Isolation and Flow Cytometric Analysis of Glioma-infiltrating Peripheral Blood Mononuclear Cells

Note: Please review the entire protocol prior to performing experiments. Approval for the use of vertebrate animals from the appropriate institutional committee on the use and welfare of animals must be obtained prior to proceeding.

1. Preparation of Tumor Cells for Intracranial Engraftment

1. Working in a class II biological safety cabinet, start by preparing GL26-Cit-NT/gal1i cell culture media by supplementing a 500 ml bottle of Dulbecco’s Modified Eagle Medium (DMEM) with 10% sterile-filtered heat-inactivated fetal bovine serum (FBS), 2 mM L-glutamine, 100 U/ml penicillin, 100 µg/ml streptomycin, 600 µg/ml G418 sulfate (for selection of the mCitrine expression vector) and 3 µg/ml of puromycin dihydrochloride (for selection of the non-targeting or gal-1-specific shRNAs).
2. Culture GL26-Cit-NT and/or GL26-Cit-gal1i cells (Figure 1A and 1B) in a tissue culture cabinet set to 37 °C and 5% CO₂ for 1-2 days prior to the tumor engraftment procedure or until the flasks reach 50-80% confluency.
3. On the day of surgery, remove the cell culture media from the glioma cells using a 10 ml serological pipette and a pipette gun and discard the media into a waste beaker.
   1. Invert the flask top-side-down and add 10 ml of DPBS to the topside of the flask using a 10 ml serological pipette. Slowly invert the flask to cover the cells in DPBS, then tip the flask vertically and remove and discard the DPBS into a waste beaker.
4. Add 3-4 ml of non-mammalian trypsin alternative in DPBS with EDTA (see materials list for details) to each T75 tissue culture flask and place back into the tissue culture cabinet for 2-5 min. Using a bright-field microscope, check that all of the cells have become detached from the bottom surface prior to proceeding.
5. Inhibit further enzymatic activity by diluting the trypsin alternative with 6 ml of DPBS. Pipette up and down using a 10 ml serological pipette to rinse the bottom surface of the flask and to mechanically dissociate any cellular aggregates. Aspirate the cells and place into respectively labeled 15 ml centrifuge tubes. Spin the cells down at 550 x g (max RCF) for 5 min at 4 °C (Figure 1C).
6. Remove the supernatant and gently resuspend the cells with 1 ml of un-supplemented DMEM. Make sure to pipette the cells up and down thoroughly with a P-1000 micropipette to achieve a single cell suspension.
7. Make a 1:10 dilution of the resultant cell suspension generated in 1.6 by placing 18 µl of un-supplemented DMEM into a 0.6 ml conical polypropylene microtube followed by the addition of 2 µl of the tumor cell suspension. Pipette up and down using a P-10 micropipette to homogenize the cells thoroughly.
8. Add 10 µl of the 1:10 tumor cell dilution generated in 1.7 to a separate 0.6 ml conical polypropylene microtube, then add 10 µl of Trypan blue stain to the tube. Thoroughly mix with a P-10 micropipette and add 10 µl of the resultant tumor cell/Trypan blue solution under a glass cover slip placed on top of a hemocytometer being careful not to introduce air bubbles (Figure 1D).
   1. Count the cells with a bright-field microscope using a 10X objective according to the specific protocol associated with the given hemocytometer. Be sure to factor in the 20-fold dilution of the original tumor cell suspension made during the preparation of the cells for accurate cell estimation in the original 1 ml aliquot. Use the following formula to calculate the total number of cells: total cells/ml = [((∑cells counted per square) / # of squares counted) x 20 (i.e., dilution factor) x 10,000 cells/ml].
9. After determining the total number of cells for each cell line used in the given experiment, spin them down at 550 x g (max RCF) for 5 min at 4 °C. Remove the supernatant(s) and resuspend the cells with an appropriate volume of un-supplemented DMEM in order to achieve a target concentration of 3 x 10⁴ cells per 1 µl for each cell line according to the following formula: (total # of cells / 30,000).
10. Place an equivalent volume of each cell line into respectively labeled 0.6 ml conical polypropylene tubes and place on ice (Figure 1E). Be sure to save some cells to continue propagating each cell line if separate T75s were not previously seeded.

2. Procedure for the Stereotactic Engraftment of Glioma Cells into the Striatum of C57BL/6J Mice

1. Prepare working solutions of surgical anesthesia, analgesia, and anesthesia reversal prior to surgery as follows:
   1. For ketamine/dexmedetomidine surgical anesthesia: remove 1.4 ml from a 10 ml vial of sterile 0.9% NaCl injection and replace with 600 µl of a 100 mg/ml stock solution of ketamine hydrochloride and 800 µl of a 0.5 mg/ml stock solution of dexmedetomidine hydrochloride to yield a mixed working solution of ketamine hydrochloride (6 mg/ml) and dexmedetomidine hydrochloride (0.04 mg/ml).
   2. For buprenorphine analgesia: remove 1 ml from a 10 ml vial of sterile 0.9% NaCl injection and replace with the entire 1 ml volume of a single 0.3 mg/ml stock buprenorphine hydrochloride ampul to yield a 0.03 mg/ml working solution.
   3. For atipamezole surgical anesthesia reversal: remove 1 ml from a 10 ml vial of sterile 0.9% NaCl injection and replace with 1 ml of a 5 mg/ml stock solution of atipamezole hydrochloride to yield a 0.5 mg/ml working solution.
2. Working in an approved location for mouse survival surgery, begin by setting up autoclaved or bead sterilized surgical instruments and other reagents as shown in (Figure 2A), then obtain enough 8-10 week-old female C57BL/6J mice (syngeneic with GL26 glioma) required for the study; ensure their continuous access to food and water.
3. Anesthetize the first mouse with a single intraperitoneal (i.p.) injection of the working anesthesia solution by delivering a dose of 75 mg/kg ketamine and 0.5 mg/kg dexmedetomidine (approximately 250 µl for a 20 g mouse). Next, give a 5 mg/kg subcutaneous (s.c.) injection of carprofen (approximately 100 µl for a 20 g mouse). Ensure that the mouse is non-responsive to toe and tail pinches prior to proceeding.
4. Using surgical clippers, shave the fur from the cranium. Apply povidone-iodine antiseptic solution to the shaved area, scrub with a 70% isopropyl alcohol prep pad, then re-apply povidone-iodine antiseptic solution. Next, apply a small amount of sterile petrolatum ophthalmic ointment to each eye to prevent drying.
5. Secure the skull in the stereotactic frame according to the following protocol:
   1. Carefully open the mouth by reaching in with a pair of curved dissecting forceps to gently pull out the tongue and move to one side of the mouth to prevent choking. Allow the forceps to retract while in the mouth to spread the jaw open and guide the top two incisors into the keyhole of the tooth bar on the stereotactic frame. Secure the tooth bar from pivoting horizontally or vertically by adjusting the respective screws. Ensure that the mouse’s cranium is level with the bench top.
   2. Place the ear bars securely against the postorbital bones of the cranium being careful not to apply too much inward pressure on the skull in order to prevent crushing.
   3. Place the nose bar over the snout and screw down until snug. Check that there is no vertical/lateral movement in the head by applying gentle pressure with a thumb and index finger. A mouse properly placed into a stereotactic frame is shown in (Figure 2B).
   4. Using a scalpel blade, make a 1 cm midline incision over the cranium from the frontal bone to the occipital bone. Retract the skin at the incision using Colibri retractors.
   5. Lower the microliter syringe equipped with a 33 G needle attached to the vertical arm of the stereotactic frame directly above the position of bregma (Figure 2C). From there, adjust the position of the needle +0.5 mm AP, +2.5 mm ML to reach the target site for tumor implantation. Use a 1 ml tuberculin syringe equipped with a 26 G needle to mark this location by gently excoriating the periosteum.
   6. Drill a hole into the cranium at the target site until reaching the underlying dura mater using a cordless precision power drill equipped with a 1/32” (0.8 mm) bit. While drilling, apply intermittent pressure followed by removal of the drill bit from the surface of the cranium to avoid excessive heat and force that may otherwise lead to the drill bit puncturing the underlying brain tissue.
6. Flush the microliter syringe with 0.9% NaCl in a 1.7 ml conical polypropylene microtube to ensure that the needle is not clogged. Gently flick the 0.6 ml conical polypropylene microtube containing the tumor cells several times to resuspend the cells.
7. Draw up 2 µl of cells into the microliter syringe ensuring that no air bubbles are introduced into the syringe. Dispense 1 µl of the cells onto a 70% isopropyl alcohol prep pad leaving exactly 1 µl of tumor cells remaining in the syringe.
8. Bring the needle down until it touches the dura mater, then introduce the needle into the brain -3.5 mm ventrally and retract by +0.5 mm. Slowly and smoothly deliver the cells by depressing the syringe plunger over the course of 1-2 min.
9. Allow the cells to settle at the injection site for 5 min prior to slowly withdrawing the needle. Wipe any coagulated blood or brain matter from the tip of the needle with a clean 70% isopropyl alcohol prep pad. Rinse the needle and syringe thoroughly with the 0.9% NaCl from step 2.6 to ensure that the needle is not clogged for the next injection.
10. Remove the Colibri retractors and suture the scalp using three 3-0 nylon monofilament sutures. Remove the mouse from the stereotactic frame and administer 0.1 mg/kg of buprenorphine hydrochloride subcutaneously (s.c.) (approximately 67 µl of the 0.03 mg/ml working solution for a 20 g mouse) followed by 2.5 mg/kg atipamezole hydrochloride intramuscularly (i.m.) (approximately 100 µl of the 0.5 mg/ml working solution for a 20 g mouse) for post-operative pain relief and anesthesia reversal. Place post-surgical mice back into their cages under a heating lamp to recover.

3. Mouse Transcardial Perfusion and Harvesting of the Brain

1. Prepare heparinized Tyrode’s Solution according to the following protocol:
   1. Put a magnetic stirring bar into a 1 L glass screw-cap storage bottle and fill to volume with ultrapure deionized water. Place the bottle on a stirring plate.
   2. Weigh and add the following chemicals to the glass bottle while stirring: 8 g sodium chloride (NaCl), 0.264 g calcium chloride (CaCl₂•2H₂O), 0.05 g Sodium phosphate monobasic (NaH₂PO₄•2H₂O), 1.0 g D-glucose (C₆H₁₂O₆), 1.0 g sodium bicarbonate (NaHCO₃), 0.2 g potassium chloride (KCl). Allow the salts to dissolve, then add 100 µl of a 1,000U/ml stock solution of heparin sodium to the Tyrode’s solution.
   3. Remove the container from the stirring plate and extract the stirring bar using a magnetic wand. Store heparinized Tyrode’s solution at 4 °C.
2. Prepare a working solution for terminal anesthesia as follows:
   1. For ketamine/xylazine terminal anesthesia: remove 1,360 µl from a 10 ml vial of sterile 0.9% NaCl injection and replace with 1,200 µl of a 100 mg/ml stock solution of ketamine hydrochloride and 160 µl of a 100 mg/ml stock solution of xylazine hydrochloride to yield a mixed working solution of ketamine hydrochloride (12 mg/ml) and xylazine hydrochloride (1.6 mg/ml).
3. Prepare density centrifugation media mix solution by placing 90 ml of 10x PBS into 264 ml of ultrapure deionized water (3.93x) in a sterile plastic container. Titrate to pH 7.0-7.2 with HCl and filter sterilize through a 0.22 µm filter. Store at 4 °C.
4. Prepare 70% density centrifugation media by combining 18 ml of density centrifugation media mix solution with 30 ml of density centrifugation media (see materials list for details) in a 50 ml polypropylene centrifuge tube. Store at 4 °C, but bring to RT before use.
5. Prepare 37% density centrifugation media by combining 9.6 ml of DPBS with 10.4 ml 70% density centrifugation media in a 50 ml centrifuge tube. Store at 4 °C, but bring to RT before use.
6. Prepare flow buffer by placing 500 µl of FBS into 50 ml of DPBS (~1% FBS) in a 50 ml centrifuge tube. Store at store at 4 °C
7. Fill a 4.5 L polyurethane ice bucket to capacity with ice chips.
8. Make a combined solution of 1 mg/ml collagenase and 1 mg/ml DNase-I in sufficient quantity for 1 ml per brain sample in the study by diluting a 10 mg/ml DNase-I stock solution 1:10 and a 50 mg/ml collagenase stock solution 1:50 with sterile DPBS in a single 15 ml polypropylene centrifuge tube. Gently mix the solution by pipetting up and down with a P-1000 micropipette. Store on ice in the polyurethane bucket from step 3.7.
9. Obtain two 7 ml glass Dounce tissue grinders. Label one “NT” and the other “gal1i”. Place the tissue grinders in the ice bucket and add 1 ml of DPBS to each.
10. Obtain enough 15 ml centrifuge tubes to collect brain matter from each mouse in the study and place in the ice bucket.
11. Place ~150 ml of DPBS among three 50 ml centrifuge tubes into the ice bucket.
12. Obtain enough polystyrene hexagonal weighing dishes (top inner diameter (I.D.) 115 mm, base I.D. 85 mm, 203 ml volume) for each mouse in the study. The complete set up for dissection and trituration of mouse brains is shown in (Figure 3A)
13. At chosen time points, anesthetize the first tumor-bearing mouse in the study using a single i.p. injection of 150 mg/kg of ketamine hydrochloride and 20 mg/kg xylazine hydrochloride (approximately 250 µl of the combined 12 mg/ml ketamine/1.6 mg/ml xylazine working solution for a 20 g mouse) to induce deep anesthesia. Ensure that the mouse is non-response to toe and tail pinches prior to proceeding.
14. Retrieve the previously prepared Tyrode’s solution and place into a laminar flow hood dedicated to terminal animal surgical procedures. Place gas impermeable polymeric tubing attached to a carbogen tank into the Tyrode’s solution. Open the tank valve to allow 95% O₂/5% CO₂ carbogen to continuously bubble into the solution.
15. Attach a 20 G aluminum hub blunt needle to the end of additional gas impermeable polymeric tubing attached to a peristaltic pump. Place the tubing at the other end of the peristaltic pump into the Tyrode’s solution and turn the pump on to allow the tubing to fill with oxygenated and heparinized Tyrode’s solution. The set up for mouse transcardial perfusion is shown in (Figure 3B).
16. Using four 26 G needles, pin down the anesthetized mouse ventral side up by the front and hind paws on an extruded polystyrene foam block covered with a disposal absorbent towel in the laminar flow hood.
17. Grasp the skin above the peritoneal cavity using a pair of blunt dissection forceps, and using a large pair of dissection scissors make a “Y” incision by penetrating the peritoneal wall, cutting cephalically to puncture the diaphragm, then bifurcating at the sternum to terminate at the left and right axillae.
18. Use a hemostat to clamp the sternum and reflect the rib cage over the mouse’s left shoulder. Remove the pericardium with the pair of blunt dissection forceps.
19. Turn on the peristaltic pump to begin a steady drip of oxygenated and heparinized Tyrode’s solution (roughly 2.3-2.5 ml/min).
20. Insert the blunt needle into the left ventricle of the heart. Use a small pair of dissection scissors to snip the right atrium to allow for exsanguination (Figure 3C).
21. Allow the Tyrode’s solution to thoroughly perfuse the mouse circulatory system until the liver and lungs have completely blanched due to the lack of blood. Ensure that no gross amounts of blood continue to exit the right atrium prior to removing the 20 G aluminum hub blunt needle from the left ventricle.
22. Unpin the mouse from the extruded polystyrene foam block and turn the mouse ventral side down. Using a large pair of dissection scissors, separate the head from the rest of the body at the level of the neck.
23. Using a small pair of dissection scissors, cut the scalp at the midline beginning at the occipital bone working forward towards the snout in order to expose the cranium.
24. Retract the skin on both sides of the cranium using a thumb and index finger and hold the skull securely. Use a pair of bone Rongeurs to break through the cranium beginning at the occipital bone and work forward to completely expose the dorsal and lateral surfaces of the brain. Be careful to minimize damage to the brain.
25. Once the cranial bones have been removed, turn the head of the mouse ventral side up and use a small pair of dissection scissors to cut the cranial nerves at the base of the brain in order to liberate it from the skull.

4. Isolation of Glioma-infiltrating PBMCs

1. Using a clean single edged razor blade, isolate the area of the brain containing the tumor by making a sagittal cut down the center of the brain to bisect the two hemispheres. Turn the ipsilateral hemisphere medial side down and make two coronal cuts at the levels of the cerebellum and the olfactory bulb to isolate the target tissue containing the tumor implant (Figure 4A). An equivalent portion of the contralateral hemisphere may also be isolated and used as a negative control.
2. Place the target tissue into the respectively labeled glass Dounce tissue grinder containing 1 ml of DPBS, push the plunger all the way down and twist 7 times to initially disrupt the tissue. Lift the plunger to allow the liquid to settle back to the bottom of the tissue grinder. Repeat this step thrice; however only twist the plunger 4 times during the next two repeats and 3 times during the last repeat to avoid over triturating the tissue.
3. Using a P-1000 micropipette, sequentially apply three 1 ml volumes of ice-cold DPBS (prepared in 3.11) along the sides of the plunger to rinse into the tissue grinder.
4. Resuspend the triturated brain matter by pipetting up and down and place into a labeled 15 ml centrifuge tube on ice. Rinse the sides of the Dounce tissue grinder with 1 additional ml of ice-cold DPBS and add to the same 15 ml centrifuge tube. Keep all tubes containing triturated brain matter on ice until all samples have been processed.
5. Repeat steps 3.13-3.25 and 4.1-4.4 for each mouse in the study.
6. Once all samples have been processed, spin down the triturated brain matter in the 15 ml centrifuge tubes at 740 x g (max RCF) for 20 min at 4 °C.
7. Remove the supernatant with a 10 ml serological pipette and pipette gun and resuspend the pelleted brain matter in 1 ml of the previously prepared collagenase/DNase-I digestive enzymes using a P-1000. Place the 15 ml centrifuge tubes into a test tube rack and place in a 37 °C water bath for 15 min.
8. Gently agitate the samples twice throughout the incubation period by flicking the tubes to facilitate tissue disaggregation.
9. Add 6 ml of ice-cold DPBS using a 10 ml serological pipette to each tube to dilute the digestive enzymes. Pipette up and down to resuspend, and filter the total volumes through sterile 70 µm nylon mesh filters into new labeled 15 ml centrifuge tubes on ice.
10. Spin the brain cell suspension down at 740 x g (max RCF) for 20 min at 4 °C to achieve a single cell pellet (Figure 4B). After removing the 15 ml tubes from the centrifuge place them on ice and increase the temperature setting on the centrifuge to about 21 °C in preparation for the next step.
11. Fully remove the supernatant from each tube containing brain cells and initially resuspend the pellets in 1 ml of 70% density centrifugation media using a P-1000 micropipette. Then add 4 additional milliliters of 70% density centrifugation media to each tube using a 10 ml serological pipette. Screw the caps on securely and homogenize the cell suspension by gently inverting to tubes several times.
12. Remove the 15 ml centrifuge tubes containing brain cells resuspended in 70% density centrifugation media from ice, and place in a test tube rack at RT. One at a time, carefully overlay 2 ml of 37% density centrifugation media solution onto the 5 ml of 70% density centrifugation media with a P-1000 micropipette to form a clean interface between the two density centrifugation media layers (Figure 4C).
    1. Use a fine-tipped marker to indicate the position of the interface so it can be easily identified after centrifugation, when the distinction becomes less apparent.
13. Spin down the 15 ml centrifuge tubes at 740 x g (max RCF) for 20 min at RT with no break to avoid disrupting the interface.
14. After centrifugation, collect the PBMCs that have accumulated at the interface between the two density centrifugation media layers (Figure 4D) by introducing a P-200 micropipette into the tube along its side, carefully bypassing the lipid layer.
    1. Once at the level of the PBMC band, slowly extract 200 µl from the surface of the 70% density centrifugation media layer and place into a respectively labeled polypropylene FACS tube on ice. Repeat once for a total volume of 400 µl per sample.
15. Add 3 ml of flow buffer to each FACS tube containing 400 µl of PBMCs to sufficiently dilute the density centrifugation media, then spin the cells down at 660 x g (max RCF) for 20 min at 4 °C.

5. Immunolabeling of Glioma-infiltrating PBMCs

1. Prior to analyzing any experimental PBMC samples, perform a single, independent, cell surface isotype control experiment to establish baseline gates to be associated with a set of fixed PMT voltages within a flow cytometric data analysis template dedicated to the assessment of glioma-infiltrating PBMCs according to the following protocol:
   1. Engraft one C57BL/6J mouse with GL26-Cit-gal1i and another with GL26-Cit-NT according to the procedure outlined in section 2. After 72 hr of tumor growth, euthanize both mice and isolate the tumor-infiltrating PBMCs according to the procedures outlined throughout sections 3 and 4.
   2. Combine the two PBMC samples (NT and gal1i) into one volume (i.e., 100 µl), then further divide the sample evenly among three FACS tubes (i.e., 33.3 µl per tube) labeled “CD45 isotype”, “Gr-1/CD11b isotype”, and “NK1.1 isotype”. Add the following antibodies (each at a 1:100 dilution and diluted to a total volume of 200 µl in flow buffer) to the respectively labeled FACS tubes: “CD45 isotype”: Alexa Fluor 700-conjugated rat IgG2b, κ (clone: RTK4530); “Gr-1/CD11b isotype”: Alexa Fluor 700-conjugated rat anti-mouse CD45 (clone: 30-F11), PE-conjugated rat IgG2b, κ (clone: eB149/10H5), and PerCP/Cy5.5-conjugated rat IgG2b, κ (clone: RTK4530); “NK1.1 isotype”: Alexa Fluor 700-conjugated rat anti-mouse CD45 (clone: 30-F11), PE-conjugated rat anti-mouse Gr-1 (clone:RB6-8C5), PerCP/Cy5.5-conjugated rat anti-mouse CD11b (clone: M1/70), and APC-conjugated mouse IgG2a, κ (clone: eBM2a).
   3. Allow PBMCs to immunolabel on ice in the dark for 20 min. Flick the tubes once halfway through the incubation period to gently mix. Wash with 1 ml of flow buffer, spin down at 660 x g (max RCF) for 10 min at 4 °C and resuspend each FACS tube containing PBMCs with 200 µl of flow buffer.
   4. Using a flow cytometer equipped with an 85 µm integrated nozzle, analyze the “CD45 isotype“ tube first to establish a baseline CD45 interval gate. Next, run the “Gr-1/CD11b isotype” tube to establish a baseline Gr-1/CD11b quadrant gate using only true CD45⁺ cells as input. Finally, run the “NK1.1 isotype” tube to establish a baseline NK1.1 interval gate using only true CD45⁺/Gr-1^low/CD11b^+/- cells as input.
2. Use the following procedure for all future experimental analysis:
   1. Make a sufficient quantity of a 1:100 dilution of cell surface antibodies in flow buffer to achieve a 200 µl volume per sample (plus 1 for the GzmB isotype): Alexa Fluor 700-conjugated rat anti-mouse CD45 (clone: 30-F11), PE-conjugated rat anti-mouse Gr-1 (clone:RB6-8C5), PerCP/Cy5.5-conjugated rat anti-mouse CD11b (clone: M1/70), APC-conjugated mouse anti-mouse NK1.1 (clone: PK136).
   2. Carefully remove the supernatant of the spun down PBMCs from 4.15 until the meniscus is just above the bottom of each FACS tube (~50 µl) to avoid cell losses. Resuspend the PBMCs in each FACS tube using the residual flow buffer with a P-200 micropipette and remove (1/# of samples) worth of the volume from each FACS tube and combine into a new FACS tube labeled “pooled isotype”.
   3. Add 200 µl of the cell surface antibody cocktail prepared in 5.2.1 to each PBMC sample. Allow PBMCs to immunolabel on ice in the dark for 20 min. Flick the tubes once halfway through the incubation period to gently mix.
   4. Wash with 1 ml of flow buffer; spin down at 660 x g (max RCF) x 10 min at 4 °C.
   5. Resuspend all FACS tubes containing PBMCs in 200 µl of a commercially available formalin-based fixative (see materials list for details) for 15 min on ice in the dark. Flick the tubes once halfway through the incubation period to gently mix.
   6. Wash with 1 ml of a commercially available 1x permeabilization/wash buffer (see materials list for details) and spin down at 660 x g (max RCF) for 10 min at 4 °C.
   7. While PBMCs are spinning down, prepare a sufficient quantity of a 1:100 dilution of Pacific Blue-conjugated mouse anti-mouse GzmB (Clone: GB11) intracellular antibodies in 1x permeabilization/wash buffer to achieve a 200 µl volume per sample.
   8. In a separate tube, prepare a single 200 µl volume of a 1:100 dilution of Pacific Blue-conjugated mouse IgG1, κ (clone: MOPC-21) isotype control antibody.
   9. Resuspend each of the experimental PBMC samples with 200 µl of the 1:100 dilution of anti-mouse GzmB antibodies and resuspend the single FACS tube labeled “pooled isotype” with the 200 µl volume of the 1:100 dilution of isotype antibodies.
   10. Allow all PBMC samples to immunolabel on ice in the dark for 20 min. Flick the tubes once halfway through the incubation period to gently mix.
   11. Wash with 1 ml of flow buffer; spin down at 660 x g (max RCF) for 10 min at 4 °C and resuspend PBMCs with 200 µl of flow buffer for analysis.
   12. Flow cytometrically analyze glioma-infiltrating PBMCs using an 85 µm integrated nozzle and the gates and PMT voltages previously established in section 5.1²². Be sure to run the entire volume of each sample on the flow cytometer to ensure a fair comparison of the total number of glioma-infiltrating PBMCs in each sample.