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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<li>Delmonte, O. M., Fleisher, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Flow+cytometry%3A+Surface+markers+and+beyond%5BTitle%5D)+AND+%22The+Journal+of+Allergy+and+Clinical+Immunology%22%5BJournal%5D)">Flow cytometry: Surface markers and beyond.</a> The Journal of Allergy and Clinical Immunology. 143, 528-537 (2019).</li>
<li>O'Connor, J. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+relevance+of+flow+cytometry+for+biochemical+analysis%5BTitle%5D)+AND+%22IUBMB+Life%22%5BJournal%5D)">The relevance of flow cytometry for biochemical analysis.</a> IUBMB Life. 51, 231-239 (2001).</li>
<li>Menon, V., Thomas, R., Ghale, A. R., Reinhard, C., Pruszak, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Flow+cytometry+protocols+for+surface+and+intracellular+antigen+analyses+of+neural+cell+types%5BTitle%5D)+AND+%22Journal+of+Visualized.+Experiments%22%5BJournal%5D)">Flow cytometry protocols for surface and intracellular antigen analyses of neural cell types.</a> Journal of Visualized. Experiments. (94), e52241(2014).</li>
</ol>

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><tbody><tr><td>1.5 mL Microcentrifuge Tubes</td><td>Fisher Scientific</td><td>02-682-003</td><td>Basix, assorted color</td></tr><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><tr><td>500 mL Vacuum Filter/ Storage Bottle System</td><td>Corning</td><td>431097</td><td></td></tr><tr><td>70 &amp;mu;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>&amp;nbsp;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></tr><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></tr><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 &amp;micro;L 960A/10</td><td>Rainin</td><td>30389270</td><td></td></tr><tr><td>Pipette Tips GP LTS 250 &amp;micro;L 960A/10</td><td>Rainin</td><td>30389277</td><td></td></tr><tr><td>Pipette tips RT LTS 1000 &amp;micro;L FL 768A/8</td><td>Rainin</td><td>30389213</td><td></td></tr><tr><td>Rainin Pipet-Lite XLS (2, 20, 200, 1000 &amp;mu;L)</td><td>Rainin</td><td>30386597</td><td></td></tr><tr><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></tr><tr><td>Spatula &amp;amp; Probe</td><td>Fine Science Tools</td><td>10090-13</td><td>Dissection &amp;amp; Perfusion</td></tr><tr><td>Surflo Winged Infusion Set 23 G x 3/4&quot;</td><td>Termuno</td><td>SV-23BLK</td><td>Butterfly needle</td></tr><tr><td>Test Tube Rack</td><td>Fisher Scientific</td><td>14-809-37</td><td></td></tr><tr><td>Thermo Scientific Legend XTR Centrifuge</td><td>ThermoFisher</td><td>discontinued</td><td>Or other standard table top centrifuge</td></tr><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&amp;ndash;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

3.7K 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

An Optogenetic Approach for Assessing Formation of Neuronal Connections in a Co-culture System

Here we describe a protocol to generate a co-culture consisting of 2 different neuronal populations. Induced pluripotent stem cells (iPSCs) are reprogrammed from human fibroblasts using episomal vectors. Colonies of iPSCs can be observed 30 days after initiation of fibroblast reprogramming. Pluripotent colonies are manually picked and grown in neural induction medium to permit differentiation into neural progenitor cells (NPCs). iPSCs rapidly convert into neuroepithelial cells within 1 week and retain the capability to self-renew when maintained at a high culture density. Primary mouse NPCs are differentiated into astrocytes by exposure to a serum-containing medium for 7 days and form a monolayer upon which embryonic day 18 (E18) rat cortical neurons (transfected with channelrhodopsin-2 (ChR2)) are added. Human NPCs tagged with the fluorescent protein, tandem dimer Tomato (tdTomato), are then seeded onto the astrocyte/cortical neuron culture the following day and allowed to differentiate for 28 to 35 days. We demonstrate that this system forms synaptic connections between iPSC-derived neurons and cortical neurons, evident from an increase in the frequency of synaptic currents upon photostimulation of the cortical neurons. This co-culture system provides a novel platform for evaluating the ability of iPSC-derived neurons to create synaptic connections with other neuronal populations.

A protocol to generate a co-culture system consisting of neurons derived from induced pluripotent stem cells (iPSCs), primary cortical neurons and astrocytes is described. This co-culture system allows detection of the formation of synaptic contacts and circuits between new, iPSC-derived neurons and pre-existing cortical neurons expressing channelrhodopsin-2.

Here we describe a protocol to generate a co-culture consisting of 2 different neuronal populations. Induced pluripotent stem cells (iPSCs) are ...

Developmental Biology

Sex Stratified Neuronal Cultures to Study Ischemic Cell Death Pathways

Sex differences in neuronal susceptibility to ischemic injury and neurodegenerative disease have long been observed, but the signaling mechanisms responsible for those differences remain unclear. Primary disassociated embryonic neuronal culture provides a simplified experimental model with which to investigate the neuronal cell signaling involved in cell death as a result of ischemia or disease; however, most neuronal cultures used in research today are mixed sex. Researchers can and do test the effects of sex steroid treatment in mixed sex neuronal cultures in models of neuronal injury and disease, but accumulating evidence suggests that the female brain responds to androgens, estrogens, and progesterone differently than the male brain. Furthermore, neonate male and female rodents respond differently to ischemic injury, with males experiencing greater injury following cerebral ischemia than females. Thus, mixed sex neuronal cultures might obscure and confound the experimental results; important information might be missed. For this reason, the Herson Lab at the University of Colorado School of Medicine routinely prepares sex-stratified primary disassociated embryonic neuronal cultures from both hippocampus and cortex. Embryos are sexed before harvesting of brain tissue and male and female tissue are disassociated separately, plated separately, and maintained separately. Using this method, the Herson Lab has demonstrated a male-specific role for the ion channel TRPM2 in ischemic cell death. In this manuscript, we share and discuss our protocol for sexing embryonic mice and preparing sex-stratified hippocampal primary disassociated neuron cultures. This method can be adapted to prepare sex-stratified cortical cultures and the method for embryo sexing can be used in conjunction with other protocols for any study in which sex is thought to be an important determinant of outcome.

Primary disassociated embryonic hippocampal neuronal cultures are useful for investigating the signaling mechanisms involved in neuron death. Sexing the embryos before the isolation and dissociation of the hippocampus allows the preparation of separate male and female cultures, which enables the researcher to identify and investigate sex-specific cell signaling.

Sex differences in neuronal susceptibility to ischemic injury and neurodegenerative disease have long been observed, but the signaling mechanisms ...

Acute Mouse Brain Slicing to Investigate Spontaneous Hippocampal Network Activity

Acute rodent brain slicing offers a tractable experimental approach to gain insight into the organization and function of neural circuits with single-cell resolution using electrophysiology, microscopy, and pharmacology. However, a major consideration in the design of in vitro experiments is the extent to which different slice preparations recapitulate naturalistic patterns of neural activity as observed in vivo. In the intact brain, the hippocampal network generates highly synchronized population activity reflective of the behavioral state of the animal, as exemplified by the sharp-wave ripple complexes (SWRs) that occur during waking consummatory states or non-REM sleep. SWRs and other forms of network activity can emerge spontaneously in isolated hippocampal slices under appropriate conditions. In order to apply the powerful brain slice toolkit to the investigation of hippocampal network activity, it is necessary to utilize an approach that optimizes tissue health and the preservation of functional connectivity within the hippocampal network. Mice are transcardially perfused with cold sucrose-based artificial cerebrospinal fluid. Horizontal slices containing the hippocampus are cut at a thickness of 450 &#956;m to preserve synaptic connectivity. Slices recover in an interface-style chamber and are transferred to a submerged chamber for recordings. The recording chamber is designed for dual surface superfusion of artificial cerebrospinal fluid at a high flow rate to improve oxygenation of the slice. This protocol yields healthy tissue suitable for the investigation of complex and spontaneous network activity in vitro.

This protocol describes the preparation of horizontal hippocampal-entorhinal cortex (HEC) slices from mice exhibiting spontaneous sharp-wave ripple activity. Slices are incubated in a simplified interface holding chamber and recordings are performed under submerged conditions with fast-flowing artificial cerebrospinal fluid to promote tissue oxygenation and the spontaneous emergence of network-level activity.

Acute rodent brain slicing offers a tractable experimental approach to gain insight into the organization and function of neural circuits with single-cell ...

Microdissection of Mouse Brain into Functionally and Anatomically Different Regions

The brain is the command center for the mammalian nervous system and an organ with enormous structural complexity. Protected within the skull, the brain consists of an outer covering of grey matter over the hemispheres known as the cerebral cortex. Underneath this layer reside many other specialized structures that are essential for multiple phenomenon important for existence. Acquiring samples of specific gross brain regions requires quick and precise dissection steps.&#160;It is understood that at the microscopic level, many sub-regions exist and likely cross the arbitrary regional boundaries that we impose for the purpose of this dissection.
Mouse models are routinely used to study human brain functions and diseases. Changes in gene expression patterns may be confined to specific brain areas targeting a particular phenotype depending on the diseased state. Thus, it is of great importance to study regulation of transcription with respect to its well-defined structural organization. A complete understanding of the brain requires studying distinct brain regions, defining connections, and identifying key differences in the activities of each of these brain regions. A more comprehensive understanding of each of these distinct regions may pave the way for new and improved treatments in the field of neuroscience. Herein, we discuss a step-by-step methodology for dissecting the mouse brain into sixteen distinct regions. In this procedure, we have focused on male mouse C57Bl/6J (6-8 week old) brain removal and dissection into multiple regions using neuroanatomical landmarks to identify and sample discrete functionally-relevant and behaviorally-relevant&#160;brain regions. This work will help lay a strong foundation in the field of neuroscience, leading to more focused approaches in the deeper understanding of brain function.

We present a hands-on, step-by-step, rapid protocol for mouse brain removal and dissection of discrete regions from fresh brain tissue. Obtaining brain regions for molecular analysis has become routine in many neuroscience labs. These brain regions are immediately frozen to obtain high quality transcriptomic data for system level analysis.

The brain is the command center for the mammalian nervous system and an organ with enormous structural complexity. Protected within the skull, the brain ...

Hippocampal Neuronal Cultures to Detect and Study New Pathogenic Antibodies Involved in Autoimmune Encephalitis

Over the last 15 years, a new category of antibody-mediated diseases of the central nervous system (CNS) has been characterized and is now defined as &#34;autoimmune encephalitis&#34; (AE). There are currently 17 known AE syndromes, and all are associated with antibodies against the neuronal cell surface or synaptic proteins. The clinical syndromes are complex and vary according to the type of associated antibody. The best-known of these diseases is anti-N-methyl D-aspartate receptor (NMDAR) encephalitis, which is a prominent neuropsychiatric disorder associated with severe memory and behavioral impairments. The associated antibodies react with the GluN1 subunit of the NMDAR at the N-terminal domain. The approach most frequently used for the discovery and characterization of AE antibodies includes the culture of dissociated, fetal, rodent hippocampal neurons. During the process of antibody characterization, live neurons in culture are exposed to patients&#39; serum or CSF, and the detection of reactivity indicates that the serum or CSF samples of the patient contain antibodies against neuronal surface antigens. Hippocampal cultures can also be used to determine whether the antibodies in patients are potentially pathogenic by examining if they cause structural or functional alterations of the neurons. The level of success of these studies depends on the quality of the cultures and on the protocols used to obtain and detect the reactivity of patient samples. This article provides an optimized protocol for primary cell culture of fetal rat hippocampal neurons combined with immunostaining to determine the presence of antibodies in the serum or CSF of patients. An example of how to examine the potential pathogenic effects of NMDAR antibodies using cultured neurons and calcium imaging is also presented.

Autoimmune encephalitis is a new category of antibody-mediated diseases of the central nervous system. Hippocampal neurons can be used to discover and characterize these antibodies. This article provides a protocol for primary cell culture and immunostaining to determine autoantibodies in the serum and cerebrospinal fluid of patients.

Over the last 15 years, a new category of antibody-mediated diseases of the central nervous system (CNS) has been characterized and is now defined as ...

Visualizing Proteins in Primary Hippocampal Neurons by Live&#45;Cell Imaging

Using Live&#45;Cell Imaging to Measure the Effects of Pathological Proteins on Axonal Transport in Primary Hippocampal Neurons

Bidirectional transport of cargos along the axon is critical for maintaining functional synapses, neural connectivity, and healthy neurons. Axonal transport is disrupted in multiple neurodegenerative diseases, and projection neurons are particularly vulnerable because of the need to transport cellular materials over long distances and sustain substantial axonal mass. Pathological modifications of several disease-related proteins negatively affect transport, including tau, amyloid-&#946;, &#945;-synuclein, superoxide dismutase, and huntingtin, providing a potential common mechanism by which pathological proteins exert toxicity in disease. Methods to study these toxic mechanisms are necessary to understand neurodegenerative disorders and identify potential therapeutic interventions.
Here, cultured primary rodent hippocampal neurons are co-transfected with multiple plasmids to study the effects of pathological proteins on fast axonal transport using live-cell confocal imaging of fluorescently-tagged cargo proteins. We begin with the harvest, dissociation, and culturing of primary hippocampal neurons from rodents. Then, we co-transfect the neurons with plasmid DNA constructs to express fluorescent-tagged cargo protein and wild-type or mutant tau (used as an exemplar of pathological proteins). Axons are identified in live cells using an antibody that binds an extracellular&#160;domain of neurofascin, an axon initial segment protein, and an axonal region of interest is imaged to measure fluorescent cargo transport.
Using KymoAnalyzer, a freely available ImageJ macro, we extensively characterize the velocity, pause frequency, and directional cargo density of axonal transport, all of which may be affected by the presence of pathological proteins. Through this method, we identify a phenotype of increased cargo pause frequency associated with the expression of pathological tau protein. Additionally, gene-silencing shRNA constructs can be added to the transfection mix to test the role of other proteins in mediating transport disruption. This protocol is easily adaptable for use with other neurodegenerative disease-related proteins and is a reproducible method to study the mechanisms of how those proteins disrupt axonal transport.

Author Spotlight: Understanding the Impact of Pathological Proteins on Axonal Transport in Neurodegenerative Diseases

Here, we demonstrate how to combine transfection of primary hippocampal rodent neurons with live-cell confocal imaging to analyze pathological protein-induced effects on axonal transport and identify mechanistic pathways mediating these effects.

Bidirectional transport of cargos along the axon is critical for maintaining functional synapses, neural connectivity, and healthy neurons. Axonal ...

Electrophysiological Recording of Hippocampal Microglia from Adult Mice

Isolation and Whole-Cell Patch-Clamp Recording of Hippocampal Microglia from Adult Mice

Microglia are resident immune cells in the brain that interact with neurons to maintain the homeostasis of the central nervous system (CNS). Studies show that the microglial surface expresses potassium channels that regulate microglial activation, while abnormalities in these potassium channels can lead to neural diseases. Currently, whole-cell patch-clamp recordings of microglia are mostly performed on cultured primary microglia from fetal or newborn mice due to difficulties in conducting electrophysiological evaluations on acutely isolated microglia. This study introduces an easy-to-follow protocol for isolating hippocampal microglia from adult mice and performing whole-cell patch-clamp recordings on the isolated cells. Briefly, the brain was removed from a mouse after decapitation, the hippocampus was dissected bilaterally, and microglia were isolated using an adult mouse brain dissociation kit. The microglia were then purified using a magnetic-activated cell sorting (MACS) method and seeded onto coverslips. Successful microglial isolation was confirmed by immunofluorescent staining with anti-CD11 and anti-Iba1 antibodies. A cover slip was placed in a recording chamber, and the whole-cell potassium currents of the acutely isolated microglia were recorded under voltage-clamp conditions.

The present protocol describes how to isolate and purify primary hippocampal microglia from adult mice, followed by instructions for conducting whole-cell patch-clamp recordings on these acutely isolated cells.

Microglia are resident immune cells in the brain that interact with neurons to maintain the homeostasis of the central nervous system (CNS). Studies show ...

Biology

Obtaining a Single-Cell Suspension from Mouse Hippocampal Tissue

Source: Trujillo, M., et al. <a href="https://app.jove.com/v/63007">Combined Mechanical and Enzymatic Dissociation of Mouse Brain Hippocampal Tissue.</a> J. Vis. Exp. (2021)
This video demonstrates a protocol to obtain a highly viable single-cell suspension from the mouse brain hippocampus using a combination of automated mechanical dissociation and a mixture of enzymes to aid in enzymatic digestion.

All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee. 
1. Prepare Enzyme Mix 1 and 2 ...

JoVE Encyclopedia of Experiments

Neuronal Culture Techniques

Primary Neuronal Culture

Isolating Microglia from Mouse Brain Demyelinating Lesions Using Magnetic Activated Cell Sorting

Source: Zhang, H. et al. <a href="https://app.jove.com/v/63511">Isolation of Mouse Primary Microglia by Magnetic-Activated Cell Sorting in Animal Models of Demyelination.</a> J. Vis. Exp. (2022).
This video demonstrates a protocol for extracting microglial cells from demyelinating lesions in mouse brains using magnetic cell sorting. The protocol involves dissecting the lesioned brain tissue containing microglial cells, treating the tissue with specific enzymes to obtain a single-cell suspension, and utilizing microglial receptor-specific magnetic beads to isolate the microglial cells.

 All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee. 
1. Materials

	&#160;Prepare ...

Isolation of Multiple Cell Types from a Mouse Brain by Mechanical Homogenization

Source: Molina Estevez, F. J., et al. <a href="https://app.jove.com/v/60335">Simultaneous Flow Cytometric Characterization of Multiple Cell Types Retrieved from Mouse Brain/Spinal Cord Through Different Homogenization Methods.</a> J. Vis. Exp. (2019).
This video demonstrates the isolation of multiple types of cells from mouse brain tissue. Tissue is first mechanically sheared using a tissue grinder in a salt solution to maintain an osmotic balance. The tissue homogenate is then subjected to density gradient centrifugation to remove cellular debris. Cells are then resuspended in a solution for further analysis.

Disclaimer: &#160;All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1. Mechanical ...