Isolating Central Nervous System Tissues and Associated Meninges for the Downstream Analysis of Immune cells

Krista D. DiSano; Michael  R. Linzey; Nora  C. Welsh; Joshua  S. Meier; Andrew  R. Pachner; Francesca Gilli

doi:10.3791/61166

Immunology and Infection

Isolating Central Nervous System Tissues and Associated Meninges for the Downstream Analysis of Immune cells

Published: May 19, 2020 doi: 10.3791/61166

Krista D. DiSano¹, Michael R. Linzey^1,2, Nora C. Welsh^1,2, Joshua S. Meier¹, Andrew R. Pachner¹, Francesca Gilli¹

¹Department of Neurology, Geisel School of Medicine, Dartmouth-Hitchcock Medical Center, ²Program in Experimental and Molecular Medicine, Dartmouth College

Summary

This paper presents two optimized protocols for examining resident and peripherally derived immune cells within the central nervous system, including the brain, spinal cord, and meninges. Each of these protocols helps to ascertain the function and composition of the cells occupying these compartments under steady state and inflammatory conditions.

Abstract

The central nervous system (CNS) is comprised of the brain and spinal cord and is enveloped by the meninges, membranous layers serving as a barrier between the periphery and the CNS. The CNS is an immunologically specialized site, and in steady state conditions, immune privilege is most evident in the CNS parenchyma. In contrast, the meninges harbor a diverse array of resident cells, including innate and adaptive immune cells. During inflammatory conditions triggered by CNS injury, autoimmunity, infection, or even neurodegeneration, peripherally derived immune cells may enter the parenchyma and take up residence within the meninges. These cells are thought to perform both beneficial and detrimental actions during CNS disease pathogenesis. Despite this knowledge, the meninges are often overlooked when analyzing the CNS compartment, because conventional CNS tissue extraction methods omit the meningeal layers. This protocol presents two distinct methods for the rapid isolation of murine CNS tissues (i.e., brain, spinal cord, and meninges) that are suitable for downstream analysis via single-cell techniques, immunohistochemistry, and in situ hybridization methods. The described methods provide a comprehensive analysis of CNS tissues, ideal for assessing the phenotype, function, and localization of cells occupying the CNS compartment under homeostatic conditions and during disease pathogenesis.

Introduction

The central nervous system (CNS) is an immunologically specialized site. The CNS parenchyma, excluding the CSF space, the meninges, and the vasculature, is classically viewed as an immune-privileged site¹^,²^,³^,⁴^,⁵ and is relatively devoid of immune cells during homeostatic conditions²^,⁶^,⁷. In contrast, the meninges, comprised of the dura, arachnoid, and pia layers, are crucial components of the CNS compartment, actively participating in homeostatic immune surveillance and inflammatory processes during disease pathogenesis³^,⁶^,⁷^,⁸. During steady state conditions, the meninges support numerous immune sentinel cells, including innate lymphoid cells (ILC), macrophages, dendritic cells (DC), mast cells, T cells, and to a lesser extent, B cells⁹^,¹⁰^,¹¹.

The meninges are highly vascularized structures and contain lymphatic vessels that provide a lymphatic connection between the CNS and its periphery⁸^,¹²^,¹³^,¹⁴. In inflammatory conditions induced by CNS injury, infections, autoimmunity, or even neurodegeneration, peripherally derived immune cells infiltrate the parenchyma and alter the immune landscape within the meninges. Following cell infiltration, the meninges may represent a functional niche for peripherally derived immune cells, promoting immune cell aggregation, local immune cell activation, and long-term survival in the CNS compartment. Prominent meningeal inflammation is observed in multiple diseases affecting the CNS, including multiple sclerosis (MS)¹⁵^,¹⁶^,¹⁷^,¹⁸^,¹⁹, stroke²⁰^,²¹, sterile injury²²^,²³ (i.e., spinal cord injury and traumatic brain injury), migraines²⁴, and microbial infection²⁵^,²⁶^,²⁷^,²⁸^,²⁹. Thus, the characterization of resident cells and peripherally derived immune cells in the meningeal compartment is essential for understanding the role of these cells during steady state conditions and disease pathogenesis.

The extraction of the brain, spinal cord, and meninges from the cranium and vertebral bodies is technically challenging and time-consuming. There are currently no techniques available for the rapid extraction of the brain with all three meningeal layers intact. While laminectomy yields excellent spinal cord tissue morphology and preserves the meningeal layers, it is both extremely time-consuming and complicated³⁰^,³¹. Conversely, more conventional extraction methods such as the removal of the brain from the cranium and the hydraulic extrusion of the spinal cord facilitate the quick extraction of the CNS tissue, but both the arachnoid and dural meninges are lost with these techniques³⁰^,³¹. The omission of dura and arachnoid layers during conventional isolation of brain and spinal cord tissues results in an incomplete analysis of the cells within the CNS compartment. Thus, the identification of new techniques focused on the quick extraction of CNS tissues with intact meninges is crucial for the optimal analysis of the CNS compartment.

This manuscript presents two methods for the rapid extraction of the brain, spinal cord, and meninges from mice, facilitating the downstream analysis of resident cells and peripherally derived immune cells in the CNS parenchyma and meninges. These optimized protocols focus on 1) isolating single-cell suspensions for downstream analysis and 2) preparing tissue for histological processing. Obtaining single-cell suspensions from the brain, spinal cord tissue, and dural and arachnoid meninges³² allows for the simultaneous analysis of cells residing in both the parenchymal and meningeal compartments. Single-cell suspensions can be used in different applications, including cell culture assays to perform in vitro stimulation³³, enzyme-linked immunospot (ELISpot)²⁸^,³⁴^,³⁵, flow cytometry³⁶^,³³, and single-cell³⁷ or bulk transcriptomics. Additionally, the optimized protocol for decalcification of whole brains and spinal cords with intact skulls or vertebral columns, respectively, allows for the gentle decalcification of the surrounding bone, leaving the meninges intact and preserving the tissue morphology. This method allows for the selective identification of proteins or RNA using immunohistochemistry (IHC) or in situ hybridization (ISH) techniques within both the parenchymal and meningeal spaces. The characterization of the phenotype, activation state, and localization of resident cells and peripherally derived immune cells within the CNS may provide information essential to understanding how individual cell types in the CNS compartment contribute to homeostasis and disease pathogenesis.

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Protocol

All animal work utilizes protocols reviewed and approved by the Institutional Animal Care and Use Committee (IACUC) at Geisel School of Medicine at Dartmouth.

1. Processing brain and spinal cord samples for decalcification

Isolating brain and spinal cord samples
1. Euthanize the mouse via CO₂ inhalation. Ensure that the CO₂ flow rate displaces 10%–30% of the cage volume per minute.
2. Using forceps, lift the xiphoid process, and cut the abdominal wall laterally just below the rib cage with scissors, pulling up to avoid cutting underlying blood vessels or organs. Cut through the diaphragm laterally.
3. Cut the rib cage along the lateral edges parallel to the lungs up to the collarbone. Using forceps, lift the sternum and clamp the sternum with a hemostat. Place the hemostat over the head to lift the rib cage away and expose the heart.
4. Using forceps, grasp the heart near its apex and make an incision in the right atrium of the heart to provide an outlet. Insert a 25 G needle with a 10 mL syringe attached to slowly administer 10 mL of ice-cold 1x phosphate buffered saline (PBS) into the left ventricle to transcardially perfuse the mouse.
  NOTE: Perfusion should occur over 4–5 min until the liver is cleared of blood. A total of 10 mL of 1x PBS is usually enough for perfusion, but more may be utilized if necessary. A clear liver usually indicates adequate perfusion.
5. Using sharp scissors, remove the head by decapitation (Figure 1; 1). Make a midline incision in the skin (Figure 1; 2) and flip the skin over the eyes to free the skull.
6. Cut at the nasal bone to release the mandible from the skull (Figure 1; 3). Remove the mandible, tongue, and eyes. Cut along the lateral aspects of the skull to release the tissue along the external auditory meatus (Figure 1; 4). Trim to remove all excess skin, muscle, and tissue overlaying the skull.
7. Separate the rib cage from the spinal column by cutting parallel to the spine with sharp scissors (Figure 1; 5 and 6). Make a small cut at the lower lumbar region to isolate the spinal column (Figure 1; 7). Trim and remove any remaining muscle along the spine to expose the vertebrae (Figure 1; 8).
  NOTE: Removing excess tissue from the spinal column and skull is necessary to obtain adequate penetration of fixative paraformaldehyde (PFA) and ethylenediaminetetraacetic acid (EDTA) decalcification buffer.
Post-fixation, decalcification, and cryopreservation
1. Using forceps, place the brain with the intact skull or spinal column in a 15 mL conical tube containing 10 mL of 4% PFA. Place the tubes at 4 ˚C for at least 48 h for adequate fixation.
  NOTE: To avoid overfixation of the tissue, do not exceed 72 h of fixation. Fixation times are extended for bone specimens before decalcification. Adequate fixation will protect tissue from the effects of the decalcification and ensure better tissue morphology.
2. Rinse the brain or spinal cord by removing the tissue from the 4% PFA with forceps and placing the tissue in a disposable 14 mL tube with 10 mL of 1x PBS for 5 min. Transfer the brain or spinal cord into a 50 mL conical tube with 10 mL of 10% EDTA (pH = 7.2–7.4).
  NOTE: Using a larger tube with 10 mL of EDTA gives the EDTA a greater contact area with the tissue and accelerates the decalcification process.
3. Check daily if the bone is soft and pliable: remove the tissue from the EDTA solution with forceps, place it on a Petri dish, and gently test the bone softness with a 25 G needle. If the needle easily penetrates the bone, the decalcification process is complete.
4. Remove the tissue from the EDTA solution and transfer the brain or spinal cord to a 14 mL disposable tube containing 10 mL of 1x PBS and wash for 10 min. Repeat the wash.
  NOTE: Decalcification usually takes 2–3 days. The solution should be changed every 2–3 days if the bone is not yet adequately decalcified. However, prolonged incubation in EDTA after the bone has been decalcified can damage the tissue morphology.
5. Prepare 10%, 20%, and 30% sucrose solutions by adding sucrose to 1x PBS. For example, for 10% sucrose, add 10 g of sucrose and bring the volume to 100 mL using sterile 1x PBS. Store the solution at 4 ˚C for up to 1 month.
  NOTE: Sucrose solutions are prone to microorganism growth, so samples should not be stored for prolonged periods of time in these solutions.
6. Remove the tissue from the 1x PBS, place it in 10 mL of 10% sucrose solution and store it at 4 ˚C. Let the tissue sit for 24 h or until it sinks to the bottom of the tube.
7. Repeat this process, moving the tissue to a 20% sucrose solution first and finally to a 30% sucrose solution. Allow the tissue to sink in 30% sucrose (at least 24 h) and proceed to tissue embedding.
Tissue embedding and freezing
1. Using forceps, remove the tissue from the 30% sucrose, place it on a Petri dish, and tilt the dish to get rid of any excess sucrose solution on the tissue. Using a scalpel, cut the tissue into desired segments.
2. Create a thin layer of optimal cutting temperature (OCT) compound at the bottom of the cryomold and place the tissue piece(s) in the mold. Cover the tissue completely with the OCT compound, ensuring no bubbles are present.
3. Flash freeze the blocks by hovering over liquid nitrogen³⁸ or setting the blocks on a 100% isopropanol/dry ice slurry³⁹ until the block is opaque. Wrap the cryomolds in aluminum foil and store the blocks at -80 ˚C for long-term storage. Move blocks to -20 ˚C before sectioning.
  NOTE: Care must be taken when sectioning and performing histology protocols on decalcified brains as the skull and meningeal layers may be lost if the sections are handled roughly.

2. Preparation of the meninges and CNS tissues for flow cytometry staining

Extracting the skull cap and brain
1. Using sharp scissors, remove the head by decapitation (Figure 2A; 1). Using scissors, make a midline incision in the skin (Figure 2A; 2) and flip the skin over the eyes to free the skull.
2. Place the scissors within the foramen magna and begin cutting the skull laterally along the cortices towards the olfactory bulb, keeping the incisions above the external auditory meatus and mandible (Figure 2A; 3). Perform the same cuts on the opposite side, with cuts meeting at the olfactory bulb to free the skull cap from the brain (Figure 2A; 3).
3. Using forceps, peel back the skull cap and place the skull cap into a 15 mL conical tube containing 5 mL of cold RPMI medium supplemented with 25 mM HEPES. Keep the tube on ice.
4. Using curved forceps, place the forceps below the base of the brain, and lift to free the brain from the skull cap. Place the brain into a 15 mL conical tube containing 5 mL of cold RPMI supplemented with 25 mM HEPES. Keep the tube on ice until processing.
Extracting the vertebral column and spinal cord tissue
1. Using forceps and sharp scissors, separate the rib cage from the spinal column by cutting parallel to the spine (Figure 2A; 4 and 5). Make a small cut at the lower lumbar region to isolate the vertebral column (Figure 2A; 6). Trim and remove any remaining muscle along the spine to expose the vertebrae (Figure 2A; 7).
2. Place the extra fine surgical scissors within the vertebral column and cut along the lateral edge of the column (Figure 2C). Cut the opposite lateral edge completely to divide the vertebral column into an anterior and posterior portion.
  NOTE: The spinal cord will remain attached to the vertebral column.
3. Using forceps, slowly and carefully peel away the spinal cord from the vertebral column and place the tissue in a 15 mL conical tube containing 5 mL of cold RPMI with 25 mM HEPES. Transfer the anterior and posterior portions of the spinal column to a 15 mL conical tube containing 5 mL of cold RPMI with 25 mM HEPES.
Removing the meninges to prepare single-cell suspensions
1. Using forceps, remove the skull cap from the RPMI media. Using sharp forceps (#7 forceps; Table of Materials), score around the outer edge of the skull cap (Figure 2B) and peel the meninges away from the edge of the skull cap, scraping to remove the dural and arachnoid meninges. Place the meninges on a Petri dish.
  NOTE: Removal of the meninges from both the brain and spinal cord requires practice. If the user experiences difficulty extracting the meninges, use a dissecting microscope to aid in the removal.
2. Remove the vertebral column from the tube. Using sharp forceps, score around the edges of the vertebral column to free the meninges and peel away the meninges from the edge of the vertebra using curved forceps. Place the meninges on a Petri dish.
3. Place a nylon mesh strainer in a 50 mL conical tube. Move the meninges into the strainer and add 3 mL of RPMI supplemented with 25 mM HEPES. Using the plunger from a 5 mL syringe, grind the tissue and media through the strainer.
4. Using a 5 mL serological pipette, wash the strainer with an additional 2 to 3 mL of RPMI/HEPES media until all visible tissue has passed through the strainer.
  NOTE: To obtain adequate cell numbers for flow cytometric analysis, meninges from multiple animals may need to be pooled together. In this experiment (Figure 3 and Figure 4), the brain and spinal cord meninges from 4–5 mice were pooled together. If the samples are pooled, additional media will be required to grind the tissue through the nylon mesh strainer to prevent overheating of the tissue.
5. Using a 10 mL serological pipette, transfer the cells and media to a fresh 15 mL conical tube. Using a 10 mL serological pipette, wash the 50 mL conical tube with 5 mL of media to collect any remaining cells. Centrifuge at 450 x g for 5 min at 4 °C to pellet the cells.
6. Using a Pasteur pipette with vacuum, aspirate the supernatant being careful to avoid the cell pellet and resuspend the cells in an appropriate volume and buffer.
7. To count the single-cell suspension, dilute a small volume of the cells (i.e., 5–10 µL) using trypan blue exclusion dye (1:10 dilution) and RPMI. Add 10 µL of the dilution to the hemocytometer.
8. Count the cells as previously described⁴⁰^,⁴¹, averaging at least two 16 square grids for accuracy.
  NOTE: In Figure 3, for example, pooled, pelleted cells from the meninges were resuspended in 250 µL of fluorescence-activated cell sorting (FACS) buffer (1x PBS with 1% FBS) for downstream surface staining. The cells were diluted 1:10 for counting (5 µL cells, 5 µL trypan blue, 40 µL RPMI). This dilution yielded between 50–100 cells per 16 square grids, ensuring more accurate cell counting because the cells are neither too dense and overlapping nor too sparse. Nucleated cell counts from pooled brain and spinal cord meninges per mouse were as follows: For meninges from sham-treatment mice = 100,000–150,000 cells and for meninges from Theiler’s murine encephalomyelitis virus-induced demyelinating disease (TMEV-IDD) mice = 300,000–350,000 cells. Cell counts will vary depending on the precision of collection, processing, and if meningeal inflammation is present.
9. Proceed to the desired single-cell technique such as a FACS surface (Figure 3)¹⁴^,³⁶^,⁴²^,⁴³^,⁴⁴, intracellular staining protocols³³^,⁴⁵, in vitro stimulation, cell culture assays³³^,⁴⁶^,⁴⁷, ELISPOT assay²⁸^,³⁴^,³⁵, and bulk or single-cell transcriptomics³⁷^,⁴⁸.
  NOTE: Keep all tubes on ice in between processing steps.
Preparing single-cell suspensions of brain and spinal cord tissue
1. Transfer the brain or spinal cord tissue with media to the top of the 100 mm Petri dish by pouring the tissue and media from the tube. Using forceps, move the tissue to the bottom of the Petri dish. Finely mince the brain or spinal cord with a sterile razor blade. Using the razor blade, move the minced tissue to the bottom of the plate by scraping to gather the tissue.
  NOTE: For the enzymatic digestion protocol below, up to two spinal cords may be pooled together for processing. Brains should be processed individually.
2. Using a 5 mL serological pipette, add 3 mL of RPMI supplemented with 10% fetal calf serum (FCS) to the Petri dish. Using a 10 mL serological pipette, pipette up and down to resuspend the tissue in the media and transfer to a 15 mL conical tube.
  NOTE: If the downstream application is single-cell or bulk RNA sequencing analysis or cell culture, FCS lots should be tested to ensure cells are not activated prior to analysis. Alternatively, cells can be processed using 1x PBS with 0.04% BSA instead of RPMI with 10% FCS.
3. Using a 10 mL serological pipette, wash the Petri dish with an additional 2 mL of media to collect any residual tissue and transfer to a conical tube for a 5 mL total volume. Keep the tubes on ice between processing steps.
4. Using a pipette, resuspend the collagenase I powder in Hank’s Balanced Salt Solution (HBSS) media to obtain the desired concentration (i.e., 100 mg/mL). Add collagenase type I to the conical tube containing the minced tissue sample to obtain the desired final concentration (i.e., 50 µL for 1 mg/mL).
  NOTE: Higher concentrations of collagenase will increase cell yields but can cleave cell surface markers. Therefore, collagenase I lots should be titrated to determine the optimal concentration needed to obtain the highest number of viable cells with all required cell surface markers intact. For example, collagenase I was tested at final concentrations of 0.5 mg/mL, 1 mg/mL, and 2 mg/mL on single brain or spinal cord samples. Cell viability was determined using the trypan blue exclusion method and cell surface markers CD45, CD19, and CD4 were assessed by flow cytometry. A 1 mg/mL concentration of collagenase I yielded the highest live cell count while retaining all cell surface markers of interest. Thus, this concentration was used for further experiments examining these cell types.
5. Gently resuspend DNase I powder using 0.15 M sodium chloride to the desired stock concentration. Add the resuspended DNase I to the conical tube containing the minced tissue sample to obtain a final concentration of 20 U/mL.
  NOTE: DNase I lots vary by units of activity per mL. The concentration to be added to the tissue sample will change based on the stock vial’s units of activity per milliliter. The final desired concentration per sample is 20 U/mL.
6. Place the tubes in a tube rack in a 37 °C water bath and incubate for 40 min. Invert the tubes every 15 min to thoroughly mix the tissue with the enzymes. After incubation, add 500 µL of 0.1 M EDTA (pH = 7.2) to each tube for a final concentration of 0.01 M EDTA and incubate for an additional 5 min to inactivate the collagenase.
7. Using a 10 mL serological pipette, add 9 mL of RPMI supplemented with 10% FCS to each tube to bring the volume of each tube to ~14.5 mL. Centrifuge at 450 x g for 5 min at 4 ˚C. Using a Pasteur pipette with vacuum, aspirate the supernatant being careful not to touch the cell pellet.
8. Using a 5 mL serological pipette, add 3 mL of 100% stock isotonic density gradient solution to the tube containing the cell pellet. Using a 10 mL serological pipette, add additional RPMI 10% FCS media to bring the final volume to 10 mL and resuspend the cell pellet to create a 30% stock isotonic density gradient solution layer.
  NOTE: Prepare the 100% stock isotonic density gradient medium in advance, aliquot, and store at 4 °C for up to 3 months. To prepare the 100% stock isotonic density gradient solution, dilute the density gradient media (Table of Materials) with density gradient media dilution buffer. Prepare the density gradient media dilution buffer (80.0 g/L NaCl, 3.0 g/L KCl; 0.73 g/L Na₂HP0₄, 0.20 g/L KH₂HP0₄; 20.0 g/L glucose) and filter sterilize using a vacuum filter system. Make the 100% stock isotonic density gradient solution by mixing 1 part of density gradient dilution buffer and 9 parts density gradient media. Mix well.
9. Invert and mix each tube well prior to adding the 70% stock isotonic density gradient solution underlay. Insert a 1 mL serological pipette containing 1 mL of 70% stock isotonic density gradient solution into the bottom of the tube. Slowly underlay 1 mL of the solution, being careful not to make bubbles. Slowly remove the serological pipet from the tube, being careful not to disturb the gradient.
  NOTE: For the 70% underlay, the 100% stock isotonic density gradient solution should be diluted to 70% using RPMI media (i.e., 7 mL of 100% stock isotonic density gradient solution and 3 mL of RPMI media mixed well). Additionally, creating a clean, undisturbed 70% underlay is essential for the removal of myelin debris and for obtaining pure single-cell suspensions at the gradient interface following centrifugation.
10. Centrifuge at 800 x g for 30 min at 4 °C with no brake. Aspirate the supernatant, including the myelin debris layer until 2–3 mL remains in the tube, being careful not to disturb the cell layer. Harvest the cell layer between the 30/70% density gradient using a 1 mL pipette and transfer to a new 15 mL conical tube.
11. Using a 10 mL serological pipette, add RPMI 10% FCS media to bring the final volume to 15 mL. Centrifuge 450 x g for 5 min at 4 °C.
  NOTE: During this step, cell layers from two tubes can be pooled if needed. Do not pool more than two tubes or the cells will not pellet due to a high-density gradient media concentration.
12. Aspirate the supernatant carefully to not disturb the cell pellet. Resuspend the cells in an appropriate volume/buffer to count the cells using a hemocytometer (e.g., resuspend a single spinal cord in 250 µL of FACS buffer for downstream surface staining) (Figure 3).
13. Using a 1 mL pipette, transfer the cell suspensions to the top of a filter top tube (Table of Materials) and allow the cells to filter to the bottom of the tube to remove any remaining myelin debris.
14. Using trypan blue exclusion dye, dilute, and count the cells on a hemocytometer by averaging at least two 16 square grids for accuracy⁴⁰^,⁴¹.
15. Proceed with the desired single-cell analysis technique.
  NOTE: Using various forms of collagenase (i.e., D, type I, type II, type IV), immune cells, microglia (Figure 3), astrocytes, pericytes, endothelial cells⁴⁹, and neurons⁵⁰ can all be efficiently isolated. Nucleated cell counts obtained for the results were as follows using the titrated collagenase I enzyme: Whole sham-treated brain = 500,000–600,000 cells; Whole TMEV-IDD brain = 800,000–1,000,000 cells; Whole sham-treated spinal cord = 150,000–200,000 cells; Whole TMEV-IDD spinal cord = 300,000–400,000. Cell counts will vary depending on the precision of collection, processing, and whether CNS inflammation is present.

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Representative Results

This representative experiment was aimed at quantifying B and T cells and describing B and T cell localization in the meningeal and parenchymal CNS compartments in homeostatic conditions as well as in a murine progressive MS model (i.e., TMEV-IDD). TMEV-IDD was induced in 5-week-old female SJL mice by intracranial infection with 5 x 10⁶ plaque forming units (PFU) of TMEV BeAn as previously described²⁹.

The present study assessed B and T cells in the meninges, brain, and spinal cord during chronic TMEV-IDD at day 120 postinfection. Age-matched sham-treated mice were used as controls. The study was comprised of two experiments. The first focused on obtaining single-cell suspensions for flow cytometric assessment, a well-established technique to analyze and quantify cell composition by evaluating cell surface and/or intracellular antigens (n = 4 sham-treated; n = 5 TMEV-IDD). The second experiment focused on describing B and T cell localization in the CNS compartment by utilizing immunohistochemistry on decalcified brain and spinal cord tissues (n = 3 sham-treated; n = 8 TMEV-IDD).

Following the isolation of single-cell suspensions from the brain, spinal cord, and pooled meninges (brain and spinal cord) from TMEV-IDD and sham-treated mice, a surface staining protocol was applied to all samples. Briefly, single-cell suspensions were incubated with a fixable viability exclusion stain (780) for 15 min, washed, blocked with Fc block in the presence of mouse serum for 15 min, and stained with conjugated antibodies for cell surface markers, including CD45 (30-F11; PerCP-Cy5.5), CD19 (1D3; PE-CF594), and CD4 (GK1.5; PE) for 30 min as previously described²⁸^,²⁹. Cells were then washed and analyzed using a flow cytometer²⁸^,²⁹. Viability gating was conducted as previously described²⁸. CD45 expression was assessed to distinguish CD45^hi peripherally derived infiltrating immune cells (P1) from CD45^lo microglia (P2) and CD45^- neurons, astrocytes, and oligodendrocytes in the brain and spinal cord (P3; Figure 3A). In sham-treated mice, few CD45^hi cells were present in the spinal cord and brain tissue. In the meninges, the same gating cut-off for CD45^hi expression used for the brain and spinal cord data was applied to identify CD45^hi immune cells (P1; Figure 3C) and exclude nonimmune cells present in the meninges (i.e., fibroblasts, endothelial cells). In sham-treated mice, few CD45^hi cells (<0.1%) were present in the meninges. Among CD45^hi immune cells in TMEV-IDD CNS tissues, B cells and CD4 T cells were identified by surface expression of CD19 and CD4, respectively (Figure 3B–D). In all TMEV-IDD CNS tissues, increased percentages of CD45^hi immune cells were observed compared to sham-treated mice (Figure 4A). During chronic TMEV-IDD, the percentage of B cells among CD45^hi immune cells in the brain and spinal cord was higher compared to the meningeal compartment (Figure 4B).

To identify the localization of B cells and T cells within the CNS compartment during chronic TMEV-IDD, decalcified brains and spinal cords were evaluated using immunohistochemistry using the staining protocol previously described²⁹. The meninges and vasculature were demarcated using laminin, a basement membrane component²⁹^,⁵¹ and ER-TR7, a fibroblast reticular cell marker⁵²^,⁵³. During conventional extraction of brain from the skull cap, the pia layer was intact, but the remaining meningeal layers were excluded (Figure 5A). In the spinal cord, hydraulic extrusion resulted in the absence of all meningeal layers (data not shown). In both decalcified brains and spinal cords, all meningeal layers were intact (Figure 5B). To examine B and T cell localization in chronic TMEV-IDD, IgG expression was used to determine the localization of isotype-switched B cells²⁹, and CD3 was used to visualize all T cells²⁹. Costaining IgG with ER-TR7 revealed that isotype-switched IgG⁺ B cells were present in the CNS parenchyma and the meninges (Figure 6A). Cellular aggregates within ER-TR7⁺ meninges contained multiple IgG⁺ B cells and CD3⁺ T cells (Figure 6B).

The representative results show high percentages of both B cells and T cells in the parenchymal and meningeal compartments in chronic TMEV-IDD mice in contrast to age-matched sham-treated mice. IHC analysis further demonstrated that in TMEV-IDD tissue, B cells and T cells were dispersed in the parenchyma, but were closely associated in the meninges, forming inflammatory aggregates. In progressive MS patients, meningeal inflammatory aggregates are associated with adjacent tissue injury and worse disease outcomes. In chronic TMEV-IDD, persistent B cell and T cell presence in the CNS compartment and aggregation in the meninges may be associated with tissue injury and disease progression. Further studies are needed to understand how meningeal versus parenchymal inflammation affect demyelination, neurodegeneration, and clinical disability.

Figure 1: Isolating brains and spinal cords for decalcification. Blue dotted lines indicate cuts made to isolate the brain with intact skull (cuts 1-4) and vertebral column (cuts 5-8). Please click here to view a larger version of this figure.

Figure 2: Isolating brains, spinal cords, and meninges for single-cell techniques. (A) Blue dotted lines indicate cuts made to isolate the skull cap with intact meninges and brain (cuts 1-3) and vertebral column (cuts 5-7). Lateral cut 3 is made on both sides of the skull. (B) Blue line indicates incision made to score and remove the meninges in the skull cap and the cuts made to the vertebral column (both lateral sides) to isolate the spinal cord and meninges. Please click here to view a larger version of this figure.

Figure 3: Identification of CD45^hi infiltrating immune cells in CNS tissues. (A) Gating strategy for the identification of CD45^hi infiltrating immune cells (P1), CD45^lo microglia (P2), and CD45^- oligodendrocytes, astrocytes, and neurons among total live cells in spinal cords from sham-treated (red) or TMEV-IDD (blue) mice. Minimal CD45^hi cells were detected in sham-treated brains and spinal cords. (B) Gating strategy for identifying CD19⁺ B cells and CD4⁺ T cells among CD45^hi infiltrating immune cells (P1) in TMEV-IDD mice. Gating strategies were similar for brain and spinal cord tissue. (C) Gating strategy for identifying CD45^hi cells (P1) in the meninges of sham-treated (red) and chronic TMEV-IDD mice (blue). (D) Gating strategy for identifying CD19⁺ B cells and CD4⁺ T cells among CD45^hi infiltrating immune cells (P1) in the meninges of chronic TMEV-IDD mice. Please click here to view a larger version of this figure.

Figure 4: CD45^hi immune cells and CD19⁺ B cells increased in CNS tissues in chronic TMEV-IDD. Bar graphs summaries of flow cytometry data obtained from sham-treated or TMEV-IDD meninges, brain, and spinal cords show percentages of CD45^hi infiltrating immune cells (A) or percentages of CD19⁺ B cells (B). For TMEV-IDD tissues, flow cytometry data were obtained from five mice, with spinal cords and brains processed individually, while meninges were pooled for analysis. For sham-treated mice, flow cytometry data were obtained from four mice, with spinal cords and brains processed individually, while meninges were pooled for analysis. Data for spinal cords and brains are shown as mean ± SEM. Please click here to view a larger version of this figure.

Figure 5: Intact meningeal layers in decalcified brains and spinal cords. (A) Brains extracted from the skull cap or (B) decalcified brains with intact skulls and vertebral columns from chronic-TMEV-IDD mice were assessed for DAPI (blue), meningeal markers laminin (green), and ER-TR7 (red). Scale bar = 50 µm. Please click here to view a larger version of this figure.

Figure 6: Immune cell aggregation was evident in the meninges during chronic TMEV-IDD. (A) Decalcified vertebral columns from chronic-TMEV-IDD mice were examined for the presence of IgG (green) to identify isotype-switched immune cells in the parenchyma and in the ER-TR7+ meninges (red). (B) CD3⁺ T cells (green) and IgG⁺ isotype-switched B cells (red) were costained with ER-TR7⁺ meninges to assess localization within the spinal cord tissue. White arrows highlight B and T cells within the meninges. Scale bar = 50 µm. White boxes delineate areas selected for cropped images (right; scale bar= 10 µm). Please click here to view a larger version of this figure.

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Discussion

Methods for evaluating the cellular composition in the CNS compartment during homeostasis and disease are essential for understanding the physiological and pathological states of the CNS. However, despite serving as an important barrier in the CNS and housing a diverse array of immune cells, the meninges are often omitted from analysis because many conventional tissue extraction methods for the brain and spinal cord do not allow for the collection of these membranes. This omission is a critical limitation in the advancement of our understanding of the cellular composition and function of the meninges and its role in steady state and inflammatory conditions. Recent studies revealed that both resident and peripherally derived immune cells residing in the meningeal compartment play an essential role in maintaining homeostasis in the CNS, as well as in driving CNS disease pathogenesis. These studies emphasize the need to analyze not only the parenchymal compartment but also the surrounding meningeal layers. The protocols described here allow for the rapid isolation of brain and spinal cords while preserving the meninges, ultimately enabling a comprehensive downstream analysis of the CNS compartment utilizing both histologic and single-cell studies. The representative results focus on assessing immune cells in the CNS compartment; however, the protocols may be adapted to analyze microglia, astrocytes, neurons, pericytes, endothelial cells, or other CNS resident cells.

A critical step in the described methods is the careful extraction of the tissue, essential for both isolating pure single-cell suspensions from brain, spinal cord, and meninges and for obtaining CNS tissues with either intact skulls or vertebral columns, allowing high-quality tissue morphology. Practicing the technique for obtaining single-cell suspensions is the key to a successful tissue extraction because it will not only enhance the purity of the sample but also improve cell yields for downstream applications. Similarly, practice in the isolation of CNS tissues with intact skulls or vertebral columns to adequately remove excess tissue surrounding the bone is an essential step that ensures better fixation, decalcification, and cryopreservation of the tissue, all crucial components for obtaining tissue sections with high-quality morphology and intact antigen targets.

One limitation of our protocol for isolating single-cell suspensions from the meninges is that it focuses on obtaining dural and arachnoid meninges³²^,⁵⁴ while omitting the isolation of the pia, which remains attached to the brain or spinal cord via astrocyte processes. Although the cells residing in the pia mater are an essential component of the meningeal compartment, it was decided to exclude the pia mater during the single-cell suspension preparation due to the extensive amount of time required to adequately isolate it⁵⁵. Similarly, the protocol only focuses on obtaining meninges in the superior portion of the cranium and excludes the isolation of the invaginating meninges in the brain and choroid plexus. Collection of the invaginating meninges and choroid plexus can be conducted according to previously published protocols⁵⁵, although it is a time-consuming procedure. In both cases the choice to omit part of the meningeal layers was due to the amount of time it requires to isolate them. The timing of the protocol used to prepare single-cell suspensions required for downstream applications such as flow cytometry, cell culture assays, and RNA sequencing (RNAseq) is critical to improve cell viability and data quality. Therefore, depending on the specific research question, a balance should be struck between sample isolation time and the thoroughness in extracting the meninges. In the current protocol, only single-cell suspension preparations from the dura and arachnoid meninges are obtained, which limits the ability to extrapolate results to the entirety of the CNS meninges. However, if these methods are used in combination with IHC or ISH analysis of whole tissue sections with the three layers of meninges, a more thorough understanding of the cellular and molecular dynamics in the parenchymal and meningeal compartments can be gained.

Another limitation of the protocol is the low cell numbers obtained from the dural and arachnoid meninges, especially during homeostatic conditions. However, if cell numbers in the spinal cord, brain, or the meninges are deemed to be too low for analysis of a particular target cell population, the minimal cell number required for that analysis can be determined by estimating the total number of events required for a given precision (e.g., 5% coefficient of variation) as described in previous literature specializing in rare event analysis⁵⁶. These calculations can then be used to determine if samples from multiple animals must be pooled in order to acquire sufficient cell numbers to analyze the cell population of interest.

The methods described provide an essential advancement in the investigation of the cellular dynamics within the CNS compartment by extending analyses to include the commonly neglected meningeal compartment. These protocols allow for the individual extraction of the brain, spinal cord, and the dural and arachnoid meninges to generate single-cell suspensions. Additionally, the decalcification protocol allows histological analyses of tissue sections comprising brain or spinal cord tissue inclusive of the three meningeal layers. Utilizing EDTA provides a slow but gentle decalcification process, which is most appropriate for specimens where high-quality tissue morphology is required (e.g., IHC and ISH). The protocol uses a pH of 7.2–7.4 to slow the rate of decalcification and ensure the maintenance of intact tissue morphology, a clear advantage over strong and weak acids (e.g., hydrochloric acid and trichloroacetic acid), which have a shorter decalcification time, but impact the quality of the tissue morphology.

Future methods seeking to isolate single-cell suspensions from the meninges should focus on developing protocols to shorten the time required to thoroughly isolate the pial layer from CNS tissues and the invaginating meninges in the brain. Obtaining the complete complex of the three meningeal layers will not only increase the cell numbers for any analysis but will also provide a more comprehensive assessment of the resident and infiltrating immune cells occupying the meninges, which enclose the CNS. Concurrently, future protocols focused on preparing decalcified brains and spinal cords should seek to identify novel agents aimed at expediting tissue decalcification while preserving the quality of tissue morphology. However, altogether, downstream applications analyzing CNS single-cell suspensions inclusive of dual and arachnoid meninges, performed in combination with comprehensive tissue section analysis, can provide a detailed understanding of cell phenotype, function, and localization within the CNS compartment.

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Disclosures

The authors have nothing to disclose.

Acknowledgments

The authors thank the staff of the Center for Comparative Medicine and Research (CCMR) at Dartmouth for their expert care of the mice used for these studies. The Bornstein Research Fund funded this research.

Materials

Name	Company	Catalog Number	Comments
Aluminum foil	any	N/A
Bovine Serum Albumin	ThermoFisher Scientific	37002D
Centrifuge	Beckman Coulter	Allegra X-12R centrifuge
Collagenase I	Worthington	LS004196
Conical tube, 15 mL	VWR	525-1069
Conical tube, 50 mL	VWR	89039-658
Cover glass	Hauser Scientific	5000
Cryomold	VWR	18000-128
Curved forceps	Fine Science Tools	11003-14
Disposable polystyrene tube, 14 mL	Fisher Scientific	14-959-1B
Disposable Scalpel	Fisher Scientific	NC0595256
DNAse I	Worthington	LS002139
Dry ice	Airgas	N/A
Durmont #7Forceps	Fine Science Tools	11271-30
EDTA disodium salt dihydrate	Amresco	0105-500g
Ethanol, 100%	any	N/A
Fetal Bovine Serum (FBS)	Hyclone	SH30910.03
Filter top tube, 5 mL	VWR	352235
Fixable viability stain 780	Becton Dickinson	565388
Flow cytometer	Beckman Coulter	Gallios
Glucose	Fisher Chemical	D16-500
Goat anti-mouse IgG (488 conjugate)	Jackson immunoresearch	115-546-146
Goat anti-mouse IgG (594 conjugate)	Jackson immunoresearch	115-586-146
Goat anti-rabbit 488	Jackson immunoresearch	111-545-144
Goat anti-rat 594	Jackson immunoresearch	112-585-167
Goat anti-rat 650	Jackson immunoresearch	112-605-167
Hank's Balnced Salt Solution (HBSS)	Corning	21-020-CV
Hemacytometer	Andwin Scientific	02-671-51B
Hemostat	Fine Science Tools	13004-14
HEPES (N-2-hydroxyethylpiperazine-N-2-ethane sulfonic acid)	ThermoFisher Scientific	15630080
KCl	Fisher chemical	BP366-500
KH₂PO₄ (anhydrous)	Sigma Aldrich	P5655-100G
Liquid Nitrogen	Airgas	N/A
Mouse FC block (CD16/32)	Becton Dickinson	553141
Na₂HP0₄ (anhydrous)	Fisher Chemical	S374-500
NaCl	Fisher chemical	S671-500
Needle, 25 gauge	Becton Dickinson	305122
Normal mouse serum	ThermoFisher Scientific	31881
Nylon mesh strainer	VWR	352350
OCT	Sakura	4583
Paraformaldehyde, 20%	Electron Microscopy Sciences	15713-S	Diluted to 4% using 1 x PBS
Pasteur pipette, 9 inch, unplugged	Fisher Scientific	13-678-20C
PBS (1x)	Corning	21-040-CV
PE Rat Anti-Mouse CD4	Becton Dickinson	553730
PE-CF594 Rat Anti-Mouse CD19	Becton Dickinson	562329
Percoll density gradient media	GE healthcare	17-0891-01
PerCP-Cy5.5 Rat Anti-Mouse CD45	Becton Dickinson	550994
Petri dish, 100 mm	VWR	353003
pH meter	Fisher Scientific	13-636-AB150
Pipet-Aid	Drummond Scientific Corporation	4-000-101
Pipette 200 µl	Gilson	FA10005M
Pipette tips, 1 mL	USA Scientific	1111-2831
Pipette tips, 200 µl	USA Scientific	1111-1816
Pipette, 1 mL	Gilson	FA10006M
Prolong Diamond mountant with DAPI	ThermoFisher Scientific	P36962
Purified Rat Anti-Mouse CD16/CD32	Becton Dickinson	553141
Rabbit anti-mouse CD3 (SP7 clone)	Abcam	ab16669
Rabbit anti-mouse laminin	Abcam	ab11575
Rat anti-mouse ERT-R7	Abcam	ab51824
RPMI 1640	Corning	10-040-CV
Serological pipet, 1 mL	VWR	357521
Serological pipet, 10 mL	VWR	357551
Serological pipet, 5 mL	VWR	357543
Sodium hydroxide	Fisher Scientific	S318-100
Sucrose	Fisher chemical	S5-500
Surgical scissors	Fine Science Tools	14001-16
Surgical scissors, extra fine	Roboz	RS-5882
Syringe, 10 mL	Becton Dickinson	302995
Syringe, 5 mL	Becton Dickinson	309646
Trypan blue	Gibco	15250-061
Vacuum filter system	Millipore	20207749
Vacuum flask	Thomas Scientific	5340-2L
Vacuum in-line filter	Pall Corporation	4402
Vacuum line	Cole Palmer	EW-06414-20
Water bath	ThermoFisher Scientific	Versa bath

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Immunology and Infection