative injury that produces behavioral symptoms, neuropathological changes, and neurodegeneration. r-mTBI is common in youths playing sports, and these injuries occur during a period of robust synaptic reorganization and myelination, making the younger population particularly vulnerable to the long-term influences of r-mTBI.
Further, r-mTBI occurs in cases of intimate partner violence, a condition for which there are few objective screening measures. In these experiments, synaptic function was assessed in the hippocampus in juvenile rats that had experienced r-mTBI using the ACHI model. Following the injuries, a tissue slicer was utilized to make hippocampal slices to evaluate bidirectional synaptic plasticity in the hippocampus at either 1 or 7 days following the r-mTBI. Overall, the ACHI model provides researchers with an ecologically valid model to study changes in synaptic plasticity following mTBI and r-mTBI.

Here, it is demonstrated how an awake closed-head injury model can be used for examining the effects of repeated mild traumatic brain injury (r-mTBI) on synaptic plasticity in the hippocampus. The model replicates important features of r-mTBI in patients and is used in conjunction with in vitro electrophysiology.

Mild traumatic brain injuries (mTBIs) are a prevalent health issue in North America. There is increasing pressure to utilize ecologically valid models of ...

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

Summary

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Chapters in this video

Strategies for Study of Neuroprotection from Cold-preconditioning

Experimental Models to Study the Neuroprotection of Acidic Postconditioning Against Cerebral Ischemia

Recording Synaptic Plasticity in Acute Hippocampal Slices Maintained in a Small-volume Recycling-, Perfusion-, and Submersion-type Chamber System

Two-photon Imaging of Microglial Processes' Attraction Toward ATP or Serotonin in Acute Brain Slices

Targeted Neuronal Injury for the Non-Invasive Disconnection of Brain Circuitry

Minimizing Hypoxia in Hippocampal Slices from Adult and Aging Mice

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

Horizontal Hippocampal Slices of the Mouse Brain

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

Assessing Changes in Synaptic Plasticity Using an Awake Closed-Head Injury Model of Mild Traumatic Brain Injury