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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

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.

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

  1. 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.
  2. 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.
  3. 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

  1. 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.
  2. 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.
  3. 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.
  4. 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.

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,...

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 advancem...

Disclosures

The authors have nothing to disclose.

Acknowledgements

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

NameCompanyCatalog NumberComments
Aluminum foilanyN/A
Bovine Serum AlbuminThermoFisher Scientific37002D
CentrifugeBeckman CoulterAllegra X-12R centrifuge
Collagenase IWorthingtonLS004196
Conical tube, 15 mLVWR525-1069
Conical tube, 50 mLVWR89039-658
Cover glassHauser Scientific5000
CryomoldVWR18000-128
Curved forcepsFine Science Tools11003-14
Disposable polystyrene tube, 14 mLFisher Scientific14-959-1B
Disposable ScalpelFisher ScientificNC0595256
DNAse IWorthingtonLS002139
Dry iceAirgasN/A
Durmont #7ForcepsFine Science Tools11271-30
EDTA disodium salt dihydrateAmresco0105-500g
Ethanol, 100%anyN/A
Fetal Bovine Serum (FBS)HycloneSH30910.03
Filter top tube, 5 mLVWR352235
Fixable viability stain 780Becton Dickinson565388
Flow cytometerBeckman CoulterGallios
GlucoseFisher ChemicalD16-500
Goat anti-mouse IgG (488 conjugate)Jackson immunoresearch115-546-146
Goat anti-mouse IgG (594 conjugate)Jackson immunoresearch115-586-146
Goat anti-rabbit 488Jackson immunoresearch111-545-144
Goat anti-rat 594Jackson immunoresearch112-585-167
Goat anti-rat 650Jackson immunoresearch112-605-167
Hank's Balnced Salt Solution (HBSS)Corning21-020-CV
HemacytometerAndwin Scientific02-671-51B
HemostatFine Science Tools13004-14
HEPES (N-2-hydroxyethylpiperazine-N-2-ethane sulfonic acid)ThermoFisher Scientific15630080
KClFisher chemicalBP366-500
KH2PO4 (anhydrous)Sigma AldrichP5655-100G
Liquid NitrogenAirgasN/A
Mouse FC block (CD16/32)Becton Dickinson553141
Na2HP04 (anhydrous)Fisher ChemicalS374-500
NaClFisher chemicalS671-500
Needle, 25 gaugeBecton Dickinson305122
Normal mouse serumThermoFisher Scientific31881
Nylon mesh strainerVWR352350
OCTSakura4583
Paraformaldehyde, 20%Electron Microscopy Sciences15713-SDiluted to 4% using 1 x PBS
Pasteur pipette, 9 inch, unpluggedFisher Scientific13-678-20C
PBS (1x)Corning21-040-CV
PE Rat Anti-Mouse CD4Becton Dickinson553730
PE-CF594 Rat Anti-Mouse CD19Becton Dickinson562329
Percoll density gradient mediaGE healthcare17-0891-01
PerCP-Cy5.5 Rat Anti-Mouse CD45Becton Dickinson550994
Petri dish, 100 mmVWR353003
pH meterFisher Scientific13-636-AB150
Pipet-AidDrummond Scientific Corporation4-000-101
Pipette 200 µlGilsonFA10005M
Pipette tips, 1 mLUSA Scientific1111-2831
Pipette tips, 200 µlUSA Scientific1111-1816
Pipette, 1 mLGilsonFA10006M
Prolong Diamond mountant with DAPIThermoFisher ScientificP36962
Purified Rat Anti-Mouse CD16/CD32Becton Dickinson553141
Rabbit anti-mouse CD3 (SP7 clone)Abcamab16669
Rabbit anti-mouse lamininAbcamab11575
Rat anti-mouse ERT-R7Abcamab51824
RPMI 1640Corning10-040-CV
Serological pipet, 1 mLVWR357521
Serological pipet, 10 mLVWR357551
Serological pipet, 5 mLVWR357543
Sodium hydroxideFisher ScientificS318-100
SucroseFisher chemicalS5-500
Surgical scissorsFine Science Tools14001-16
Surgical scissors, extra fineRobozRS-5882
Syringe, 10 mLBecton Dickinson302995
Syringe, 5 mLBecton Dickinson309646
Trypan blueGibco15250-061
Vacuum filter systemMillipore20207749
Vacuum flaskThomas Scientific5340-2L
Vacuum in-line filterPall Corporation4402
Vacuum lineCole PalmerEW-06414-20
Water bathThermoFisher ScientificVersa bath

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