We thank Kathy Gage for her excellent editorial contribution. The illustration&#160;figures were&#160;created with BioRender.com. This study was supported by funds from the Department of Anesthesiology (Duke University Medical Center) and NIH grants NS099590, HL157354, and NS127163.

The protocol was approved by the Duke Institute Animal Care and Use Committee (IACUC). Male C57Bl/6 mice (3-4 months old; 22-28 g) were used in the current study. The details of the reagents and the equipment used are listed in the Table of Materials.
1. Single-cell suspension from mouse brain
NOTE: Figure 1 illustrates the overview of the brain cell isolation protocol.
<ol>
	<li>Anesthetize, intubate, and secure the mouse, as demonstrated previously11.</li>
	<li>Make an upper abdominal incision through the skin and underlying muscle layers, and carefully cut through the diaphragm and ribs to create the opening into the thoracic cavity.</li>
	<li>Fill a 30 mL syringe with cold PBS and attach it to the blunt perfusion needle via silicone tubing. Carefully insert the perfusion needle into the left ventricle.</li>
	<li>Perform a small nick at the right atrium to allow blood drainage and then perfuse12 the mouse at a flow rate of ~ 10 mL/min for 3-4 min. 
	NOTE: To improve blood clearance in the brain, increasing the volume of perfusion solution and adding heparin may be considered.</li>
	<li>Decapitate the mouse and make a midline incision through the skin on the top of the head to expose the skull.</li>
	<li>Carefully scrape away any connective tissue overlying the skull using blunt forceps.</li>
	<li>Use sharp dissection scissors to carefully cut through the skull sutures along the midline and on both sides, creating a flap.</li>
	<li>Gently lift the skull flap to expose the brain, and use blunt forceps to carefully remove the brain from the cranial cavity.</li>
	<li>Immerse the brain in a Petri dish containing ice-cold PBS and remove any remaining meninge tissues. 
	NOTE: To better preserve cell viability, it is recommended to perform all following steps on ice or at 4 &#176;C.</li>
	<li>Place the brain in a pre-cooled 15 mL glass Dounce homogenizer containing 20 mL (whole brain) or 12 mL (half brain) of ice-old HBSS buffer.</li>
	<li>Use the loose Dounce pestle to gently and slowly dissociate the brain tissue with around 100-120 strokes on ice. 
	NOTE: This step is critical to obtaining a large quantity of healthy brain cells. This step takes approximately 10 min. Stop homogenization when no substantial brain tissue chunks are visible in the suspension. At this point, the pestle can easily move down without any force.</li>
	<li>Filter the resulting brain tissue suspension through a 70 &#181;m cell strainer in a 50 mL centrifuge tube.</li>
	<li>Rinse the Dounce tube with 5 mL HBSS, and proceed with filtration to maximize the cell collection.</li>
	<li>Centrifuge the filtered tissue suspension at 550 x g for 6 min at 4 &#176;C.</li>
	<li>Remove the supernatant using a vacuum aspirator, and resuspend the cell pellet in 1 mL of 30% isotonic density gradient solution. 
	NOTE: To make 15 mL of 30% isotonic density gradient solution, mix 4.5 mL of 100% stock density gradient medium, 1.5 mL of 10&#215; PBS, and 9 mL of ddH2O.</li>
	<li>Transfer the cell suspension into a 15 mL centrifuge tube, and add 9 mL (whole brain) or 6 mL (half brain) of 30% density gradient solution.</li>
	<li>Gently invert the tube to ensure thorough mixing.</li>
	<li>Immediately centrifuge the tube at 845 x g with acceleration and brake set at level 3, for 20 min at 4 &#176;C.</li>
	<li>Gently remove the tube from the centrifuge without shaking, and carefully aspirate the upper myelin layer using a 1 mL tip connected to the vacuum. 
	NOTE: It is critically important not to disturb this layer. Any remaining myelin can negatively impact cell health and compromise downstream analyses.</li>
	<li>Aspirate the supernatant, leaving 1-2 mL behind, and resuspend the cell pellet with a minimum of 10 mL cold HBSS.</li>
	<li>Centrifuge at 550 x g for 6 min at 4 &#176;C.</li>
	<li>Wash one more time with a minimum of 7 mL of cold HBSS.</li>
	<li>Remove as much of the supernatant as possible, and resuspend the cells for flow cytometry analysis using 100-200 &#181;L of flow cytometry buffer. 
	NOTE: The flow cytometry FACS buffer used for this study contains 0.5% BSA and 2 mM EDTA in 1&#215; PBS. A 96-well V-bottom plate for cell staining is recommended to minimize cell loss during the washing steps.</li>
	<li>Perform flow cytometry analysis using a standard protocol established in individual laboratories13.
	<ol>
		<li>Briefly, mix 80 &#181;L of cell suspension with 20 &#181;L of the antibody mixture in a 96-well V-bottom plate and incubate for 25 min at 4 &#176;C.</li>
		<li>Wash with 200 &#181;L of FACS buffer and then resuspend the cells with 100 &#181;L of HBSS plus cell viability staining reagent.</li>
		<li>After a 15 min incubation at 4 &#176;C, wash with FACS buffer and resuspend the cells in 200 &#181;L of FACS buffer for flow cytometry analysis.</li>
	</ol>
	</li>
</ol>
2. Preparation of bone marrow single-cell suspension from mouse calvaria
NOTE: Figure 2 depicts the overview of the skull bone marrow isolation procedure.
<ol>
	<li>Perfuse the mouse as described in step 1.4.</li>
	<li>Decapitate the mouse and make a midline skin incision on the scalp to expose the skull.</li>
	<li>Cut through the skull, starting from the foramen magnum and moving forward along the lateral sides of the calvaria using sharp scissors. Then, lift and remove the calvaria. 
	NOTE: The brain can be harvested from the same mouse and processed using step 1, enabling analysis of immune cells from both the brain and skull bone marrow.</li>
	<li>Carefully peel off the dura mater from the skull under a dissecting microscope using blunt forceps. 
	NOTE: It is easier to start at the edges of the occipital bone, slowly detach, and progress steadily towards the frontal area, working inward.</li>
	<li>Place the calvaria in the dish on ice and use a 1 mL pipette to rinse the calvaria, ensuring the removal of blood residues from its surface.</li>
	<li>Transfer the calvaria to a new dish with fresh, cold PBS sufficient to immerse the calvaria.</li>
	<li>Use sterile scissors to cut the calvaria into approximately 3 mm x 3 mm fragments in cold PBS. 
	NOTE: Cropping methods may affect the efficiency of calvaria bone marrow extraction. It is recommended that the same cropping method be used to minimize variability. It is also important to cut open the sutures. Figure 2B illustrates one approach.</li>
	<li>Poke a hole in the bottom of a 500 &#181;L microcentrifuge tube using an 18 G needle.</li>
	<li>Insert this tube into a 1.5 mL microcentrifuge tube containing 20 &#181;L of cold PBS.</li>
	<li>Transfer the calvaria fragments into the 500 &#181;L tube. 
	NOTE: The bone fragments should be loosely packed to increase extraction efficiency.</li>
	<li>Centrifuge at 10,000 x g for 30 s at 4 &#176;C. 
	NOTE: To maximize cell recovery, use an 18 G needle to gently stir the calvaria fragments, and repeat this step 1-2 more times.</li>
	<li>Resuspend the collected calvaria bone marrow cells in 500 &#181;L of Red Blood Cell lysis buffer and incubate at room temperature for 30 s.</li>
	<li>Transfer the suspension to a 15 mL centrifuge tube containing 4 mL of cold PBS.</li>
	<li>Centrifuge at 400 x g for 6 min at 4 &#176;C.</li>
	<li>Wash one more time with 5 mL of cold PBS.</li>
	<li>Resuspend the cell pellet in an appropriate buffer. For flow cytometry, we typically use 500 &#181;L of FACS buffer.</li>
	<li>For flow cytometry analysis, perform cell counting and use approximately 1 x 106 cells for antibody staining. 
	NOTE: Ensure to include the Fc block step before antibody staining13.</li>
</ol>

Obtaining Single-Cell Suspensions from Murine Brain and Skull Bone Marrow

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E., Garcia-Bonilla, L., Iadecola, C., Anrather, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immune+compartments+at+the+brain's+borders+in+health+and+neurovascular+diseases.">Immune compartments at the brain's borders in health and neurovascular diseases.</a> Semin Immunopathol. 45 (3), 437-449 (2023).</li><li>Herisson, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+vascular+channels+connect+skull+bone+marrow+and+the+brain+surface+enabling+myeloid+cell+migration.">Direct vascular channels connect skull bone marrow and the brain surface enabling myeloid cell migration.</a> Nat Neurosci. 21 (9), 1209-1217 (2018).</li><li>Mazzitelli, J. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Skull+bone+marrow+channels+as+immune+gateways+to+the+central+nervous+system.">Skull bone marrow channels as immune gateways to the central nervous system.</a> Nat Neurosci. 26 (12), 2052-2062 (2023).</li><li>Cugurra, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Skull+and+vertebral+bone+marrow+are+myeloid+cell+reservoirs+for+the+meninges+and+CNS+parenchyma.">Skull and vertebral bone marrow are myeloid cell reservoirs for the meninges and CNS parenchyma.</a> Science. 373 (6553), 7844 (2021).</li><li>Brioschi, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Heterogeneity+of+meningeal+b+cells+reveals+a+lymphopoietic+niche+at+the+CNS+borders.">Heterogeneity of meningeal b cells reveals a lymphopoietic niche at the CNS borders.</a> Science. 373 (6553), 9277 (2021).</li><li>Smyth, L. C. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+direct+connections+between+the+dura+and+the+brain.">Identification of direct connections between the dura and the brain.</a> Nature. 627, 165-173 (2024).</li><li>Mazzitelli, J. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cerebrospinal+fluid+regulates+skull+bone+marrow+niches+via+direct+access+through+dural+channels.">Cerebrospinal fluid regulates skull bone marrow niches via direct access through dural channels.</a> Nat Neurosci. 25 (5), 555-560 (2022).</li><li>Pulous, F. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cerebrospinal+fluid+can+exit+into+the+skull+bone+marrow+and+instruct+cranial+hematopoiesis+in+mice+with+bacterial+meningitis.">Cerebrospinal fluid can exit into the skull bone marrow and instruct cranial hematopoiesis in mice with bacterial meningitis.</a> Nat Neurosci. 25 (5), 567-576 (2022).</li><li>Li, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mouse+cardiac+arrest+model+for+brain+imaging+and+brain+physiology+monitoring+during+ischemia+and+resuscitation.">Mouse cardiac arrest model for brain imaging and brain physiology monitoring during ischemia and resuscitation.</a> J Vis Exp. 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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protocol+for+analysis+of+mouse+neutrophil+netosis+by+flow+cytometry.">Protocol for analysis of mouse neutrophil netosis by flow cytometry.</a> STAR Protoc. 2 (4), 100948 (2021).</li><li>Su, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Meningeal+immunity+and+neurological+diseases:+New+approaches,+new+insights.">Meningeal immunity and neurological diseases: New approaches, new insights.</a> J Neuroinflammation. 20 (1), 125 (2023).</li><li>Niu, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanical+isolation+of+neonatal+and+adult+mouse+dura+leukocytes+for+flow+cytometry+analysis.">Mechanical isolation of neonatal and adult mouse dura leukocytes for flow cytometry analysis.</a> STAR Protoc. 4 (2), 102272 (2023).</li><li>Roussel-Queval, A., Rebejac, J., Eme-Scolan, E., Paroutaud, L. 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Here, two simple yet effective protocols are presented for isolating immune cells from the brain and skull bone marrow. These protocols can reliably yield a large quantity of viable immune cells that may be suitable for diverse downstream applications, in particular for flow cytometry.
To study neuroinflammation in various brain disorders, many protocols for immune cell preparations from the brain have been established and used in different laboratories15,16,17. The common approaches involve enzyme-based cell dissociation followed by gradient separation. Typically, in these protocols, brain tissue is minced and then incubated in an enzyme digestion solution containing both collagenase and DNase for 30-60 min at 37 &#176;C with gentle shaking. After digestion, cells undergo further dissociation using a mechanical method such as pipetting or Dounce homogenization. After passing through a cell strainer (e.g., 70 &#181;m strainer), cells are then subjected to gradient centrifugation (often 30%/70% density gradient solution) to remove myelin, red blood cells, and debris and to enrich leukocytes. However, this approach requires a prolonged enzymatic incubation at 37 &#176;C that may substantially alter the transcriptome and proteome of brain immune cells. Indeed, a systematic investigation suggested that the enzymatic approach introduces profound alterations in the transcriptome and proteotype of specific brain cells18. This could lead to artifacts in subsequent transcriptomic analyses such as scRNA-seq. In contrast, the protocol reported here uses a mechanical dissociation approach at cold temperatures throughout the procedure, thus suppressing cellular metabolism. This ensures better preservation of the transcriptional and proteomic state, minimizing potential artifacts. During protocol development, we noticed that using a large Dounce homogenizer for homogenizing brain tissue is critical to achieving higher cell counts with lower cell death. It is likely that during Dounce homogenization, the massive presence of extracellular matrices, cell debris, and cytoplasmic components is toxic to cells. Using an excessive amount of buffer helps to dilute these toxic substances and, thus, mitigates their harmful effects, leading to better cell viability. The flow cytometry data confirmed an approximate 85% viability rate of immune cells prepared with this protocol. Moreover, it is notable that centrifugation with only a 30% density gradient solution effectively enriches brain immune cells and removes myelin. This can also reduce the cost of cell preparation.
Studies on immune cells in the skull bone marrow are still limited. A method commonly used to isolate cells from the mouse skull calvarium involves cutting the skull into small pieces and then mechanically crushing the pieces with a pestle6. In the current protocol, brief centrifugation is used. This method is adapted from a published protocol for preparing bone marrow from femurs and tibias19. For these long bones, one needs only to cut off one end of the bone to collect bone marrow by centrifugation. For the skull, this step is modified by cutting it into several small pieces to open the marrow cavity. It is recommended to use a consistent pattern for cutting the skull in order to obtain reproducible cell counts. Moreover, to maximize the recovery of bone marrow cells, the skull pieces can undergo 2-3 rounds of centrifugation with mixing between each centrifugation.
Both protocols have limitations. First, the use of 30% density gradient solution centrifugation may not effectively remove the remaining red blood cells in the brain, which may occur due to incomplete perfusion. Moreover, if myelin structure is modulated under certain conditions, a 30% density gradient solution may not be an optimal concentration for their removal. In this case, another working concentration of density gradient solution needs to be established. Second, although flow cytometry analysis indicates good cell viability of the brain immune cells using the described protocol, it still needs to be evaluated for other applications, such as transcriptomic analysis in which actinomycin D may be used. Third, the calvaria is used to prepare skull bone marrow. However, it must be noted that immune cells exist in many other parts of the skull. Lastly, the meninges, especially the dura mater, harbor diverse immune cells and play roles in brain homeostasis and dysfunction20. For immune analysis of the meninges, a protocol published elsewhere may be adopted17,21,22,23.
In summary, two robust protocols for isolating immune cells from both the mouse brain and skull are described. These protocols are expected to be useful for obtaining a large quantity of high-quality cells, suitable for various downstream applications.

The brain, the central hub of the nervous system, is protected by the skull. Beneath the skull are three layers of connective tissue known as the meninges - the dura mater, arachnoid mater, and pia mater. Cerebrospinal fluid (CSF) circulates in the subarachnoid space between the arachnoid mater and pia mater, cushioning the brain and also removing waste via the glymphatic system1,2. Together, this unique architecture provides a secure and supportive environment that maintains the stability of the brain and shields it from potential injury.
The brain has long been considered immune-privileged. However, this notion has been partially abandoned as mounting evidence indicates that, in addition to resident microglia in the parenchyma, the borders of the brain, including the choroid plexus and meninges, host a diverse array of immune cells3. These cells play critical roles in maintaining homeostasis, surveilling brain health, and initiating the immune response to brain injury. Notably, recent findings indicate that the skull is involved in meninges immunity, and may contribute to the immune response in the brain after injury. In 2018, Herisson et al. made a seminal discovery of direct vascular channels that link the skull bone marrow to the meninges, thereby establishing an anatomical route for leukocyte migration4,5. Later, Cugurra et al. demonstrated that many myeloid cells (e.g., monocytes and neutrophils) and B cells in the meninges do not originate from the blood6. Using techniques such as calvaria bone-flap transplantation and selective irradiation regimens, the authors provided compelling evidence that the skull bone marrow serves as a local source for myeloid cells in the meninges as well as CNS parenchyma after CNS injury6. Further, another study proposed that meningeal B cells are constantly supplied by the skull bone marrow7. More recently, a novel structure, termed the arachnoid cuff exit (ACE), has been identified as a direct gateway between the dura mater and the brain for immune cell trafficking8.
These exciting findings have important implications for the origin of infiltrating immune cells into the injured brain (e.g., after ischemic stroke). A large body of evidence has indicated that after stroke, many immune cells infiltrate the brain, contributing to both acute brain damage and chronic brain recovery. The conventional notion&#160;is that these cells are circulating leukocytes in the blood that infiltrate the brain, which is largely facilitated by stroke-induced blood-brain barrier damage. However, this notion has been challenged. In one study, immune cells in the skull and the tibia of mice were labeled differently, and at 6 h after stroke, a significantly greater decrease in neutrophils and monocytes was found in the skull vs. the tibia, and more skull-derived neutrophils were present in the ischemic brain. These data suggest that in the acute stroke phase, neutrophils in the ischemic brain primarily originate from the skull bone marrow4. Interestingly, CSF may guide this migration. Indeed, two recent reports demonstrated that CSF can directly relay signaling cues from the brain into the skull bone marrow via skull channels, and instruct cell migration and hematopoiesis in the skull bone marrow after CNS injury9,10.
In light of these recent findings, it has become important to analyze changes in immune cells in both the brain and the skull bone marrow, when studying the immune response to brain disorders. In such investigations, sufficient numbers of high-quality immune cells are needed for downstream analyses such as cell sorting, flow cytometry analysis, and single-cell RNA sequencing (scRNA-seq). Here, the overall goal is to present two optimized procedures for preparing single-cell suspensions from brain tissue and skull bone marrow. It is important to note that the calvaria (frontal bone, occipital bone, and parietal bones) of the skull are typically used to extract bone marrow, and this bone marrow is specifically referred to as skull bone marrow throughout this study.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.5 mL microcentrifuge tubes</td>
			<td>VWR</td>
			<td>76332-066</td>
			<td></td>
		</tr>
		<tr>
			<td>1.5 mL microcentrifuge tubes</td>
			<td>VWR</td>
			<td>76332-068</td>
			<td></td>
		</tr>
		<tr>
			<td>15 mL conical tubes</td>
			<td>Thermo Fisher Scientific</td>
			<td>339651</td>
			<td></td>
		</tr>
		<tr>
			<td>18 G x 1 in BD PrecisionGlide Needle</td>
			<td>BD Biosciences</td>
			<td>305195</td>
			<td></td>
		</tr>
		<tr>
			<td>1x HBSS</td>
			<td>Gibco</td>
			<td>14175-095</td>
			<td></td>
		</tr>
		<tr>
			<td>50 mL conical tubes</td>
			<td>Thermo Fisher Scientific</td>
			<td>339653</td>
			<td></td>
		</tr>
		<tr>
			<td>96-well V-bottom microplate&#160;</td>
			<td>SARSTEDT</td>
			<td>82.1583</td>
			<td></td>
		</tr>
		<tr>
			<td>AURORA&#160; flow cytometer</td>
			<td>Cytek bioscience</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Fisher</td>
			<td>BP9706-100</td>
			<td></td>
		</tr>
		<tr>
			<td>CD11b-AF594</td>
			<td>BioLegend</td>
			<td>101254</td>
			<td>1:500 dilution</td>
		</tr>
		<tr>
			<td>CD19-BV785</td>
			<td>BioLegend</td>
			<td>115543</td>
			<td>1:500 dilution</td>
		</tr>
		<tr>
			<td>CD19-FITC</td>
			<td>BioLegend</td>
			<td>115506</td>
			<td>1:500 dilution</td>
		</tr>
		<tr>
			<td>CD3-APC</td>
			<td>BioLegend</td>
			<td>100312</td>
			<td>1:500 dilution</td>
		</tr>
		<tr>
			<td>CD3-PE</td>
			<td>BioLegend</td>
			<td>100206</td>
			<td>1:500 dilution</td>
		</tr>
		<tr>
			<td>CD45-Alex 700</td>
			<td>BioLegend</td>
			<td>103128</td>
			<td>1:500 dilution</td>
		</tr>
		<tr>
			<td>CD45-BV421</td>
			<td>Biolegend</td>
			<td>103133</td>
			<td>1:500 dilution</td>
		</tr>
		<tr>
			<td>Cell Strainer 70 um</td>
			<td>Avantor</td>
			<td>732-2758</td>
			<td></td>
		</tr>
		<tr>
			<td>Dressing Forceps&#160;</td>
			<td>V. Mueller</td>
			<td>NL1410</td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA</td>
			<td>Invitrogen</td>
			<td>15575-038</td>
			<td></td>
		</tr>
		<tr>
			<td>Fc Block</td>
			<td>Biolegend</td>
			<td>101320</td>
			<td>1:100 dilution</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>Roboz</td>
			<td>RS-5047</td>
			<td></td>
		</tr>
		<tr>
			<td>LIVE/DEAD Fixable Blue Dead Cell Stain Kit</td>
			<td>Thermo Fisher Scientific</td>
			<td>N7167</td>
			<td>1:500 dilution</td>
		</tr>
		<tr>
			<td>Ly6G-BV421</td>
			<td>BioLegend</td>
			<td>127628</td>
			<td>1:500 dilution</td>
		</tr>
		<tr>
			<td>Ly6G-PerCp-cy5.5</td>
			<td>BioLegend</td>
			<td>127615</td>
			<td>1:500 dilution</td>
		</tr>
		<tr>
			<td>NK1.1-APC-cy7</td>
			<td>BioLegend</td>
			<td>108723</td>
			<td>1:500 dilution</td>
		</tr>
		<tr>
			<td>Percoll (density gradient medium)</td>
			<td>Cytiva</td>
			<td>17089101</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate buffer saline (10x)</td>
			<td>Gibco</td>
			<td>70011-044</td>
			<td></td>
		</tr>
		<tr>
			<td>RBC Lysis Buffer (10x)</td>
			<td>BioLegend</td>
			<td>420302</td>
			<td></td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>SKLAR</td>
			<td>64-1250</td>
			<td></td>
		</tr>
		<tr>
			<td>WHEATON Dounce Tissue, 15 mL Size</td>
			<td>DWK Life Sciences</td>
			<td>357544</td>
			<td></td>
		</tr>
	</tbody>
</table>

isolating immune cells from mouse brain and skull

Mounting evidence indicates that the immune response triggered by brain disorders (e.g., brain ischemia and autoimmune encephalomyelitis) occurs not only in the brain, but also in the skull. A key step toward analyzing changes in immune cell populations in both the brain and skull bone marrow after brain damage (e.g., stroke) is to obtain sufficient numbers of high-quality immune cells for downstream analyses. Here, two optimized protocols are provided for isolating immune cells from the brain and skull bone marrow. The advantages of both protocols are reflected in their simplicity, speed, and efficacy in yielding a large quantity of viable immune cells. These cells may be suitable for a range of downstream applications, such as cell sorting, flow cytometry, and transcriptomic analysis. To demonstrate the effectiveness of the protocols, immunophenotyping experiments were performed on stroke brains and normal brain skull bone marrow using flow cytometry analysis, and the results aligned with findings from published studies.

To prepare immune cells from the mouse brain tissue, the protocol generally yields cells with high viability (84.1%&#160;&#177; 2.3% [mean&#160;&#177; SD]). Approximately 70%-80% of these cells are CD45 positive. In the normal mouse brain, nearly all CD45+ cells are microglia (CD45LowCD11b+), as expected. This protocol has been used in the laboratory for various applications, including flow cytometry analysis, fluorescence-activated cell sorting (FACS), and scRNA-seq analysis. As an example, a flow cytometry analysis was conducted in a stroke model (Figure 3). Mice were subjected to a transient stroke model, 30 min filament middle cerebral artery occlusion (MCAO)14. On day 3 after stroke, cells were prepared from mouse brains and stained with common immune cell surface markers. The gating strategy is depicted in Figure 3A. Consistent with prior research13,15, a marked increase in CD45High cells in the ipsilateral hemisphere was observed (Figure 3B,C), indicating infiltration of the immune cell into the brain.
For the skull bone marrow protocol, a substantial yield of skull bone marrow cells (around 2 x 106 cells) was consistently achieved with excellent viability (93.8% &#177;&#160;1.8% [mean&#160;&#177; SD]). As expected, most of these cells are CD45 positive. To illustrate the utility of this protocol, a comparative study was performed to characterize the immune cell composition in both the femur and skull bone marrow of naive mice (Figure 4). As depicted in the representative plots, the immune cell composition was similar between the femur and skull bone marrow (Figure 4A,B). Interestingly, the frequency of neutrophils appears to be higher in the femur than in the skull bone (Figure 4C). However, this finding requires further validation, as the sample size was small for this pilot study.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66861/66861fig01.jpg" /> 
Figure 1: Overview of the brain cell isolation protocol. <a href="https://www.jove.com/files/ftp_upload/66861/66861fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66861/66861fig02.jpg" /> 
Figure 2: The skull bone marrow cell isolation protocol. (A) Overview of the procedure. (B) The recommended pattern for skull cutting. <a href="https://www.jove.com/files/ftp_upload/66861/66861fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66861/66861fig03.jpg" /> 
Figure 3: Representative flow cytometry analysis of the immune cells in the stroke brain. (A) An example of the gating strategy. The plots depict the following immune cell populations: microglia (CD45LowCD11b+), NK cells (CD45HighNK1.1+), neutrophils (CD45HighLy6G+), T cells (CD45HighCD11b-CD3+), and B cells (CD45HighCD11b-CD19+). This example represents an analysis of brain immune cells in the ipsilateral hemisphere on day 3 after stroke. (B,C) Infiltrating immune cells into the stroke brain. Young male mice were subjected to transient middle cerebral artery occlusion (MCAO). On day 3, brains were collected and split into left (contralateral) and right (ipsilateral) hemispheres. Brain cells were prepared according to the described protocol and then subjected to flow cytometry analysis. Representative plots illustrate increased infiltrating immune cells into the ipsilateral hemisphere (CD45High cells; B). Quantitative data are shown in the bar graphs (C). The cell numbers were calculated based on the cell count and volume recorded in the flow cytometry data. The data are presented as mean &#177; SEM. *p &#60; 0.05. <a href="https://www.jove.com/files/ftp_upload/66861/66861fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66861/66861fig04.jpg" /> 
Figure 4: Representative flow cytometry analysis of the skull bone marrow cells. For comparison, analysis of bone marrow cells from the femur was included. The plots depict the following immune cell populations: NK cells (CD45+NK1.1+), neutrophils (CD45+CD11b+Ly6G+), T cells (CD45+CD11b-CD3+), and B cells (CD45+CD11b-CD19+). (A) Representative plots for femur bone marrow. (B) Representative plots for skull bone marrow. (C) Comparison of the immune cell composition between femur and skull bone marrow. The data are presented as mean &#177; SEM. *p &#60; 0.05. <a href="https://www.jove.com/files/ftp_upload/66861/66861fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Author Spotlight: Investigating the Complex Immune Response Following Brain Ischemia

To investigate the immune response to brain disorders, one common approach is to analyze changes in immune cells. Here, two simple and effective protocols are provided for isolating immune cells from murine brain tissue and skull bone marrow.

Isolating Immune Cells from Mouse Brain and Skull

Mounting evidence indicates that the immune response triggered by brain disorders (e.g., brain ischemia and autoimmune encephalomyelitis) occurs not only ...

isolating-immune-cells-from-mouse-brain-and-skull

Universidad San Sebastian

Research

JoVE Journal

Neuroscience

2.2K Views.  Duke University Medical Center. To investigate the immune response to brain disorders, one common approach is to analyze changes in immune cells. Here, two simple and effective protocols are provided for isolating immune cells from murine brain tissue and skull bone marrow.

Video: Author Spotlight: Investigating the Complex Immune Response Following Brain Ischemia

Isolating LacZ-expressing Cells from Mouse Inner Ear Tissues using Flow Cytometry

Isolation of specific cell types allows one to analyze rare cell populations such as stem/progenitor cells. Such an approach to studying inner ear tissues presents a unique challenge because of the paucity of cells of interest and few transgenic reporter mouse models. Here, we describe a protocol using fluorescence-conjugated probes to selectively label LacZ-positive cells from the neonatal cochleae.
The most common underlying pathology of sensorineural hearing loss is the irreversible damage and loss of cochlear sensory hair cells, which are required to transduce sound waves to neural impulses. Recent evidence suggests that the murine auditory and vestibular organs harbor stem/progenitor cells that may have regenerative potential1,2. These findings warrant further investigation, including identifying specific cell types with stem/progenitor cell characteristics. The Wnt signaling pathway has been demonstrated to play a critical role in maintaining stem/progenitor cell populations in several organ systems3-7. We have recently identified Wnt-responsive Axin2-expressing cells in the neonatal cochlea, but their function is largely unknown8.
To better understand the behavior of these Wnt-responsive cells in vitro, we have developed a method of isolating Axin2-expressing cells from cochleae of Axin2-LacZ reporter mice9. Using flow cytometry to isolate Axin2-LacZ positive cells from the neonatal cochleae, we could in turn execute a variety of experiments on live cells to interrogate their behavior as stem/progenitor cells. Here, we describe in detail the steps for the microdissection of neonatal cochlea, dissociation of these tissues, labeling of the LacZ-positive cells using a fluorogenic substrate, and cell sorting. Techniques for dissociating cochleae into single cells and isolating cochlear cells via flow cytometry have been described2,10-12. We have made modifications to these techniques to establish a novel protocol to isolate LacZ-expressing cells from the neonatal cochlea.

Flow cytometry is a powerful tool allowing for the isolation and study of specific cell populations. This protocol describes steps for isolating LacZ-expressing cells from cochlear tissues from neonatal transgenic mice. Dissociated cochlear cells were labeled using fluorescent-conjugated substrates of &#946;-galactosidase prior to separation via flow cytometry.

Isolation of specific cell types allows one to analyze rare cell populations such as stem/progenitor cells. Such an approach to studying inner ear tissues ...

A Polished and Reinforced Thinned-skull Window for Long-term Imaging of the Mouse Brain

In vivo imaging of cortical function requires optical access to the brain without disruption of the intracranial environment. We present a method to form a polished and reinforced thinned skull (PoRTS) window in the mouse skull that spans several millimeters in diameter and is stable for months. The skull is thinned to 10 to 15 &mu;m in thickness with a hand held drill to achieve optical clarity, and is then overlaid with cyanoacrylate glue and a cover glass to: 1) provide rigidity, 2) inhibit bone regrowth and 3) reduce light scattering from irregularities on the bone surface. Since the skull is not breached, any inflammation that could affect the process being studied is greatly reduced. Imaging depths of up to 250 &mu;m below the cortical surface can be achieved using two-photon laser scanning microscopy. This window is well suited to study cerebral blood flow and cellular function in both anesthetized and awake preparations. It further offers the opportunity to manipulate cell activity using optogenetics or to disrupt blood flow in targeted vessels by irradiation of circulating photosensitizers.

We present a method to form an imaging window in the mouse skull that spans millimeters and is stable for months without inflammation of the brain. This method is well suited for longitudinal studies of blood flow, cellular dynamics, and cell/vascular structure using two-photon microscopy.

In vivo imaging of cortical function requires optical access to the brain without disruption of the intracranial environment. We present a method to form ...

Mitochondria-associated ER Membranes (MAMs) and Glycosphingolipid Enriched Microdomains (GEMs): Isolation from Mouse Brain

Intracellular organelles are highly dynamic structures with varying shape and composition, which are subjected to cell-specific intrinsic and extrinsic cues. Their membranes are often juxtaposed at defined contact sites, which become hubs for the exchange of signaling molecules and membrane components1,2,3,4. The inter-organellar membrane microdomains that are formed between the endoplasmic reticulum (ER) and the mitochondria at the opening of the IP3-sensitive Ca2+ channel are known as the mitochondria associated-ER membranes or MAMs4,5,6. The protein/lipid composition and biochemical properties of these membrane contact sites have been extensively studied particularly in relation to their role in regulating intracellular Ca2+ 4,5,6. The ER serves as the primary store of intracellular Ca2+, and in this capacity regulates a myriad of cellular processes downstream of Ca2+ signaling, including post-translational protein folding and protein maturation7. Mitochondria, on the other hand, maintain Ca2+ homeostasis, by buffering cytosolic Ca2+ concentration thereby preventing the initiation of apoptotic pathways downstream of Ca2+ unbalance4,8. The dynamic nature of the MAMs makes them ideal sites to dissect basic cellular mechanisms, including Ca2+ signaling and regulation of mitochondrial Ca2+ concentration, lipid biosynthesis and transport, energy metabolism and cell survival 4,9,10,11,12. Several protocols have been described for the purification of these microdomains from liver tissue and cultured cells13,14. 
	Taking previously published methods into account, we have adapted a protocol for the isolation of mitochondria and MAMs from the adult mouse brain. To this procedure we have added an extra purification step, namely a Triton X100 extraction, which enables the isolation of the glycosphingolipid enriched microdomain (GEM) fraction of the MAMs. These GEM preparations share several protein components with caveolae and lipid rafts, derived from the plasma membrane or other intracellular membranes, and are proposed to function as gathering points for the clustering of receptor proteins and for protein&#x2013;protein interactions4,15.

Mitochondria-associated ER Membranes MAMs and Glycosphingolipid Enriched Microdomains GEMs: Isolation from Mouse Brain

This procedure illustrates how to isolate from the adult mouse brain the mitochondria-associated ER membranes or MAMs and the glycosphingolipid-enriched microdomain fractions from MAMs and mitochondrial preparations.

Intracellular organelles are highly dynamic structures with varying shape and composition, which are subjected to cell-specific intrinsic and extrinsic ...

Isolation and Culture of Endothelial Cells from the Embryonic Forebrain

Embryonic brain endothelial cells can serve as an important tool in the study of angiogenesis and neurovascular development and interactions. The two vascular networks of the embryonic forebrain, pial and periventricular, are spatially distinctive and have different origins and growth patterns. Endothelial cells from the pial and periventricular vascular networks have unique gene expression profiles and functions. Here we present a step-by-step protocol for isolation, culture, and verification of pure populations of endothelial cells from the periventricular vascular network (PVECs) of the embryonic forebrain (telencephalon). In this approach, telencephalon devoid of pial membrane obtained from embryonic day 15 mice is minced, digested with collagenase/dispase, and dispersed mechanically into a single cell suspension. PVECs are purified from cell suspension using positive selection with anti-CD-31/PECAM-1 antibody conjugated to MicroBeads using a strong magnetic separation method. Purified cells are cultured on collagen 1&#160;coated culture dishes in endothelial cell culture medium until they become confluent and further subcultured. PVECs obtained with this protocol exhibit cobblestone and spindle shaped phenotypes, as visualized by phase-contrast light microscopy and fluorescence microscopy. Purity of PVEC cultures was established with endothelial cell markers. In our hands, this method reliably and consistently yields pure populations of PVECs. This protocol will benefit studies aimed at gaining mechanistic insights into forebrain angiogenesis, understanding PVEC interactions, and cross-talks with neuronal cell types and holds tremendous potential for therapeutic angiogenesis.

This video demonstrates an easy and reliable strategy for preparation of pure cultures of endothelial cells from the embryonic forebrain within 10-12 days and will be useful for research focused on many aspects of cerebral angiogenesis.

Embryonic brain endothelial cells can serve as an important tool in the study of angiogenesis and neurovascular development and interactions. The two ...

Exposure of the Pig CNS for Histological Analysis: A Manual for Decapitation, Skull Opening, and Brain Removal

Pigs have become increasingly popular in large-animal translational neuroscience research as an economically and ethically feasible substitute to non-human primates. The large brain size of the pig allows the use of conventional clinical brain imagers and the direct use and testing of neurosurgical procedures and equipment from the human clinic. Further macroscopic and histological analysis, however, requires postmortem exposure of the pig central nervous system (CNS) and subsequent brain removal. This is not an easy task, as the pig CNS is encapsulated by a thick, bony skull and spinal column. The goal of this paper and instructional video is to describe how to expose and remove the postmortem pig brain and the pituitary gland in an intact state, suitable for subsequent macroscopic and histological analysis.

The goal of this paper and instructional video is to describe how to expose and remove the postmortem pig brain and pituitary gland in an intact state, suitable for subsequent macroscopic and histological analysis.

Pigs have become increasingly popular in large-animal translational neuroscience research as an economically and ethically feasible substitute to ...

Analysis of Immune Cells in Single Sciatic Nerves and Dorsal Root Ganglion from a Single Mouse Using Flow Cytometry

Nerve-resident immune cells in the peripheral nervous system (PNS) are essential to maintaining neuronal integrity in a healthy nerve. The immune cells of the PNS are affected by injury and disease, affecting the nerve function and the capacity for regeneration. Neuronal immune cells are commonly analyzed by immunofluorescence (IF). While IF is essential for determining the location of the immune cells in the nerve, IF is only semi-quantitative and the method is limited to the number of markers that can be analyzed simultaneously and the degree of surface expression. In this study, flow cytometry was used for quantitative analysis of leukocyte infiltration into sciatic nerves or dorsal root ganglions (DRGs) of individual mice. Single cell analysis was performed using DAPI and several proteins were analyzed simultaneously for either surface or intracellular expression. Both sciatic nerves from one mouse that were treated according to this protocol generated &#8805; 30,000 single nucleated events. The proportion of leukocytes in the sciatic nerves, determined by expression of CD45, was approximately 5% of total cell content in the sciatic nerve and approximately 5-10% in the DRG. Although this protocol focuses primarily on the immune cell population within the PNS, the flexibility of flow cytometry to measure a number of markers simultaneously means that the other cells populations present within the nerve, such as Schwann cells, pericytes, fibroblasts, and endothelial cells, can also be analyzed using this method. This method therefore provides a new means for studying systemic effects on the PNS, such as neurotoxicology and genetic models of neuropathy or in chronic diseases, such as diabetes.

Quantitative analysis of cell content within the murine sciatic nerve is difficult due to the scarcity of the tissue. This protocol describes a method for tissue digestion and preparation that provides sufficient cells for flow cytometry analysis of immune cell populations from nerves of individual mice.

Nerve-resident immune cells in the peripheral nervous system (PNS) are essential to maintaining neuronal integrity in a healthy nerve. The immune cells of ...

Neuronal Differentiation from Mouse Embryonic Stem Cells In vitro

The neural differentiation of mouse embryonic stem cells (mESCs) is a potential tool for elucidating the key mechanisms involved in neurogenesis and potentially aid in regenerative medicine. Here, we established an efficient and low cost method for neuronal differentiation from mESCs in vitro, using the strategy of combinatorial screening. Under the conditions defined here, the 2-day embryoid body formation + 6-day retinoic acid induction protocol permits fast and efficient differentiation from mESCs into neural precursor cells (NPCs), as seen by the formation of well-stacked and neurite-like A2lox and 129 derivatives that are Nestin positive. The healthy state of embryoid bodies and the timepoint at which retinoic acid (RA) is applied, as well as the RA concentrations, are critical in the process. In the subsequent differentiation from NPCs into neurons, N2B27 medium II (supplemented by Neurobasal medium) could better support the long term maintenance and maturation of neuronal cells. The presented method is highly efficiency, low cost and easy to operate, and can be a powerful tool for neurobiology and developmental biology research.

Here, we established a low cost and easy to operate method that directs fast and efficient differentiation from embryonic stem cells into neurons. This method is suitable for popularization among laboratories and can be a useful tool for neurological research.

The neural differentiation of mouse embryonic stem cells (mESCs) is a potential tool for elucidating the key mechanisms involved in neurogenesis and ...

Derivation, Expansion, Cryopreservation and Characterization of Brain Microvascular Endothelial Cells from Human Induced Pluripotent Stem Cells

Brain microvascular endothelial cells (BMECs) can be differentiated from human induced pluripotent stem cells (iPSCs) to develop ex vivo cellular models for studying blood-brain barrier (BBB) function. This modified protocol provides detailed steps to derive, expand, and cryopreserve BMECs from human iPSCs using a different donor and reagents than those reported in previous protocols. iPSCs are treated with essential 6 medium for 4 days, followed by 2 days of human endothelial serum-free culture medium supplemented with basic fibroblast growth factor, retinoic acid, and B27 supplement. At day 6, cells are sub-cultured onto a collagen/fibronectin matrix for 2 days. Immunocytochemistry is performed at day 8 for BMEC marker analysis using CLDN5, OCLN, TJP1, PECAM1, and SLC2A1. Western blotting is performed to confirm BMEC marker expression, and absence of SOX17, an endodermal marker. Angiogenic potential is demonstrated with a sprouting assay. Trans-endothelial electrical resistance (TEER) is measured using chopstick electrodes and voltohmmeter starting at day 7. Efflux transporter activity for ATP binding cassette subfamily B member 1 and ATP binding cassette subfamily C member 1 is measured using a multi-plate reader at day 8. Successful derivation of BMECs is confirmed by the presence of relevant cell markers, low levels of SOX17, angiogenic potential, transporter activity, and TEER values ~2000 &#937; x cm2. BMECs are expanded until day 10 before passaging onto freshly coated collagen/fibronectin plates or cryopreserved. This protocol demonstrates that iPSC-derived BMECs can be expanded and passaged at least once. However, lower TEER values and poorer localization of BMEC markers was observed after cryopreservation. BMECs can be utilized in co-culture experiments with other cell types (neurons, glia, pericytes), in three-dimensional brain models (organ-chip and hydrogel), for vascularization of brain organoids, and for studying BBB dysfunction in neuropsychiatric disorders.

This protocol details an adapted method to derive, expand, and cryopreserve brain microvascular endothelial cells obtained by differentiating human induced pluripotent stem cells, and to study blood brain barrier properties in an ex vivo model.

Brain microvascular endothelial cells (BMECs) can be differentiated from human induced pluripotent stem cells (iPSCs) to develop ex vivo cellular models ...

In vivo Imaging of Fully Active Brain Tissue in Awake Zebrafish Larvae and Juveniles by Skull and Skin Removal

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at cellular and subcellular resolution. Advances in molecular and imaging technologies have allowed us to gain numerous detailed insights into cellular and molecular mechanisms of brain development in the transparent zebrafish embryo. Recently, processes of refinement of neuronal connectivity that occur at later larval stages several weeks after fertilization, which are for example control of social behavior, decision making or motivation-driven behavior, have moved into focus of research. At these stages, pigmentation of the zebrafish skin interferes with light penetration into brain tissue, and solutions for embryonic stages, e.g., pharmacological inhibition of pigmentation, are not feasible anymore.
Therefore, a minimally invasive surgical solution for microscopy access to the brain of awake zebrafish is provided that is derived from electrophysiological&#160;approaches. In teleosts, skin and soft skull cartilage can be carefully removed by micro-peeling these layers, exposing underlying neurons and axonal tracts without damage. This allows for recording neuronal morphology, including synaptic structures and their molecular contents, and the observation of physiological changes such as Ca2+ transients or intracellular transport events. In addition, interrogation of these processes by means of pharmacological inhibition or optogenetic manipulation is feasible. This brain exposure approach provides information about structural and physiological changes in neurons as well as the correlation and interdependence of these events in live brain tissue in the range of minutes or hours. The technique is suitable for in vivo brain imaging of zebrafish larvae up to 30 days post fertilization, the latest developmental stage tested so far. It, thus, provides access to such important questions as synaptic refinement and scaling, axonal and dendritic transport, synaptic targeting of cytoskeletal cargo or local activity-dependent expression. Therefore, a broad use for this mounting and imaging approach can be anticipated.

Here we present a method to image the zebrafish embryonic brain in vivo upto larval and juvenile stages. This microinvasive procedure, adapted from electrophysiological approaches, provides access to cellular and subcellular details of mature neuron and can be combined with optogenetics and neuropharmacological studies for characterizing brain function and drug intervention.

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at ...

Micro-Drive Implant for Recording Daily Neural Activity in Juvenile Mice

A Lightweight Drive Implant for Chronic Tetrode Recordings in Juvenile Mice

In vivo electrophysiology provides unparalleled insight into the sub-second-level circuit dynamics of the intact brain and represents a method of particular importance for studying mouse models of human neuropsychiatric disorders. However, such methods often require large cranial implants, which cannot be used in mice at early developmental time points. As such, virtually no studies of in vivo physiology have been performed in freely behaving infant or juvenile mice, despite the fact that a better understanding of neurological development in this critical window would likely provide unique insights into age-dependent developmental disorders such as autism or schizophrenia. Here, a micro-drive design, surgical implantation procedure, and post-surgery recovery strategy are described that allow for chronic field and single-unit recordings from multiple brain regions simultaneously in mice as they age from postnatal day 20 (p20) to postnatal day 60 (p60) and beyond, a time window roughly corresponding to the human ages of 2 years old through to adulthood. The number of recording electrodes and final recording sites can be easily modified and expanded, thus allowing flexible experimental control of the in vivo monitoring of behavior- or disease-relevant brain regions across development.

Author Spotlight: A Lightweight Drive Implant for Chronic Tetrode Recordings in Juvenile Mice  

Here, we describe a micro-drive design, surgical implantation procedure, and post-surgery recovery strategy that allow for chronic field and single-unit recordings from multiple brain regions simultaneously in juvenile and adolescent mice across a critical developmental window from postnatal day 20 (p20) to postnatal day 60 (p60) and beyond.