Assessing the Expression of Major Histocompatibility Complex Class I on Primary Murine Hippocampal Neurons by Flow Cytometry

All procedures were performed in compliance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health and according to the International Guiding Principles for Biomedical Research Involving Animals. The protocol was approved by the University of North Carolina at Charlotte Institutional Animal Care and Use Committee (Protocol #19-020).

1. Preparing for culture

NOTE: These procedures should be done under sterile conditions in a tissue culture-designated biosafety cabinet. See Table 1 for media and solutions.

1. Sterilize all dissection tools by autoclaving in self-sealing sterilization bags.
2. Prepare working stocks of poly-D-lysine for treating 12-well culture dishes.
   1. Dissolve poly-D-lysine in 1x borate buffer to a concentration of 100 µg/mL (10x concentrated). Store approximately 1 mL aliquots at -20 ˚C until needed.
   2. Dilute 10x concentrated poly-D-lysine in dPBS to 10 µg/mL (1x) and filter sterilize.
   3. Treat 12-well culture plates with 0.5 mL per well of 1x poly-D-lysine at room temperature for at least 1 h or overnight for using the next day. Ensure the volume of poly-D-lysine is enough to fully cover the bottom of the culture area.
   4. Just prior to the use, remove poly-D-lysine from culture plate, rinse 3x in sterile water, and keep in sterile water until ready to plate cells. Remove all traces of liquid by a thorough aspiration prior to plating cells.
3. Prepare Neuron Growth Media and FACS buffer. Filter sterilize and store at 4 ˚C until use. Neuron Growth Media can be kept for approximately 2 wks; FACS buffer can be kept for approximately 4 wks.

2. Dissecting embryonic hippocampus

NOTE: This procedure can be performed on the benchtop as it requires the use of a stereo microscope for the removal of meninges and microdissection of the hippocampus. Adhere to strict aseptic technique to minimize potential contamination.

1. Euthanize a timed-pregnant C57BL/6J female mouse at embryonic day 18 in a CO₂ chamber.
2. Spray the abdominal skin and fur with 70% EtOH, then pinch skin with tissue forceps and make small incision with surgical scissors. Incise the peritoneum to expose the viscera and uterus.
3. Grasp the uterus with standard pattern forceps, lift out of cavity, and cut the connection to the mesometrium with fine scissors. Transfer uterus with embryos to 100 mm sterile plastic culture dish placed on ice.
4. Using fine scissors and fine forceps, carefully cut the uterus and remove the embryonic sacs to release embryos. Transfer embryos to a new 100 mm sterile plastic culture dish containing dPBS placed on ice.
5. Decapitate embryonic pups using fine scissors, and transfer heads to new 100 mm sterile plastic culture dish containing Hibernate-E medium placed on ice.
6. To remove the brain, hold the head with a pair of fine forceps in one hand. Using curved Dumont #7 forceps in other hand, insert the tip of the forceps at the base of the skull and pinch the bone and overlying skin moving anteriorly along the midline without piercing the brain.
7. Using curved forceps pull apart the skin and skull to expose the brain. Sweep the curved forceps under the brain from the exposed olfactory bulb to the cerebellum to lift the brain out of the skull. Transfer the brain to a new 100 mm sterile plastic culture dish containing Hibernate-E medium placed on ice. Repeat for each brain.
   NOTE: All brains may be collected in a single culture dish unless needed to be separate for experiment-specific purposes.
8. Under a stereo dissection microscope, working with one brain at a time and two pairs of sterile Dumont #5 forceps, pinch off olfactory bulbs and pull away meninges. Thorough removal of the meninges is necessary to avoid culture contamination by other cells and to allow further dissection.
9. Once meninges are properly removed, the superior side of the cortex opens laterally, which will expose the hippocampus. Using Dumont #5 forceps, pinch the hippocampus away from the attached cortex, and carefully transfer isolated hippocampus to a sterile 15 mL conical tube containing 5 mL of Hibernate-E medium on ice.
10. Repeat the dissection for both brain hemispheres for each embryonic mouse pup and combine all hippocampi into a single 15 mL conical tube, unless needed separately for experiment specific purposes.
    1. To culture cortical neurons, the cortex may be separated from the subcortex and processed identically to hippocampal neurons.
       NOTE: At this point, protocol may be paused. Store brains or dissected hippocampi in Hibernate-E medium supplemented with B27 (2%) at 4 ˚C for up to 1 month, though extended delay may compromise the number of viable cells.

3. Dissociating and culturing hippocampal neurons

NOTE: All procedures should be performed under sterile conditions in a tissue culture designated biosafety cabinet.

1. Prepare 0.5 mL of papain dissociation solution per embryo for hippocampal dissociation. The volume of dissociation solution needed will vary depending on the number of embryonic brains being dissected.
   NOTE: For cortical neuron culture, prepare 1.0 mL of papain solution per embryo.
   1. Dissolve 20 µL of papain suspension per 1 mL of the Hibernate E medium by vortexing for approximately 3 min.
   2. After papain is dissolved, add 1 µL of DNase I per 1 mL of the solution. Do not vortex the solution after DNase I has been added as this can deactivate the enzyme.
2. Centrifuge 15 mL conical tube containing hippocampal tissue (step 2.10) at 1, 000 x g for 5 min. Remove the supernatant and add papain solution. Resuspend the tissue by inverting several times. Incubate at 37 ˚C for 30 min, inverting to resuspend the tissue every 10 min.
   1. If culturing cortical neurons, transfer all cortices to a new 100 mm sterile culture dish, carefully aspirate media from plate, and mince tissue with fine scissors or razor blade. Add papain solution to the culture dish and transfer cortical tissue with papain solution to 15 mL conical tube using a sterile transfer pipette. Incubate tissue and enzyme solution at 37 ˚C for 30 min, agitating every 10 min.
3. During 30 min incubation, prepare 3 fire-polished glass Pasteur pipettes: 1 fully open, 1 half open, and 1 quarter open.
4. After the 30 min incubation, centrifuge the 15 mL conical tube containing digested brain tissues at 125 x g for 10 min. Remove the enzyme solution and replace with an equal volume of fresh Hibernate E medium.
5. Using fully open Pasteur pipette, triturate the tissue 10x. Let the tissue settle for 2 min, then transfer the supernatant cell suspension to a sterile 50 mL conical tube. Add an equal volume of Hibernate E medium back to the tissue.
6. Using half open Pasteur pipette, triturate tissue 10x. Let the tissue settle for 2 min, then transfer the supernatant cell suspension to the same 50 mL conical tube as in the previous step. Add an equal volume of Hibernate E medium back to the tissue.
7. Using quarter open Pasteur pipette, triturate tissue 10x. Let the tissue settle for 2 min, then transfer the supernatant cell suspension again to the same 50 mL conical as in previous steps. Discard any remaining tissue that hasn’t dissociated.
8. Centrifuge the 50 mL conical tube containing the supernatant from the cell suspension at 125 x g for 5 min. If not already done, wash poly-D-lysine-coated plates with sterile water and remove all traces of liquid by thorough aspiration during centrifugation.
9. After centrifugation, discard the supernatant and resuspend the cell pellet in 5 mL of Neuron Growth Media. Count live cells using trypan blue and hemocytometer.
   1. If culturing cortical neurons, add at least 10 mL of Neuron Growth Media for less dense cells and more accurate counting.
10. Dilute cells with Neuron Growth Media for a final plating density of 5 x 10⁵ viable cells per ml and add 1 mL of diluted cell suspension to each well of a 12 well plate. Maintain neurons at 37 ˚C with 5% CO₂.
    NOTE: Typically, the hippocampi from 1 embryonic mouse brain, i.e., 2 hippocampi, yield approximately 1 x 10⁶ viable cells. This number is provided for experiment-planning purposes only and should not be considered a substitution for counting cells for each culture.
11. Change the growth media the day after culture by removing half of the volume of media from each well, i.e., 0.5 mL, and add an equal volume of fresh media to the side of the well to avoid disrupting the cultured cells. Complete half media changes twice weekly for the lifespan of the culture.

4. Assessing MHCI expression by flow cytometry

1. Prepare treatment by diluting Interferon beta (IFNβ) or an equal volume of diluent in Neuron Growth Media. Since media change will be a half-volume change, prepare treatment media with 2x concentrated IFNβ. i.e., to treat 3 wells of a 12 well plate with 100 U/mL of IFNβ, prepare 1.5 mL with 200 U/mL IFNβ or and equal volume of IFNβ diluent for media-treated controls.
2. Remove 0.5 mL from each well of neurons and add 0.5 mL of Neuron Growth Media with or without IFNβ to each well. Incubate in normal growth conditions (37 ˚C, 5% CO₂) for 6-72 h, depending on experimental conditions and signal optimization.
3. Following treatment time, remove all culture media and gently wash once in cold Neurobasal media lacking supplements (B27, L-glutamine, Pen-Strep).
4. Add 0.5 mL of non-supplemented cold Neurobasal media containing 1 µg/mL of Fc block and 1 µg/mL of fluorescent conjugated anti-MHCI antibody to each well. Incubate at 4 ˚C protected from light for 45 min.
5. Remove antibody-containing media and carefully wash once in cold dPBS. Add 0.5 mL room-temperature enzyme-free cell dissociation buffer to each well and agitate to dislodge cells. Watch for dissociation using inverted tissue culture microscope.
6. Once cells are detached from culture dish, add 0.5 mL of FACS buffer to each well, triturate cells to disperse clumps, and transfer total volume of each well to individual 1.7 mL microcentrifuge tubes.
7. Centrifuge at 1, 000 x g for 5 min. Remove the supernatant, resuspend the cell pellet in 100 µL of FACS buffer, and transfer total volume of each tube to individual wells of a 96 well U-bottom plate.
8. Add 100 µL of fixative reagent to each well. Triturate several times to avoid cells clumping and incubate for 15 min at room temperature protected from light.
9. Centrifuge at 500 x g for 5 min. Remove the supernatant and resuspend in 200 µl of FACS buffer.
10. Centrifuge at 500 x g for 5 min. Remove the supernatant and resuspend in 100 µL of permeabilization reagent containing fluorescent-conjugated anti-NeuN antibody (1:100 dilution) to each well. Mix well, then incubate at room temperature protected from light with rocking for 20 min.
11. After incubation, centrifuge at 500 x g for 5 min. Remove the supernatant and resuspend in 100 µL of FACS buffer. Repeat 2x.
12. After the final wash, resuspend cells in 100 µL of 2% paraformaldehyde diluted in FACS buffer. Mix well to prevent cells from clumping. Store at 4 ˚C protected from light until ready to read on a flow cytometer. Read samples as soon as possible, within 1 week.

5. Quantifying and evaluating data

1. Measure the compensation controls for each fluorescent dye and correct any spectral overlaps. Ideal compensation controls include cultured neurons that are unstained, stained with anti-MHCI only, and stained with anti-NeuN only.
2. If possible, record 100,000 events for each sample (at least 10,000) and save as FCS files.
3. To analyze data, using appropriate analysis software, set up sequential gates, as depicted in Figure 1A-C to select for NeuN-positive neurons.
   1. Plot SSC-A (log) vs FSC-A (linear). Draw a gate on the cellular population (P1) to eliminate cellular debris from the analysis.
   2. Within P1 cellular population, plot FSC-H (linear) vs FSC-A (linear). Draw a gate on single cell population.
   3. Within the single cell population, plot SSC-A (log) vs NeuN (log). Draw a gate on NeuN-positive population using unstained or MHCI-only stained cells as a guide.
   4. Plot a histogram of MHCI fluorescence with cellular events normalized to mode on y-axis. Using unstained or NeuN-only stained neurons as a guide, draw a horizontal gate to quantify the percent neurons positive for MHCI.
   5. Export the percent cells positive for MHCI, as well as the median fluorescence intensity (MFI) for MHCI on the NeuN-positive population for statistical evaluation and graphical drawing.