We thank Aimee Rogers for providing hands-on training and continued product support. We thank Dr. Amanda Burke for ongoing troubleshooting and clarifying discussions. We thank Meredith Joheim and the UAMS Science Communication Group for the grammatical editing and formatting of this manuscript.&#160;This study was supported by NIH R25GM083247 and NIH 1R01CA258673 (A.R.A).

Experiments were conducted in accordance with the ethical standards approved by the Institutional Animal Care and Use Committee at UAMS. 6-month-old female C57Bl6/J wild-type mice were purchased and group-housed (4 mice per cage) under a constant 12 h light/dark cycle.
1. Preparation of reagents
<ol>
	<li>Prepare fixable live/dead stain stock solution. Reconstitute the fluorescent stain with 20 &#181;L of dimethyl sulfoxide (DMSO).</li>
	<li>Wrap the vial in foil, label it as &#34;Reconstituted&#34;, and store it at -20 &#176;C for up to six months.</li>
	<li>Prepare a 0.9% saline solution with heparin. Dilute the contents of one vial of heparin sodium (10,000 USP units per 10 mL) in 1 L of double-distilled water (ddH2O).</li>
	<li>Prepare enough for approximately 45 mL per animal and store at 4 &#176;C for up to one week.</li>
	<li>Make 1% paraformaldehyde (PFA).
	<ol>
		<li>In a fume hood, heat a hot plate to 50 &#176;C. In a Microwave, heat 100 mL of ddH2O in a glass beaker to approximately 60 &#176;C. Add a magnetic stir bar and transfer to the hot plate.</li>
		<li>In the fume hood, weigh out 1 g of PFA and add to the beaker of ddH2O. Add 0.1125 g of NaOH crystals and mix until dissolved (5-10 min).</li>
		<li>Add 0.4 g of NaPO4- monobasic and mix until dissolved (2-5 min). Vacuum filter the solution and adjust pH to 7.4 with HCl and NaOH.</li>
		<li>Cool on ice or at 4&#176;C for 30 min before storing. 
		NOTE: Aliquots of 1.5 mL can be stored at -20 &#176;C for one year. Avoid freeze-thaw cycles. If, after thawing, the solution becomes cloudy or a precipitate has formed, the solution should not be used. 
		CAUTION: Toxic, flammable. Always work with PFA under a ventilated hood wearing proper personal protective equipment.</li>
	</ol>
	</li>
	<li>Resuspend lyophilized Enzyme A with 1 mL of Buffer A. Do not vortex the solution. 
	NOTE: Enzyme A and Buffer A as well as Buffers A, Y, and Z are reagents in the commercial Adult Brain Dissociation Kit17.</li>
	<li>Divide Enzyme P into aliquots of 50 &#181;L and resuspend Enzyme A into 10 &#181;L aliquots. Per kit instructions, store at -20 &#176;C for up to six months. Avoid freeze-thaw cycles.</li>
</ol>
2. Day of experiment
<ol>
	<li>Cool the tabletop centrifuge to 4 &#176;C.</li>
	<li>Place aliquot(s) of PFA in the fridge for gradual thawing.</li>
	<li>Place the reconstituted live/dead stain in the dark (e.g., a drawer) to thaw at room temperature.</li>
	<li>Prepare the bovine serum albumin (BSA) Buffer. Add 0.5 g of BSA to 100 mL of 1x Dulbecco&#39;s phosphate-buffered solution without calcium and magnesium (D-PBS), pH 7.2.</li>
	<li>Add a stir bar and mix on a stir plate for 30 min. Transfer to 50 mL conical tubes and store at 4 &#176;C. 
	NOTE: Always use freshly prepared BSA buffer.</li>
	<li>Prepare live/dead stain working dilution. Add 1 &#181;L of the reconstituted live/dead stain stock solution to 360 &#181;L D-PBS and store it in the dark (e.g., a drawer or box) at room temperature. Prepare 50 &#181;L of the working dilution per sample.</li>
</ol>
3. Perfusion
<ol>
	<li>Place the saline solution with heparin on ice.</li>
	<li>Turn on oxygen, set the flowmeter indicator ball on the small animal anesthesia vaporizer system to 1 L/min. Ensure there is adequate oxygen pressure and isoflurane.</li>
	<li>Adjust the vaporizer dial to 3.5% (for induction and maintenance).</li>
	<li>Prime the perfusion pump lines with the saline/heparin solution. Set the speed to 6 mL/min.</li>
	<li>Place the mouse in the induction chamber, turn on the breather, and wait several minutes until the mouse is unresponsive. Confirm sufficient depth of anesthesia through the absence of pedal withdrawal to noxious pinch.</li>
	<li>Place the mouse on its back on the dissection tray with its nose in the nose cone. Perform a secondary confirmation of full anesthetization through the absence of pedal withdrawal to noxious pinch. Pin all four paws to the tray.</li>
	<li>Spray the animal&#39;s abdomen with 20% ethanol.</li>
	<li>Using forceps, pinch the lower abdomen and lift the skin. Use scissors to cut through fur and skin to the bottom of the ribcage.</li>
	<li>Make two diagonal incisions from below the ribcage toward each shoulder.</li>
	<li>Carefully resect the diaphragm (avoiding the lungs and heart). Resect the ribcage to expose the heart.</li>
	<li>Carefully sever any connective tissue around the heart. 
	NOTE: Steps 3.10-3.11 are critical; perform with proficiency and dexterity.</li>
	<li>Use the scissors to clip the right atrium (dark lobe on the upper left of the heart). Turn off the flow of isoflurane to the breather.</li>
	<li>Hold the heart steady with forceps. With the bevel of the butterfly needle facing up, pierce the left ventricle while keeping the needle level and parallel to the animal.</li>
	<li>Hold the needle in place, turn on the pump, and perfuse at least 30 mL of the saline/heparin solution until the fluid leaving the heart is opaque and the liver and lungs pale in color. 
	NOTE: Steps 3.13-3.14 are critical; perform with proficiency and dexterity.</li>
	<li>Turn off the pump, remove the needle, and transfer the mouse to the dissection area.</li>
</ol>
4. Dissection
<ol>
	<li>Using large surgical scissors, decapitate the head.</li>
	<li>Cut the fur from the back of the head up to the eyes. Peel the skin back to expose the skull.</li>
	<li>Clip the skull between the eyes. Make two cuts at the back of the skull, at the 10 and 2 o&#39;clock positions, then make one long cut (keep tips up to avoid damaging the brain) along the midsagittal line of the skull to the original cut between the eyes.</li>
	<li>Use forceps to peel the two halves of the skull away to the sides. Use a spatula to remove the brain and place it into a 60 mm glass Petri dish on ice filled with cold D-PBS (Figure 1).</li>
	<li>Use a scalpel or razor to separate each hemisphere. Then remove the olfactory bulbs and cerebellum.</li>
	<li>Use forceps to remove the midbrain until the hippocampus is exposed.</li>
	<li>Secure the brain with forceps. Using a second set of forceps, gently tease the hippocampus out of each hemisphere, and transfer both hippocampi to a labeled 1.5 mL tube containing cold D-PBS.</li>
	<li>Place the sample tube containing the two hippocampi from the mouse on ice.</li>
</ol>
5. Prepare Enzyme Mix 1 and 2 for each sample
NOTE: For volumes greater than 2 mL, use a 10 mL serologic pipette; for volumes, 200 &#181;L-2 mL, use a 1000 &#181;L pipette; for volumes, 21-199 &#181;L, use a 200 &#181;L pipette; for volumes, 2-20 &#181;L, use a 20 &#181;L pipette; for volumes under 2 &#181;L, use a 0-2 &#181;L pipette.
<ol>
	<li>For each sample, thaw one aliquot each of Enzyme P and Enzyme A at room temperature.</li>
	<li>For Enzyme mix 1, combine 50 &#181;L of Enzyme P and 1900 &#181;L of Buffer Z in a labeled C Tube (Table of Materials).</li>
	<li>For Enzyme mix 2, add 20 &#181;L of Buffer Y to the thawed 10 &#181;L aliquot of Enzyme A.</li>
</ol>
6. Adult brain dissociation protocol17
NOTE: When working with samples, tubes should be placed in a tube rack at room temperature while BSA and D-PBS remain on ice unless otherwise noted.
<ol>
	<li>Switch on the dissociator.</li>
	<li>Use forceps to transfer the hippocampi tissue pieces to the C Tube.</li>
	<li>Transfer 30 &#181;L of Enzyme mix 2 into the C Tube. Twist the cap until tension is felt, then tighten until it clicks.</li>
	<li>Place the C Tube upside down into a position of the dissociator; the sample will be assigned the Selected status (Figure 2). Secure the heater over the C Tube.</li>
	<li>Press the folder icon, select Favorites folder, scroll to and select the 37C_ABDK_02 program. Click on OK to apply the program to all selected C tubes, then tap on Start (Figure 2).</li>
	<li>Label one 50 mL conical tube per sample.</li>
	<li>Place a 70 &#181;m cell strainer on each 50 mL conical tube and wet with 2 mL of BSA buffer.</li>
	<li>Upon completion of the program, remove the heater and the C tube from the dissociator.</li>
	<li>Add 4 mL of BSA buffer to the sample and apply the mixture to the cell strainer on the 50 mL conical tube.</li>
	<li>Add 10 mL of D-PBS to the C Tube, close it, and swirl the solution gently. Apply it to the cell strainer on the 50 mL conical tube.</li>
	<li>Discard the cell strainer and the C Tube. Centrifuge the suspension at 300 x g for 10 min at 4 &#176;C. Then, aspirate and discard the supernatant.</li>
</ol>
7. Debris removal
<ol>
	<li>Resuspend the pellet with 1550 &#181;L of cold D-PBS and transfer the suspension to a labeled 15 mL conical tube.</li>
	<li>Add 450 &#181;L of cold Debris Removal Solution&#160;and pipette up and down (do not vortex). 
	NOTE: Debris Removal Solution is a reagent in the commercial Adult Brian Dissociation Kit17.</li>
	<li>Gently overlay 1 mL of cold D-PBS on top of the cell suspension, keeping the tip against the wall of the conical tube. Repeat until the total overlay is 2 mL. 
	NOTE: This step is critical; perform with proficiency and dexterity.</li>
	<li>Centrifuge at 3000 x g for 10 min at 4 &#176;C with full acceleration and full brake. 
	NOTE: If the phases are not clearly separated, repeat steps 7.2-7.3. Centrifuge a final time at 1000 x g for 10 min at 4 &#176;C.</li>
	<li>The suspension should now consist of three distinct layers (Figure 3). Aspirate the topmost layer. Sweep the pipette tip back and forth to aspirate the white middle layer. Remove as much of the middle layer as possible without disturbing the bottommost layer. 
	NOTE: This step is critical; perform with proficiency and dexterity.</li>
	<li>Add 2 mL of cold D-PBS and pipette up and down to mix.</li>
	<li>Centrifuge at 1000 x g for 10 min at 4 &#176;C with full acceleration and full brake. Aspirate and discard the supernatant. Resuspend the pellet in 1 mL of BSA buffer. 
	&#8203;NOTE: Cells can be resuspended in the appropriate buffer then magnetically labeled and isolated in preparation for single-cell sequencing at this point.</li>
</ol>
8. Cell count
<ol>
	<li>Perform cell counting as per the manufacturer&#39;s protocol of available cell counter (one option is noted in the Table of Materials)</li>
</ol>
9. Live/dead stain
<ol>
	<li>Centrifuge the remaining 900 &#181;L (from 7.7) at 1000 x g for 10 min at 4 &#176;C with full acceleration and full brake.</li>
	<li>While the sample is spinning, label one flow tube per sample and wrap it in foil to limit light exposure.</li>
	<li>Aspirate and discard the supernatant.</li>
	<li>Resuspend the pellet in 50 &#181;L of diluted live/dead stain (previously prepared). 
	NOTE: This step should be performed in a low-light setting. Turn off overhead room lights to achieve this.</li>
	<li>Transfer each sample to the corresponding labeled flow tube and incubate at room temperature for 8-10 min in the dark (e.g., a drawer or box).</li>
	<li>Add 500 &#181;L of BSA buffer and centrifuge at 1000 x g for 10 min at 4 &#176;C with full acceleration and full brake.</li>
	<li>Aspirate and discard the supernatant. 
	&#8203;NOTE: The pellet may not be visible; leave a small amount of buffer behind so as not to unintentionally aspirate the pellet. Cells can be resuspended in the appropriate buffer, blocked, and stained with cell-specific antibodies at this point. See Supplemental File 1 for sample protocol18.</li>
</ol>
10. Fixation (optional)
<ol>
	<li>Resuspend the pellet in 200 &#181;L of 1% PFA (previously prepared). Incubate for 15 min at 4 &#176;C.</li>
	<li>Wash by adding 500 &#181;L of D-PBS and centrifuge at 300 x g for 10 min at 4 &#176;C.</li>
	<li>Aspirate the supernatant. 
	NOTE: The pellet may not be visible; leave a small amount of buffer behind so as not to unintentionally aspirate the pellet.</li>
	<li>Resuspend the pellet in 200 &#181;L of D-PBS and store at 4 &#176;C for up to 3 days.</li>
</ol>
11. Flow cytometry
<ol>
	<li>Label the filter caps on the new tubes.</li>
	<li>Using a 1 mL pipette, pipette each sample onto the filter cap.</li>
	<li>Centrifuge briefly at 200 x g at 4 &#176;C, only allowing the centrifuge to reach 200 x g before stopping the run.</li>
	<li>Proceed to flow cytometry core for downstream analysis.</li>
</ol>

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The authors have nothing to disclose.

Several steps in this neural dissociation protocol require proficient technique and dexterity&#8211;perfusion, supernatant aspiration, and myelin removal. Throughout the perfusion process, the internal organs must remain intact (aside from removing the diaphragm and clipping the heart); this includes avoiding the upper chambers of the heart with the butterfly needle. While the amount of saline with heparin needed varies, transparent fluid flowing from the heart indicates the process is complete. The brain must be complet...

The hippocampus was first described by a Bolognese anatomist, Giulio Cesare Aranzio, in the 1500&#39;s1. In naming this newfound structure, Aranzio was likely inspired by its uncanny resemblance to the seahorse of the genus Hippocampus1. The hippocampus is involved in stress responses but is widely known for its role in learning and memory. More specifically, the hippocampus is responsible for the encoding and retrieval of declarative and spatial memory1.
The hippocampus, or hippocampus proper, is divided into the CA1 (cornu ammonis), CA2, and CA3 subfields1. Compared to the rest of the nervous system, the hippocampus has several unique defining characteristics, including its plasticity and potential for ongoing neurogenesis2. Neurogenesis is the process of the proliferation and differentiation of neural stem cells, followed by their integration into the pre-existing neuronal network. Neurogenesis is restricted to the subgranular zone of the dentate gyrus and subventricular zone of the lateral ventricles (and the olfactory bulbs)3. While neurogenesis is abundant in embryogenesis, it is a lifelong process3,4. As such, this discussion will focus on adult neurogenesis in the hippocampus.
The subventricular and subgranular zones are neurogenic niches containing ependymal and vascular cells, as well as immature and mature lineages of neural stem cells5. Microglia contribute to these niches as immune cells to regulate neurogenesis6. Neural progenitor cells are nonstem cell progenies of neural stem cells7. Three types of neural progenitors are present in the subventricular zone: radial glia-like type B cells, type C transit-amplifying progenitors, and type A neuroblasts3,8. The slowly dividing type B neural progenitor cells in the subventricular zone can differentiate into rapidly dividing type C cells8. Subsequently, type C cells differentiate into type A cells8. These neuroblasts migrate through the rostral migratory stream to the olfactory bulb before differentiating into interneurons or oligodendrocytes9. These olfactory bulb interneurons are key to olfactory short-term memory, and associative learning, whereas the oligodendrocytes myelinate axons of the corpus callosum9. The majority of adult neurogenesis occurs in the subgranular zone of the dentate gyrus, where radial type 1 and nonradial type 2 neural progenitors are found3. Most neural progenitor cells are destined to become dentate granule neurons and astrocytes10. Connected by gap junctions, astrocytes form networks to modulate plasticity, synaptic activity, and neuronal excitability5. As the primary excitatory neuron of the dentate gyrus, granule cells provide input from the entorhinal cortex to the CA3 region11.
Neural stem cell populations can be isolated using immunomagnetic or immunofluorescent isolation strategies12,13. Neural tissue is particularly difficult to dissociate; efforts to do so often result in samples with poor cell viability and/ or fail to produce the necessary single-cell suspension for downstream analysis. Neural dissociation can be conducted via mechanical dissociation (such as using filters, chopping techniques, or pipette trituration), enzymatic digestion, or a combination of techniques14,15. In a study evaluating neural dissociation methods, the viability and quality of manual mechanical dissociation by pipette trituration versus combinations of pipette trituration and digestion with various enzymes were compared15. Quality was graded based on the amount of cell clumps and DNA or subcellular debris in the prepared suspension15. Suspensions of glial tumors subjected to manual mechanical dissociation alone had significantly lower cell viability than treatments with dispase or a combination of DNase, collagenase, and hyaluronidase15. Volovitz et al. acknowledged the variation in viability and quality between the different methods and emphasized that inadequate dissociation may reduce the accuracy of downstream analysis15.
In a separate study, the authors compared over 60 different methods and combinations of dissociation of cultured neuronal cells14. These methods included eight different variations of manual mechanical dissociation by pipette trituration, a comparison of incubation with five individual enzymes at three different intervals, and various combinations of mechanical dissociation with enzymatic digestion or the combination of two enzymes14. None of the mechanical methods yielded a single-cell suspension14. Four of the single enzyme treatments, ten of the combination enzymatic treatments, and four of the combinations of mechanical dissociation with enzymatic digestion yielded a single-cell suspension14. Enzymatic digestion with TrypLE followed by Trypsin-EDTA most effectively dissociated samples14. Incidentally, samples treated with TrypLE and/or Trypsin-EDTA tended to form gelatinous clumps14. While this study was performed on cultured cells, it speaks to the shortcomings of pipette trituration or enzymatic digestion alone.
Side-by-side comparisons of manual versus automated mechanical dissociation are lacking. However, one group ran flow cytometry to compare manual and semi-automated mechanical dissociation of whole mouse brains in conjunction with commercial papain or trypsin enzymatic dissociation kits16. Processing with the dissociator more consistently yielded viable cells16. Following dissociation, the authors also isolated Prominin-1 cells, neuronal precursor cells, and microglia16. For two of the three isolated cell populations, the purity of the isolated cells was slightly higher when samples were processed with the dissociator, as compared to manually16. Rei&#223; et al. noted that person-to-person variability in pipetting technique hinders reproducibility of viable cell population yield in tissue dissociation16. The authors concluded that automated mechanical dissociation standardizes sample processing16.
The method of dissociation outlined in this manuscript is a combination of fully automated mechanical dissociation and enzymatic digestion, using solutions accompanying a commercial adult brain dissociation kit17. Unlike standard protocols, this optimized protocol reduces sample manipulation, yields a highly viable single-cell suspension, and is intended for processing minimal amounts of starting tissue.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
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			<td>1.5 mL Microcentrifuge Tubes</td>
			<td>Fisher Scientific</td>
			<td>02-682-003</td>
			<td>Basix, assorted color</td>
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		<tr>
			<td>15 mL Falcon Tubes</td>
			<td>Becton Dickinson Labware Europe</td>
			<td>352009</td>
			<td>Polystyrene</td>
		</tr>
		<tr>
			<td>25 mL Serological Pipets</td>
			<td>Fisher Scientific</td>
			<td>14-955-235</td>
			<td></td>
		</tr>
		<tr>
			<td>5 mL Round Bottom Polystyrene Test Tube</td>
			<td>Falcon</td>
			<td>352052</td>
			<td></td>
		</tr>
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			<td>500 mL Vacuum Filter/ Storage Bottle System</td>
			<td>Corning</td>
			<td>431097</td>
			<td></td>
		</tr>
		<tr>
			<td>70 &#956;m cell strainer</td>
			<td>Fisher Scientific</td>
			<td>08-771-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Adult Brain Dissociation Kit</td>
			<td>Miltenyi Biotec</td>
			<td>&#160;130-107-677</td>
			<td>Contains Enzyme P, Buffer Z, Buffer Y, Enzyme A, Buffer A, Debris Removal Solution</td>
		</tr>
		<tr>
			<td>Aluminum Foil</td>
			<td>Fisher Scientific</td>
			<td>01-213-105</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-ACSA-2-PE-Vio770, mouse, clone REA969</td>
			<td>Miltenyi Biotec</td>
			<td>130-116-246</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-Myelin Basic Protein</td>
			<td>Sigma-Aldrich</td>
			<td>M3821-100UG</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-PSA-NCAM-PE, human, mouse and rat, Clone 2-2B</td>
			<td>Miltenyi Biotec</td>
			<td>130-117-394</td>
			<td></td>
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		<tr>
			<td>BD LSRFortessa</td>
			<td>BD</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Sigma-Aldrich</td>
			<td>A7906-50G</td>
			<td></td>
		</tr>
		<tr>
			<td>CD11b-VioBlue, mouse, Clone REA592</td>
			<td>Miltenyi Biotec</td>
			<td>130-113-810</td>
			<td></td>
		</tr>
		<tr>
			<td>CD31 Antibody</td>
			<td>Miltenyi Biotec</td>
			<td>130-111-541</td>
			<td></td>
		</tr>
		<tr>
			<td>Ceramic Hot Plate Stirrer</td>
			<td>Fisher Scientific</td>
			<td>11-100-100SH</td>
			<td></td>
		</tr>
		<tr>
			<td>Dimethyl Sulfoxide</td>
			<td>Fisher Scientific</td>
			<td>BP231-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Pharmco by Greenfield Global</td>
			<td>111000200</td>
			<td></td>
		</tr>
		<tr>
			<td>Falcon 50 mL Conical Centrifuge Tubes</td>
			<td>Fisher Scientific</td>
			<td>14-432-22</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Scissors - Sharp</td>
			<td>Fine Science Tools</td>
			<td>14060-09</td>
			<td>Perfusion</td>
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		<tr>
			<td>FlowJo</td>
			<td>BD</td>
			<td></td>
			<td>(v10.7.0)</td>
		</tr>
		<tr>
			<td>gentleMACS C Tubes</td>
			<td>Miltenyi Biotec</td>
			<td>130-093-237</td>
			<td></td>
		</tr>
		<tr>
			<td>gentleMACS Octo Dissociator with Heaters</td>
			<td>Miltenyi Biotec</td>
			<td>130-096-427</td>
			<td></td>
		</tr>
		<tr>
			<td>Gibco DPBS (1X)</td>
			<td>ThermoFisher Scientific</td>
			<td>14190144</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Beaker</td>
			<td>Fisher Scientific</td>
			<td>02-555-25A</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin sodium</td>
			<td>Fresenius Kabi</td>
			<td>504011</td>
			<td></td>
		</tr>
		<tr>
			<td>LIVE/DEAD Fixable Aqua Dead Cell Stain Kit</td>
			<td>ThermoFisher</td>
			<td>L34965</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnetic Stir Bar</td>
			<td>Fisher Scientific</td>
			<td>14-513-51</td>
			<td></td>
		</tr>
		<tr>
			<td>Noyes Spring Scissors</td>
			<td>Fine Science Tools</td>
			<td>15012-12</td>
			<td>Dissection</td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma-Aldrich</td>
			<td>441244-3KG</td>
			<td>Prilled, 95%</td>
		</tr>
		<tr>
			<td>Pipette tips GP LTS 20 &#181;L 960A/10</td>
			<td>Rainin</td>
			<td>30389270</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette Tips GP LTS 250 &#181;L 960A/10</td>
			<td>Rainin</td>
			<td>30389277</td>
			<td></td>
		</tr>
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			<td>Pipette tips RT LTS 1000 &#181;L FL 768A/8</td>
			<td>Rainin</td>
			<td>30389213</td>
			<td></td>
		</tr>
		<tr>
			<td>Rainin Pipet-Lite XLS (2, 20, 200, 1000 &#956;L)</td>
			<td>Rainin</td>
			<td>30386597</td>
			<td></td>
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			<td>RBXMO FITC XADS</td>
			<td>Fisher Scientific</td>
			<td>A16167</td>
			<td></td>
		</tr>
		<tr>
			<td>Round Ice Bucket with Lid</td>
			<td>Fisher Scientific</td>
			<td>07-210-129</td>
			<td></td>
		</tr>
		<tr>
			<td>Round-Bottom Tubes with Cell Strainer Cap</td>
			<td>Falcon</td>
			<td>100-0087</td>
			<td></td>
		</tr>
		<tr>
			<td>S1 Pipet Fillers</td>
			<td>ThermoFisher Scientific</td>
			<td>9541</td>
			<td></td>
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			<td>Spatula &#38; Probe</td>
			<td>Fine Science Tools</td>
			<td>10090-13</td>
			<td>Dissection &#38; Perfusion</td>
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			<td>Surflo Winged Infusion Set 23 G x 3/4&#34;</td>
			<td>Termuno</td>
			<td>SV-23BLK</td>
			<td>Butterfly needle</td>
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		<tr>
			<td>Test Tube Rack</td>
			<td>Fisher Scientific</td>
			<td>14-809-37</td>
			<td></td>
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		<tr>
			<td>Thermo Scientific Legend XTR Centrifuge</td>
			<td>ThermoFisher</td>
			<td>discontinued</td>
			<td>Or other standard table top centrifuge</td>
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		<tr>
			<td>Variable-Flow Peristaltic Pump</td>
			<td>Fisher Scientific</td>
			<td>13-876-2</td>
			<td>Low-flow model</td>
		</tr>
		<tr>
			<td>VetFlo Starter Kit for Mice</td>
			<td>Kent Scientific</td>
			<td>VetFlo-MSEKIT</td>
			<td>Anesthesia mask, tubing, induction chamber, charcoal canisters</td>
		</tr>
		<tr>
			<td>VetFlo Vaporizer Single Channel Anesthesia System</td>
			<td>Kent Scientific</td>
			<td>VetFlo-1210S</td>
			<td>0.2&#8211;4 LPM</td>
		</tr>
		<tr>
			<td>Vi-CELL XR Cell Viability Analyzer</td>
			<td>Beckman Coulter Life Sciences</td>
			<td>731196</td>
			<td>Cell Counting</td>
		</tr>
		<tr>
			<td>Vi-CELL XR 4 Bags of Sample Vials</td>
			<td>Beckman Coulter Life Sciences</td>
			<td>383721</td>
			<td>Cell Counting</td>
		</tr>
	</tbody>
</table>

combined mechanical and enzymatic dissociation of mouse brain hippocampal tissue

This neural dissociation protocol (an adaptation of the protocol accompanying a commercial adult brain dissociation kit) optimizes tissue processing in preparation for detailed downstream analysis such as flow cytometry or single-cell sequencing. Neural dissociation can be conducted via mechanical dissociation (such as using filters, chopping techniques, or pipette trituration), enzymatic digestion, or a combination thereof. The delicate nature of neuronal cells can complicate efforts to obtain the highly viable, true single-cell suspension with minimal cellular debris that is required for single-cell analysis. The data demonstrate that this combination of automated mechanical dissociation and enzymatic digestion consistently yields a highly viable (&gt;90%) single-cell suspension, overcoming the aforementioned difficulties. While a few of the steps require manual dexterity, these steps lessen sample handling and potential cell loss. This manuscript details each step of the process to equip other laboratories to successfully dissociate small quantities of neural tissue in preparation for downstream analysis.

Samples were processed with a flow cytometer at a core facility, and the resulting data were evaluated with a software package for flow analysis. Previously, compensation controls were analyzed-the live/dead stain and negative control. If multiple fluorochromes are used, fluorescence minus one (FMO) controls and single-stain controls should be prepared for each antibody. Compensation for spectral overlap for the experimental samples was calculated based on the analyzed controls. For cell population identification, a hier...

Watch this Scientific Journal Video about Combined Mechanical and Enzymatic Dissociation of Mouse Brain Hippocampal Tissue at JoVE.com

Combined Mechanical and Enzymatic Dissociation of Mouse Brain Hippocampal Tissue

This neural cell dissociation protocol is intended for samples with a low amount of starting material and yields a highly viable single-cell suspension for downstream analysis, with optional fixation and staining steps.

This neural dissociation protocol (an adaptation of the protocol accompanying a commercial adult brain dissociation kit) optimizes tissue processing in ...

combined-mechanical-enzymatic-dissociation-mouse-brain-hippocampal

Research

JoVE Journal

Neuroscience

3.6K Views.  University of Arkansas for Medical Sciences. This neural cell dissociation protocol is intended for samples with a low amount of starting material and yields a highly viable single-cell suspension for downstream analysis, with optional fixation and staining steps.

Video: Combined Mechanical and Enzymatic Dissociation of Mouse Brain Hippocampal Tissue

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 Mouse Brain Tissue for Immunoelectron Microscopy

Transmission electron microscopy (TEM) is extremely useful for visualizing microglial, oligodendrocytic, astrocytic, and neuronal subcellular compartments (dendrite, dendritic spine, axon, axon terminal, perikaryon), as well as their intracellular organelles and cytoskeleton, in the central nervous system at high spatial resolution. Combined with TEM, pre-embedding immunocytochemistry allows the discrimination of cellular elements with few distinctive features and identification criteria (e.g., microglial perikarya and processes, when using an antibody against the microglia-specific marker Iba1 (ionized calcium binding adaptor molecule 1; as presented here)), identifying the neurotransmitter contents of cellular elements (e.g., serotonergic) and their ultrastructural localization of soluble or membrane-bound proteins (e.g., 5 HT1A and EphA4 receptors). Here, we describe a protocol for transcardiac perfusion of mice with acrolein fixative, removal and sectioning of the brain, as well as immunoperoxidase-diaminobenzidine (DAB) staining, resin embedding, and ultrathin sectioning of the brain sections. Upon completion of these procedures, the immunostained material is ready for examination with TEM. When rigorously performed, this technique provides an excellent compromise between optimal ultrastructural preservation and immunocytochemical detection.

We describe a protocol for transcardiac perfusion of mice, removal and sectioning of the brain, as well as immunoperoxidase staining, resin embedding, and ultrathin sectioning of the brain sections. Upon completion of these procedures, the immunostained material is ready for examination with transmission electron microscopy.

Transmission electron microscopy (TEM) is extremely useful for visualizing microglial, oligodendrocytic, astrocytic, and neuronal subcellular compartments ...

Nucleofection and Primary Culture of Embryonic Mouse Hippocampal and Cortical Neurons

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and synapse formation and function. An advantage of culturing these neurons is that they readily polarize, forming distinctive axons and dendrites, on a two dimensional substrate at very low densities. This property has made them extremely useful for determining many aspects of neuronal development. Furthermore, by providing glial conditioning for these neurons they will continue to develop, forming functional synaptic connections and surviving for several months in culture. In this protocol we outline a technique to dissect, culture and transfect embryonic mouse hippocampal and cortical neurons. Transfection is accomplished by electroporating DNA into the neurons before plating via nucleofection. This protocol has the advantage of expressing fluorescently-tagged fusion proteins early in development (~4-8hrs after plating) to study the dynamics and function of proteins during polarization, axon outgrowth and branching. We have also discovered that this single transfection before plating maintains fluorescently-tagged fusion protein expression at levels appropriate for imaging throughout the lifetime of the neuron (&gt; 2 months in culture). Thus, this methodology is useful for studying protein localization and function throughout CNS development with little or no disruption of neuronal function.

This protocol outlines the steps required to dissect, transfect via electroporation and culture mouse hippocampal and cortical neurons. Short-term cultures may be used for studies of axon outgrowth and guidance, while long-term cultures can be used for studies of synaptogenesis and dendritic spine analysis.

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and ...

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange (ABE)

Palmitoylation is a post-translational lipid modification involving the attachment of a 16-carbon saturated fatty acid, palmitate, to cysteine residues of substrate proteins through a labile thioester bond [reviewed in1]. Palmitoylation of a substrate protein increases its hydrophobicity, and typically facilitates its trafficking toward cellular membranes. Recent studies have shown palmitoylation to be one of the most common lipid modifications in neurons1, 2, suggesting that palmitate turnover is an important mechanism by which these cells regulate the targeting and trafficking of proteins. The identification and detection of palmitoylated substrates can therefore better our understanding of protein trafficking in neurons.
Detection of protein palmitoylation in the past has been technically hindered due to the lack of a consensus sequence among substrate proteins, and the reliance on metabolic labeling of palmitoyl-proteins with 3H-palmitate, a time-consuming biochemical assay with low sensitivity. Development of the Acyl-Biotin Exchange (ABE) assay enables more rapid and high sensitivity detection of palmitoylated proteins2-4, and is optimal for measuring the dynamic turnover of palmitate on neuronal proteins. The ABE assay is comprised of three biochemical steps (Figure 1): 1) irreversible blockade of unmodified cysteine thiol groups using N-ethylmaliemide (NEM), 2) specific cleavage and unmasking of the palmitoylated cysteine's thiol group by hydroxylamine (HAM), and 3) selective labeling of the palmitoylated cysteine using a thiol-reactive biotinylation reagent, biotin-BMCC. Purification of the thiol-biotinylated proteins following the ABE steps has differed, depending on the overall goal of the experiment.
Here, we describe a method to purify a palmitoylated protein of interest in primary hippocampal neurons by an initial immunoprecipitation (IP) step using an antibody directed against the protein, followed by the ABE assay and western blotting to directly measure palmitoylation levels of that protein, which is termed the IP-ABE assay. Low-density cultures of embryonic rat hippocampal neurons have been widely used to study the localization, function, and trafficking of neuronal proteins, making them ideally suited for studying neuronal protein palmitoylation using the IP-ABE assay. The IP-ABE assay mainly requires standard IP and western blotting reagents, and is only limited by the availability of antibodies against the target substrate. This assay can easily be adapted for the purification and detection of transfected palmitoylated proteins in heterologous cell cultures, primary neuronal cultures derived from various brain tissues of both mouse and rat, and even primary brain tissue itself.

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange ABE

The reversible addition of palmitate to proteins is an important regulator of intracellular protein trafficking. This is of particular interest in neurons where many synaptic proteins are palmitoylated. We utilize a simple biochemical method to detect palmitoylated proteins in cultured neurons, which can be adapted for multiple cell types and tissues.

Palmitoylation is a post-translational lipid  modification involving the attachment of a 16-carbon saturated fatty acid,  palmitate, to cysteine residues ...

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 ...

Modification of a Colliculo-thalamocortical Mouse Brain Slice, Incorporating 3-D printing of Chamber Components and Multi-scale Optical Imaging

The ability of the brain to process sensory information relies on both ascending and descending sets of projections. Until recently, the only way to study these two systems and how they interact has been with the use of in vivo preparations. Major advances have been made with acute brain slices containing the thalamocortical and cortico-thalamic pathways in the somatosensory, visual, and auditory systems. With key refinements to our recent modification of the auditory thalamocortical slice1, we are able to more reliably capture the projections between most of the major auditory midbrain and forebrain structures: the inferior colliculus (IC), medial geniculate body (MGB), thalamic reticular nucleus (TRN), and the auditory cortex (AC). With portions of all these connections retained, we are able to answer detailed questions that complement the questions that can be answered with in vivo preparations. The use of flavoprotein autofluorescence imaging enables us to rapidly assess connectivity in any given slice and guide the ensuing experiment. Using this slice in conjunction with recording and imaging techniques, we are now better equipped to understand how information processing occurs at each point in the auditory forebrain as information ascends to the cortex, and the impact of descending cortical modulation. 3-D printing to build slice chamber components permits double-sided perfusion and broad access to networks within the slice and maintains the widespread connections key to fully utilizing this preparation.

Using multiple angles to cut the mouse pup brain, we improve upon a previously-described acute brain slice which captures the connections between most of the major auditory midbrain and forebrain structures.

The ability of the brain to process sensory information relies on both ascending and descending sets of projections. Until recently, the only way to study ...

Combined Optogenetic and Freeze-fracture Replica Immunolabeling to Examine Input-specific Arrangement of Glutamate Receptors in the Mouse Amygdala

Freeze-fracture electron microscopy has been a major technique in ultrastructural research for over 40 years. However, the lack of effective means to study the molecular composition of membranes produced a significant decline in its use. Recently, there has been a major revival in freeze-fracture electron microscopy thanks to the development of effective ways to reveal integral membrane proteins by immunogold labeling. One of these methods is known as detergent-solubilized Freeze-fracture Replica Immunolabeling (FRIL).
The combination of the FRIL technique with optogenetics allows a correlated analysis of the structural and functional properties of central synapses. Using this approach it is possible to identify and characterize both pre- and postsynaptic neurons by their respective expression of a tagged channelrhodopsin and specific molecular markers. The distinctive appearance of the postsynaptic membrane specialization of glutamatergic synapses further allows, upon labeling of ionotropic glutamate receptors, to quantify and analyze the intrasynaptic distribution of these receptors. Here, we give a step-by-step description of the procedures required to prepare paired replicas and how to immunolabel them. We will also discuss the caveats and limitations of the FRIL technique, in particular those associated with potential sampling biases. The high reproducibility and versatility of the FRIL technique, when combined with optogenetics, offers a very powerful approach for the characterization of different aspects of synaptic transmission at identified neuronal microcircuits in the brain.
Here, we provide an example how this approach was used to gain insights into structure-function relationships of excitatory synapses at neurons of the intercalated cell masses of the mouse amygdala. In particular, we have investigated the expression of ionotropic glutamate receptors at identified inputs originated from the thalamic posterior intralaminar and medial geniculate nuclei. These synapses were shown to relay sensory information relevant for fear learning and to undergo plastic changes upon fear conditioning.

This article illustrates how the expression of neurotransmitter receptors can be quantified and the pattern analyzed at synapses with identified pre and postsynaptic elements using a combination of viral transduction of optogenetic tools and the freeze-fracture replica immunolabeling technique.

Freeze-fracture electron microscopy has been a major technique in ultrastructural research for over 40 years. However, the lack of effective means to ...

Characterizing Multiscale Mechanical Properties of Brain Tissue Using Atomic Force Microscopy, Impact Indentation, and Rheometry

To design and engineer materials inspired by the properties of the brain, whether for mechanical simulants or for tissue regeneration studies, the brain tissue itself must be well characterized at various length and time scales. Like many biological tissues, brain tissue exhibits a complex, hierarchical structure. However, in contrast to most other tissues, brain is of very low mechanical stiffness, with Young&#39;s elastic moduli E on the order of 100s of Pa. This low stiffness can present challenges to experimental characterization of key mechanical properties. Here, we demonstrate several mechanical characterization techniques that have been adapted to measure the elastic and viscoelastic properties of hydrated, compliant biological materials such as brain tissue, at different length scales and loading rates. At the microscale, we conduct creep-compliance and force relaxation experiments using atomic force microscope-enabled indentation. At the mesoscale, we perform impact indentation experiments using a pendulum-based instrumented indenter. At the macroscale, we conduct parallel plate rheometry to quantify the frequency dependent shear elastic moduli. We also discuss the challenges and limitations associated with each method. Together these techniques enable an in-depth mechanical characterization of brain tissue that can be used to better understand the structure of brain and to engineer bio-inspired materials.

We present a set of techniques to characterize the viscoelastic mechanical properties of brain at the micro-, meso-, and macro-scales.

To design and engineer materials inspired by the properties of the brain, whether for mechanical simulants or for tissue regeneration studies, the brain ...

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 ...

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 ...