Real-time Live-cell Flow Cytometry to Investigate Calcium Influx, Pore Formation, and Phagocytosis by P2X7 Receptors in Adult Neural Progenitor Cells

Animals were treated in accordance with the Australian Code of Practice for the Care and Use of Animals for Scientific Purposes and approved for use by the Griffith University Animal Ethics committee.

1. Neurosphere Culture of Neural Progenitor Cells from the Adult SVZ and Hippocampus

NOTE: The dissection protocol presented here is based on work by Walker and Kempermann, and a detailed protocol for the dissection of neural progenitor cells from adult mice is available elsewhere¹⁴. Culture conditions have been modified from Babu and colleagues¹⁵. Adult female C57BL/6 mice aged 8–12 weeks were used.

1. In a biological safety cabinet, prepare the culture medium (neural basal medium) supplemented with neural stem cell proliferation supplement, 2 mM glutamine, 20 ng/mL epidermal growth factor (EGF), 10 ng/mL basic fibroblast growth factor (bFGF), and 2 µg/mL heparin.
2. Euthanize two mice by CO₂ inhalation. Alternatively, anesthetize the mice according to institutional guidelines and immediately euthanize them by cervical dislocation. Spray their heads with 70% ethanol and decapitate them. Transfer each head to a sterile tube containing Hank’s balanced salt solution (HBSS) with 1x penicillin/streptomycin solution (P/S).
3. Perform dissections in a laminar flow hood under a dissection microscope.
4. Using forceps and dissection scissors, remove tissue and bone to expose the brain, and transfer it to a sterile dish containing HBSS with P/S.
5. Position the brain ventral side up and use a scalpel blade to make a complete coronal incision across the optic chiasm. Hold the brain steady with the forceps and use one swift, clean, downward motion. Avoid sawing to minimize cell death and to help maintain tissue structure.
   NOTE: If not held securely, the brain may tilt forward during the cut, compromising the number of progenitor cells obtained from the SVZ.
6. Isolate the SVZ from the rostral half of the brain.
   1. Locate both ventricles, separated by the septum, with the corpus callosum forming a white bridge above them.
   2. Use forceps to remove the septum separating the two ventricles and isolate the lateral walls of the anterior lateral ventricles. Do this by cutting away above, below, to the sides, and at the front with fine dissection scissors. There will remain just a small cup-like shape.
   3. Prepare a clean Petri dish lid with a few drops of HBSS in the middle and transfer the SVZ to the liquid. Do not allow the tissue to dry or to touch a dry surface. Stand to the side while dissecting the hippocampi.
7. Isolate the hippocampi from the caudal half of the brain.
   1. Make a midline cut between the hemispheres to sever the corpus callosum. Use the lateral ventricles as a guide and gently unfold the cortex to expose the hippocampus. Once the cortex has been unrolled, the hippocampus can be seen as a dense, white, curved structure.
   2. Use fine dissection scissors or forceps to isolate the hippocampus from the neighboring tissue.
   3. With forceps, remove excess white matter, blood vessels, and any membranous tissue covering the hippocampus.
   4. Prepare a clean Petri dish lid prepared with a few drops of HBSS and transfer the hippocampus to the liquid. Repeat for the other hemisphere.
8. Once the hippocampus and the SVZ from two mice have been isolated and transferred to their respective Petri dish lids, mechanically dice the tissue using a scalpel blade. Chop the tissue in one Petri dish lid for approximately 1 min, rotating every 10 s until the tissue appears smooth and only fine pieces remain. Take a clean scalpel blade and repeat for the other dish, dicing the tissue for 1 min.
9. Using a 1 mL pipette, transfer all the tissue from each Petri dish lid to separate tubes containing 1 mL of 0.25% trypsin-ethylenediaminetetraacetic acid (EDTA). Use one tube for the SVZ and one tube for the hippocampus.
   1. Do this by first pipetting about half of the trypsin-EDTA to the tissue in the dish to minimize air bubbles and to prevent the tissue coming into contact with the dry pipette tip as it is transferred to the tube. Rinse the dish and the scalpel blade with the rest of the trypsin-EDTA to collect as much tissue as possible and add it to the tube.
10. Incubate the tissue with the trypsin in a 37 °C water bath for 30 min, agitating the tube every 10 min to properly dissociate the tissue.
11. Triturate the tissue using a fire-polished glass Pasteur pipette to create a single cell suspension. Take care not to overtriturate as this will cause excessive cell lysis. This is a critical step for optimal tissue dissociation.
12. Observe the tube contents during the trituration process and stop at the earliest signs that the majority of tissue clumps have been removed. The trypsin solution should become slightly cloudy when the cells have gone into single-cell suspension, although some clumps may remain.
    NOTE: As an indication, passing the suspension up and down 10x–15x is adequate.
13. Add culture medium up to 5 mL to neutralize the trypsin, and spin at 300 x g for 3 min. Wash a further 2x with HBSS and pass the medium through a 70 µm cell strainer to remove any tissue clumps.
14. Transfer all the cells in 15 mL of medium to a T75 cell culture flask.
    NOTE: Spheres should form in the passage zero (P0) SVZ culture after 7–10 days and after 15–20 days in the hippocampal culture. Refrain from washing or feeding the cells during this time to maximize the number of neural progenitor cells in culture. If more progenitors are required, or to decrease the P0 incubation time, it is possible to increase the number of mice sacrificed.
15. Maintain cultures at 37 °C with 5% CO₂ and, following the initial P0 culture phase, passage every 7–10 days as necessary, using a biological safety cabinet and standard tissue culture practices.
    NOTE: The P0 hippocampal culture should generate enough spheres to passage into a T25 flask; the SVZ culture will generate many more spheres and can be passaged into a T75. If the P0 hippocampal spheres have adhered, use a 200 µL pipette to gently lift the sphere.
16. Passage the spheres when they reach 150 to 200 µm in diameter. Collect the spheres and medium in a 15 mL tube and allow the spheres to settle by gravity for approximately 5 min. Alternatively, spin at a low speed (100 x g) for 2 min.
17. Remove the medium and dissociate the cells with dissociation reagent for 7–10 min, depending on the size of the spheres.
    NOTE: The dissociation reagent used will have significant effects on the outcome of the culture, so please refer to the Table of Materials for specifics.
18. Wash the cells by adding 5 mL of HBSS to the cell solution and centrifuge at 300 x g for 3 min. Remove the supernatant, resuspend the cells in culture medium, and seed at approximately 150,000 cells in 5 mL of medium in a T25 cell culture flask or equivalent. Maintain the cultures at 37 °C and 5% CO₂.
19. Confirm neural progenitor cell status by identifying the expression of stem cell markers such as Sox2 and ASCL1 before proceeding to any downstream protocol⁹.
    NOTE: This can be done by the researcher’s preferred method (e.g., immunochemistry by microscopy, flow cytometry or western blot, or by using quantitative polymerase chain reaction (qPCR)).

2. Preparation of a Single-cell Suspension for Analysis by Flow Cytometry

1. Prepare the required media. These include the following.
   1. Prepare Na⁺ medium containing 145 mM NaCl, 5 mM KOH, 10 mM HEPES, 5 mM D-glucose, 0.1% bovine serum albumin (BSA), and 0.1 mM CaCl₂. This medium is also used in a calcium-free form, with 0.1 mM CaCl₂ omitted.
   2. Prepare K⁺ medium containing 145 mM KCl, 5 mM KOH, 10 mM HEPES, 5 mM D-glucose, and 0.1% BSA.
      NOTE: These media were extensively optimized and are detailed in previous publications^4,5.
   3. Prepare Stock ATP by weighing enough ATP powder for around 20 mL of a 100 mM stock. Dissolve the powder in 17 mL of KCl buffer (145 mM KCl, 5 mM KOH, and 10 mM HEPES, pH 7.5) and slowly, while stirring, add 2 mL of 18% (w/v) tetramethylammonium hydroxide (TMA) to the solution to bring the pH to 6.8–7.0.
   4. Adjust the final volume to 20 mL. Do not overshoot the pH past 7.0. Store the stock at -80 °C; note that the ATP aliquots are stable for at least 6 months.
      NOTE: The free molecular weight of anhydrous ATP is 551.14 g/mol, and this does not include the molecular weight of the water and disodium molecules, which may vary from batch to batch and should be taken into consideration for the calculations. TMA is toxic, and care must be taken when handling. Read the safety data sheet and prepare the stocks in a fume cabinet.
   5. Prepare stock BzATP by dissolving the BzATP in ultrapure H₂O for a final stock concentration of 10 mM.
2. Create a single-cell suspension as described in steps 1.16 and 1.17. Count the cells using a hemocytometer or automatic cell counter. Resuspend the cells in the required medium (e.g., Na⁺ medium, culture medium) for the experiment being conducted (as described below) and place the cells on ice in the meantime.
3. For each experiment, ensure there is enough sample to include controls for forward and side scatter, as well as for the calibration of voltage and compensation settings. Note, as an indication, that spheres grown in a T75 flask typically yield around 8 x 10⁶ cells per flask.
4. Set the flow cytometer settings using control samples.
   1. Use forward and side scatter to selectively gate the living cells.
      NOTE: Forward scatter provides information regarding cell size based on light diffraction, while side scatter provides a measure of internal complexity or granularity. Flow events with small forward and side scatter may be considered as dead cells.
   2. Adjust the voltage and gain of the flow cytometer according to the manufacturer’s instructions. Run a trial sample to ensure the capture of data at the maximum fluorescence intensity. No compensation is required for single channel acquisitions.

3. Measuring Calcium Influx by Live-cell flow Cytometry

1. Following the preparation of a single-cell suspension, resuspend the cells in 1 mL of calcium-free Na⁺ medium and load them with 2 ng/mL of calcium indicator dye according to the manufacturer’s protocol (refer to the Table of Materials) with 10 µL of 5% pluronic acid. Incubate the cells for 30 min at 37 °C.
2. Wash the cells by adding 3–5 mL of calcium-free Na⁺ medium and centrifuging gently (200 x g for 4 min). Remove the supernatant and resuspend the cells in calcium-free Na⁺ medium, washing a second time.
3. Resuspend the cells in 1 mL of calcium-free Na⁺ medium, place them on ice, and allow them to de-esterify for 30 min.
4. Wash 1x more by adding 3–5 mL of K⁺ medium and centrifuging (200 x g for 4 min); then, resuspend the cells in K⁺ medium and aliquot them into fluorescence assisted cell sorting (FACS) tubes at a concentration of 1 x 10⁶ cells per 500 µL per FACS tube.
   NOTE: The number of FACS tubes per sample will depend on the number of treatments and repeats that are included.
5. Place the FACS tubes on ice until the cells are ready to be analyzed. Do not leave the cells on ice for an extended period but begin the assay as soon as possible.
6. For some samples, preincubate the cells with P2X7 receptor-specific inhibitor AZ10606120 (1 µM for 10–15 min) or A438079 (10 µM for 30 min) at 37 °C prior to analysis.
7. A few minutes prior to running the first sample, add CaCl₂ (to a final concentration of 1 mM in the FACS tube) and place the tube in a 37 °C water bath to recover.
8. Drop a clean, small magnetic stirrer into the FACS tube and position the tube in the time module linked to a circulating 37 °C water bath to control the sample temperature. Select a low stirring speed to ensure movement of the sample without introducing a vortex effect. Place the water jacket tube adapter onto the sample platform and close the lever arm of the FACS machine.
9. Initiate sample acquisition and run the sample for 3 min at around 1,000 events per second.
10. At the 40 s mark, quickly remove the tube and add the P2X7 agonist, either 1 mM ATP or 300 µM BzATP, and replace the tube to continue the acquisition.
11. While the first sample is recording, prepare the second sample with CaCl₂ and place it in 37 °C to allow sufficient time for the cells to warm up prior to analysis. Once the first sample has finished, clean the intake by running a water sample, and then the acquisition of the second sample can begin as described in steps 3.8 and 3.9. Always clean the intake between samples.

4. Measuring Pore Formation by Live-cell Flow Cytometry

1. Create a single-cell suspension as described in steps 1.16 and 1.17. Save a few milliliters of the old medium, use it to resuspend the cells at a concentration of 1 x 10⁶ cells per 100 µL per FACS tube, and place it on ice until ready.
2. Prior to running the assay, add 900 µL of K⁺ medium for a final volume of 1 mL and place the tubes in a 37 °C water bath for 10 min to recover.
3. If applicable, preincubate the cells with treatments, including the P2X7-specific inhibitors AZ10606120 (1 µM for 10–15 min) or A438079 (10 µM for 30 min).
4. Immediately prior to running the assay, add 25 µM ethidium bromide to the FACS tube; then, add the magnetic stirrer, place it on the FACS machine according to step 3.7, and begin the acquisition.
   NOTE: Ethidium bromide is toxic, and care must be taken when handling it. Dispose of used FACS tubes appropriately.
5. To induce the formation of pores in the cell membrane, add 1 mM ATP or 100 µM BzATP 40 s after the start of the acquisition.
6. Run the samples at around 1,000 events per second for 6 min.
7. While the first sample is running, take the second sample from the ice and place it in the 37 °C water bath to allow sufficient time for the cells to recover prior to the analysis. Once the first sample has finished with the acquisition, clean the intake by running a water sample; then, the second sample can be placed on the machine to begin recording as described in steps 3.8 and 3.9.

5. Measuring Phagocytosis by Live-cell Flow Cytometry

1. Create a single-cell suspension as described in steps 1.16 and 1.17. Resuspend the cells in conditioned medium and aliquot them into FACS tubes at a concentration of at least 1 x 10⁶ cells per 100 µL per FACS tube. Dilute the cells to a final concentration of 1 x 10⁶ cells/mL with Na⁺ medium (e.g., add 900 µL of Na⁺ medium) and place the cells on ice until the analysis is performed.
2. Use 1 µm of YG latex beads (microspheres) as phagocytic targets for real-time phagocytosis assays.
   NOTE: Other colors may also be substituted, but differently sized beads were found to be inadequate targets of phagocytosis⁴.
3. Prior to running the first sample, transfer the cells to a 37 °C water bath and incubate them for approximately 7–10 min to allow the cells to recover.
4. Add any treatments requiring preincubation to their respective tubes, including 1 mM ATP for 15 min, 300 µM oxidized ATP (oxATP) for 40 min, 20 µM cytochalasin D for 20 min, and 4% paraformaldehyde (PFA) for 20 min.
   1. No preincubation is required for 5% human serum. If treatments are added at approximately the same time, the samples can be run in reverse order while the others continue to incubate. For example, run the controls and serum first, then the ATP-treated sample, followed by cytochalasin D and PFA, and oxATP last.
5. Place the sample on the cytometer with the magnetic stirrer as described in steps 3.8 and 3.9 and, then, initiate sample acquisition.
6. Remove the sample tube from the machine 15–20 s after the start of the acquisition, and add 5 µL of undiluted YG beads. Return the sample FACS tube and continue the acquisition. Run the samples for 7–8 min at around 1,000 events/s.
7. While the first sample is running, take the second sample from the ice and place it in a 37 °C water bath to allow sufficient time for the cells to recover prior to analysis. Once the first sample is finished, clean the intake by running a water sample and, then, begin acquisition on the second sample as described in steps 3.8 and 3.9.

6. Data Analysis

1. Export the data to a spreadsheet. The data analysis will depend on the experimental question.
   NOTE: Be aware that different runs may have different baseline intensities, so it is important to run the assay for a designated time at the start (around 40 s) prior to adding any agonists and to normalize the data by calculating the change in fluorescence (the fluorescence at any given time point [F] divided by the fluorescence at time point zero [F0], or F/F0).
2. To quantify the rate or kinetics of the P2X7 function in question, calculate the area under the curve or the sum of the trapezoids created under the curve for each 10 s time period¹⁶.
3. To determine the effects of treatments, average the fluorescence intensity over the final 10–20 s of recording and compare the treatments. Determine the significance by t-test or analysis of variance.