I thank Dr. Carla J. Shatz for advice and support, and Dr. Barbara K. Brott and Michelle K. Drews for critically reading the manuscript. The work is supported by NIH EY02858 and the Mathers Charitable Foundation grants to CJS.

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
<ol>
	<li>Prepare 1 L of 1x aCSF&#160;(in mM): 125 NaCl, 26 NaHCO3, 2.3 KCl, 1.26 KH2PO4, 1.3 Mg2SO4&#183;7H2O, 2.5 CaCl2, 25 glucose (Table 1). Use this solution for the recovery chamber and subsequent recordings. 
	NOTE:&#160;Store aCSF as a 10x stock solution that contains NaCl, NaHCO3, KCl, and KH2PO4. Store Mg2SO4&#183;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 M&#937; water. 
	NOTE: Depending on the brain region or mouse strain, chilled aCSF can be also used for transcardial perfusion with success11,15,26.</li>
	<li>Prepare 300 mL of NMDG-aCSF&#160;(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:&#160;Store NMDG-aCSF as a 3x stock solution at 4 &#176;C. Prepare working solution on the day of the experiment; add choline bicarbonate and glucose before adjusting the final volume with 18 M&#937; 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-aCSF13,20.</li>
	<li>Set the vibrating microtome cutting tray and mounting disk to -20 &#176;C.</li>
	<li>Prepare the recovery chamber.
	<ol>
		<li>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&#160;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.</li>
	</ol>
	</li>
	<li>Prepare the transcardial perfusion and cutting solution.
	<ol>
		<li>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!</li>
		<li>Place the bottle with chilled NMDG-aCSF on ice and bubble. The solution should be between 0&#8210;2 &#176;C. 
		NOTE: Slushy NMDG-&#160;aCSF implies having most of the solution as liquid, while a small fraction of slushy ice present will keep the solution near 0 &#176;C throughout perfusion and cutting. Care should be taken to keep away ice crystals from the brain during cutting.</li>
	</ol>
	</li>
	<li>Prepare the tissue-mounting disk.
	<ol>
		<li>Take the disk out of the freezer, wipe it dry if needed.</li>
		<li>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.
		<ol>
			<li>For 5% agar plate, dissolve 5 g of agar in 100 mL of 18 M&#937; water by microwaving, and pour into a clean petri dish. Keep at 4 &#176;C.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Take the cutting tray out of the freezer, place it into the microtome, surround it with ice, and load the blade.</li>
</ol>
2. Transcardial perfusion and brain extraction
<ol>
	<li>Overdose mice via an intraperitoneal injection of anesthetic&#160;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 M&#937; 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.</li>
	<li>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.</li>
	<li>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&#160;is solution coming out of the nose of the animal.</li>
	<li>Perfusion
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Insert the 27 G needle into the left ventricle; the mouse&#8217;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.</li>
		<li>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.</li>
		<li>Run the pump for few more minutes. If using rectal thermometer, the body temperature should fall down to 28&#8210;29 &#176;C, and animal&#8217;s nose should be cold to the touch.</li>
	</ol>
	</li>
	<li>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&#176;, a spatula, a &#8220;60&#176;&#8221; tool (Figure 1A,B), and a small soft brush.
	<ol>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>
3. Slicing
<ol>
	<li>Take the brain out of the NMDG-aCSF and place it on a piece of filter paper.</li>
	<li>Cut and remove a 60&#176; wedge from the rostral end of the forebrain using a &#8220;60&#176;&#8221; tool centered at the midline. Cut surfaces will be used for mounting to achieve the proper angle for transversal hippocampal slices as discussed below&#160;(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&#176; cut (Figure 1A).</li>
	<li>Separate hemispheres down the midline with the scalpel.</li>
	<li>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.</li>
	<li>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&#160;(Figure 1B).</li>
	<li>Submerge the disk with hemispheres into a cutting chamber containing ice-cold carbogenated NMDG-aCSF.</li>
	<li>Cut 400 &#181;m sections. Cutting should be done in less than 10 min. Different microtomes will require different settings to achieve this time. A total of 8&#8210;10 slices from the dorsal hippocampus region will be obtained.</li>
</ol>
4. Recovery
<ol>
	<li>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.</li>
	<li>Incubate at room temperature (22&#8210;24 &#176;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 &#176;C for 30 min followed by room temperature incubation. Rewarming at 37 &#176;C can introduce cytotoxic edema13.</li>
</ol>

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The author has nothing to disclose.

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&#160;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. ...

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 time2,3,4. 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 activity5,6. 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-decapitation4,7,8. 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 &#176;C to 31 &#176;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 edema10,11. 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 regions13,14,15,16.
Numerous brain-slicing protocols involve using cold temperatures only during the slice-cutting step, sometimes in combination with Na+ ion replacement strategy16,17. 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&#8217;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 slices11,20, perirhinal cortex21, rat hippocampus10,22,23, 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 hypoxia13,20. 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.

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			<td>made in-house</td>
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			<td>#10 scalpel blade</td>
			<td>Bard-Parker (Aspen Surgical)</td>
			<td>371110</td>
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		</tr>
		<tr>
			<td>1M CaCl2</td>
			<td>Fluka Analytical</td>
			<td>21114</td>
			<td></td>
		</tr>
		<tr>
			<td>95%O2/5%CO2</td>
			<td>Praxiar or another local supplier</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Acepromazine maleate (AceproJect)</td>
			<td>Henry Schein</td>
			<td>5700850</td>
			<td></td>
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			<td>Agar</td>
			<td>Fisher</td>
			<td>BP1423-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Beakers, measuring cylinders, reagent bottles</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Brushes size 00-2</td>
			<td>Ted Pella</td>
			<td></td>
			<td>Crafts stores are another source of soft brushes, with larger selection and better quality than Ted Pella.</td>
		</tr>
		<tr>
			<td>CCD camera</td>
			<td>Olympus</td>
			<td>XM10</td>
			<td></td>
		</tr>
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			<td>Pfalz &#38; Bauer</td>
			<td>C21240</td>
			<td></td>
		</tr>
		<tr>
			<td>Cyanoacrilate glue</td>
			<td>Krazy glue</td>
			<td></td>
			<td>Singles</td>
		</tr>
		<tr>
			<td>Decapitation scissors</td>
			<td>FST</td>
			<td>14130-17</td>
			<td></td>
		</tr>
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			<td>Feather blades</td>
			<td>Feather</td>
			<td>FA-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Filter paper #2</td>
			<td>Whatman</td>
			<td></td>
			<td>Either rounds or pieces cut from a bigger sheet work well.</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>A. Dumont &#38; Fils</td>
			<td>Inox 3c</td>
			<td></td>
		</tr>
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			<td>Glass bubblers (Robu glass borosillicate microfilter candles) - porosity 3</td>
			<td>Robuglas.com</td>
			<td>18103 or 18113</td>
			<td>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.</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma-Aldrich</td>
			<td>G8270</td>
			<td></td>
		</tr>
		<tr>
			<td>HCl</td>
			<td>Fisher</td>
			<td>A144SI-212</td>
			<td></td>
		</tr>
		<tr>
			<td>Ice buckets</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma-Aldrich</td>
			<td>P4504</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine HCl (KetaVed)</td>
			<td>VEDCO</td>
			<td>NDC 50989-996-06</td>
			<td></td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>Sigma-Aldrich</td>
			<td>P0662</td>
			<td></td>
		</tr>
		<tr>
			<td>Leica Tissue slicer VT1000S</td>
			<td></td>
			<td></td>
			<td>The cutting settings are 1 mm horizontal blade amplitude, frequency dial at 9, and speed setting at 2</td>
		</tr>
		<tr>
			<td>Magnetic stirrers and stir bars</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Mg2SO4 x 7H2O</td>
			<td>Sigma-Aldrich</td>
			<td>230391</td>
			<td></td>
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			<td>MgCl2</td>
			<td>Sigma-Aldrich</td>
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			<td>MilliQ water machine</td>
			<td>Millipore</td>
			<td></td>
			<td>Source for 18 Mohm water</td>
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			<td>Na-ascorbate</td>
			<td>Sigma-Aldrich</td>
			<td>A4035</td>
			<td></td>
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		<tr>
			<td>Na-pyruvate</td>
			<td>Sigma-Aldrich</td>
			<td>P8574</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma-Aldrich</td>
			<td>S3014</td>
			<td></td>
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			<td>NaHCO3</td>
			<td>EMD</td>
			<td>SX0320-1</td>
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			<td>Needle 27G1/2</td>
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			<td>NMDG</td>
			<td>Sigma-Aldrich</td>
			<td>M2004</td>
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			<td>Paper tape</td>
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			<td>Peristaltic pump</td>
			<td>Cole-Parmer</td>
			<td>#7553-70</td>
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			<td>Peristaltic pump head</td>
			<td>Cole-Parmer</td>
			<td>Masterflex #7518-00</td>
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			<td>Personna blades</td>
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			<td>Personna double edge</td>
			<td>Amazon</td>
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			<td>pH meter</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Recovery chamber</td>
			<td>in-house made</td>
			<td></td>
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			<td>Scalpel blade handle size 3</td>
			<td>Bard-Parker (Aspen Surgical)</td>
			<td>371030</td>
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			<td>Scissors angled blade</td>
			<td>FST</td>
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			<td>Spatulas</td>
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			<td></td>
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		</tr>
		<tr>
			<td>Transfer pipettes</td>
			<td>Samco Scientific</td>
			<td>225</td>
			<td></td>
		</tr>
		<tr>
			<td>Upright microscope</td>
			<td>Olympus BX51WI</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Xylazine HCl (XylaMed)</td>
			<td>VetOne</td>
			<td>510650</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

We applied the above protocol to generate hippocampus slices from CamKIIa-Cre+;WT&#160;mice on a mixed genetic background C57Bl/ 6 x SV/ 129J, at P &#62; 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 (&#62;90%) is routi...

Watch this Scientific Journal Video about Minimizing Hypoxia in Hippocampal Slices from Adult and Aging Mice at JoVE.com

Minimizing Hypoxia in Hippocampal Slices from Adult and Aging Mice

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 ...

minimizing-hypoxia-in-hippocampal-slices-from-adult-and-aging-mice

Research

JoVE Journal

Neuroscience

7.4K Views.  Stanford University. 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.

Video: Minimizing Hypoxia in Hippocampal Slices from Adult and Aging Mice

Preparation of Oligomeric &beta;-amyloid1-42 and Induction of Synaptic Plasticity Impairment on Hippocampal Slices

Impairment of synaptic connections is likely to underlie the subtle amnesic changes occurring at the early stages of Alzheimer s Disease (AD). &beta;-amyloid (A&beta;), a peptide produced in high amounts in AD, is known to reduce Long-Term Potentiation (LTP), a cellular correlate of learning and memory. Indeed, LTP impairment caused by A&beta; is a useful experimental paradigm for studying synaptic dysfunctions in AD models and for screening drugs capable of mitigating or reverting such synaptic impairments. Studies have shown that A&beta; produces the LTP disruption preferentially via its oligomeric form. Here we provide a detailed protocol for impairing LTP by perfusion of oligomerized synthetic A&beta;1-42 peptide onto acute hippocampal slices. In this video, we outline a step-by-step procedure for the preparation of oligomeric A&beta;1-42. Then, we follow an individual experiment in which LTP is reduced in hippocampal slices exposed to oligomerized A&beta;1-42 compared to slices in a control experiment where no A&beta;1-42 exposure had occurred.

Preparation of Oligomeric &#946;-amyloid1-42 and Induction of Synaptic Plasticity Impairment on Hippocampal Slices

One feature of Alzheimer's Disease is the elevation of A&beta;1-42 peptide. Here we provide a protocol for preparing synthetic A&beta;1-42 oligomers, which impairs hippocampal Long-Term Potentiation, a cellular correlate of memory. This procedure is useful for investigating mechanisms of A&beta;-induced pathology and drug screening.

Impairment of synaptic connections is likely to underlie the subtle amnesic changes occurring at the early stages of Alzheimer s Disease (AD). ...

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

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

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

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

Isolation and Culture of Hippocampal Neurons from Prenatal Mice

Primary cultures of rat and murine hippocampal neurons are widely used to reveal cellular mechanisms in neurobiology. By isolating and growing individual neurons, researchers are able to analyze properties related to cellular trafficking, cellular structure and individual protein localization using a variety of biochemical techniques. Results from such experiments are critical for testing theories addressing the neural basis of memory and learning. However, unambiguous results from these forms of experiments are predicated on the ability to grow neuronal cultures with minimum contamination by other brain cell types. In this protocol, we use specific media designed for neuron growth and careful dissection of embryonic hippocampal tissue to optimize growth of healthy neurons while minimizing contaminating cell types (i.e. astrocytes). Embryonic mouse hippocampal tissue can be more difficult to isolate than similar rodent tissue due to the size of the sample for dissection. We show detailed dissection techniques of hippocampus from embryonic day 19 (E19) mouse pups. Once hippocampal tissue is isolated, gentle dissociation of neuronal cells is achieved with a dilute concentration of trypsin and mechanical disruption designed to separate cells from connective tissue while providing minimum damage to individual cells. A detailed description of how to prepare pipettes to be used in the disruption is included. Optimal plating densities are provided for immuno-fluorescence protocols to maximize successful cell culture. The protocol provides a fast (approximately 2 hr) and efficient technique for the culture of neuronal cells from mouse hippocampal tissue.

We provide a protocol for the culture of highly purified hippocampal neurons from prenatal mouse brains without the use of a feeder glial cell layer.

Primary cultures of rat and murine hippocampal neurons are widely used to reveal cellular mechanisms in neurobiology. By isolating and growing individual ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

Paired Whole Cell Recordings in Organotypic Hippocampal Slices

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct electrophysiological characterization of the synaptic connections between individual neurons, but also pharmacological manipulation of either the presynaptic or the postsynaptic neuron. When carried out in organotypic hippocampal slice cultures, the probability that two neurons are synaptically connected is significantly increased. This preparation readily enables identification of cell types, and the neurons maintain their morphology and properties of synaptic function similar to that in native brain tissue. A major advantage of paired whole cell recordings is the highly precise information it can provide on the properties of synaptic transmission and plasticity that are not possible with other more crude techniques utilizing extracellular axonal stimulation. Paired whole cell recordings are often perceived as too challenging to perform. While there are challenging aspects to this technique, paired recordings can be performed by anyone trained in whole cell patch clamping provided specific hardware and methodological criteria are followed. The probability of attaining synaptically connected paired recordings significantly increases with healthy organotypic slices and stable micromanipulation allowing independent attainment of pre- and postsynaptic whole cell recordings. While CA3-CA3 pyramidal cell pairs are most widely used in the organotypic slice hippocampal preparation, this technique has also been successful in CA3-CA1 pairs and can be adapted to any neurons that are synaptically connected in the same slice preparation. In this manuscript we provide the detailed methodology and requirements for establishing this technique in any laboratory equipped for electrophysiology.

Pair recordings are simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling precise electrophysiological and pharmacological characterization of the synapses between individual neurons. Here we describe the detailed methodology and requirements for establishing this technique in organotypic hippocampal slice cultures in any laboratory equipped for electrophysiology.

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct ...

Investigation of Synaptic Tagging/Capture and Cross-capture using Acute Hippocampal Slices from Rodents

Synaptic tagging and capture (STC) and cross-tagging are two important mechanisms at cellular level that explain how synapse-specificity and associativity is achieved in neurons within a specific time frame. These long-term plasticity-related processes are the leading candidate models to study the basis of memory formation and persistence at the cellular level. Both STC and cross-tagging involve two serial processes: (1) setting of the synaptic tag as triggered by a specific pattern of stimulation, and (2) synaptic capture, whereby the synaptic tag interacts with newly synthesized plasticity-related proteins (PRPs). Much of the understanding about the concepts of STC and cross-tagging arises from the studies done in CA1 region of the hippocampus and because of the technical complexity many of the laboratories are still unable to study these processes. Experimental conditions for the preparation of hippocampal slices and the recording of stable late-LTP/LTD are extremely important to study synaptic tagging/cross-tagging. This video article describes the experimental procedures to study long-term plasticity processes such as STC and cross-tagging in the CA1 pyramidal neurons using stable, long-term field-potential recordings from acute hippocampal slices of rats.

This video article describes experimental procedures to study long-term plasticity and its associative processes such as synaptic tagging, capture and cross-tagging in the CA1 pyramidal neurons using acute hippocampal slices from rodents.

Synaptic tagging and capture (STC) and cross-tagging are two important mechanisms at cellular level that explain how synapse-specificity and associativity ...

Evaluation of Synapse Density in Hippocampal Rodent Brain Slices

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

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

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

Preparing Fresh Retinal Slices from Adult Zebrafish for Ex Vivo Imaging Experiments

The retina is a complex tissue that initiates and integrates the first steps of vision. Dysfunction of retinal cells is a hallmark of many blinding diseases, and future therapies hinge on fundamental understandings about how different retinal cells function normally. Gaining such information with biochemical methods has proven difficult because contributions of particular cell types are diminished in the retinal cell milieu. Live retinal imaging can provide a view of numerous biological processes on a subcellular level, thanks to a growing number of genetically encoded fluorescent biosensors. However, this technique has thus far been limited to tadpoles and zebrafish larvae, the outermost retinal layers of isolated retinas, or lower resolution imaging of retinas in live animals. Here we present a method for generating live ex vivo retinal slices from adult zebrafish for live imaging via confocal microscopy. This preparation yields transverse slices with all retinal layers and most cell types visible for performing confocal imaging experiments using perfusion. Transgenic zebrafish expressing fluorescent proteins or biosensors in specific retinal cell types or organelles are used to extract single-cell information from an intact retina. Additionally, retinal slices can be loaded with fluorescent indicator dyes, adding to the method&#39;s versatility. This protocol was developed for imaging Ca2+ within zebrafish cone photoreceptors, but with proper markers it could be adapted to measure Ca2+ or metabolites in M&#252;ller cells, bipolar and horizontal cells, microglia, amacrine cells, or retinal ganglion cells. The retinal pigment epithelium is removed from slices so this method is not suitable for studying that cell type. With practice, it is possible to generate serial slices from one animal for multiple experiments. This adaptable technique provides a powerful tool for answering many questions about retinal cell biology, Ca2+, and energy homeostasis.

Preparing Fresh Retinal Slices from Adult Zebrafish for Ex Vivo Imaging Experiments

Imaging retinal tissue can provide single-cell information that cannot be gathered from traditional biochemical methods. This protocol describes preparation of retinal slices from zebrafish for confocal imaging. Fluorescent genetically encoded sensors or indicator dyes allow visualization of numerous biological processes in distinct retinal cell types.

The retina is a complex tissue that initiates and integrates the first steps of vision. Dysfunction of retinal cells is a hallmark of many blinding ...

Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools to selectively manipulate brain rhythms. Here we describe an approach combining projection-specific optogenetics with extracellular electrophysiology for high-fidelity control of hippocampal theta oscillations (5-10 Hz) in behaving mice. The specificity of the optogenetic entrainment is achieved by targeting channelrhodopsin-2 (ChR2) to the GABAergic population of medial septal cells, crucially involved in the generation of hippocampal theta oscillations, and a local synchronized activation of a subset of inhibitory septal afferents in the hippocampus. The efficacy of the optogenetic rhythm control is verified by a simultaneous monitoring of the local field potential (LFP) across lamina of the CA1 area and/or of neuronal discharge. Using this readily implementable preparation we show efficacy of various optogenetic stimulation protocols for induction of theta oscillations and for the manipulation of their frequency and regularity. Finally, a combination of the theta rhythm control with projection-specific inhibition addresses the readout of particular aspects of the hippocampal synchronization by efferent regions.

We describe the use of optogenetics and electrophysiological recordings for selective manipulations of hippocampal theta oscillations (5-10 Hz) in behaving mice. The efficacy of the rhythm entrainment is monitored using local field potential. A combination of opto- and pharmacogenetic inhibition addresses the efferent readout of hippocampal synchronization.

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools ...

Single Synapse Indicators of Glutamate Release and Uptake in Acute Brain Slices from Normal and Huntington Mice

Synapses are highly compartmentalized functional units that operate independently on each other. In Huntington&#39;s disease (HD) and other neurodegenerative disorders, this independence might be compromised due to insufficient glutamate clearance and the resulting spill-in and spill-out effects. Altered astrocytic coverage of the presynaptic terminals and/or dendritic spines as well as a reduced size of glutamate transporter clusters at glutamate release sites have been implicated in the pathogenesis of diseases resulting in symptoms of dys-/hyperkinesia. However, the mechanisms leading to the dysfunction of glutamatergic synapses in HD are not well understood. Improving and applying synapse imaging we have obtained data shedding new light on the mechanisms impeding the initiation of movements. Here, we describe the principle elements of a relatively inexpensive approach to achieve single synapse resolution by using the new genetically encoded ultrafast glutamate sensor iGluu, wide-field optics, a scientific CMOS (sCMOS) camera, a 473 nm laser and a laser positioning system to evaluate the state of corticostriatal synapses in acute slices from age appropriate healthy or diseased mice. Glutamate transients were constructed from single or multiple pixels to obtain estimates of i) glutamate release based on the maximal elevation of the glutamate concentration [Glu] next to the active zone and ii) glutamate uptake as reflected in the time constant of decay (TauD) of the perisynaptic [Glu]. Differences in the resting bouton size and contrasting patterns of short-term plasticity served as criteria for the identification of corticostriatal terminals as belonging to the intratelencephalic (IT) or the pyramidal tract (PT) pathway. Using these methods, we discovered that in symptomatic HD mice ~40% of PT-type corticostriatal synapses exhibited insufficient glutamate clearance, suggesting that these synapses might be at risk to excitotoxic damage. The results underline the usefulness of TauD as a biomarker of dysfunctional synapses in Huntington mice with a hypokinetic phenotype.

We present a protocol to evaluate the balance between glutamate release and clearance at single corticostriatal glutamatergic synapses in acute slices from adult mice. This protocol uses the fluorescent sensor iGluu for glutamate detection, a sCMOS camera for signal acquisition and a device for focal laser illumination.

Minimizing Hypoxia in Hippocampal Slices from Adult and Aging Mice

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Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice

Single Synapse Indicators of Glutamate Release and Uptake in Acute Brain Slices from Normal and Huntington Mice