Real-Time In Vitro Migration Assay for Primary Murine CD8+ T Cells

The animal protocols were approved by the University Committee on Animal Resources at the University of Rochester. The mice in this study were maintained in the pathogen-free space of the University of Rochester animal facility. Male/female C57BL/6 mice, aged 6-12 weeks (15-30 g), were used for the present study. Mouse tissue isolation can be performed on a benchtop with gloves to cover hands and a facemask to cover the nose and mouth, or inside a biosafety cabinet. All cell culture and plate preparation must be performed in a biosafety cabinet. The reagents and equipment used in this study are listed in the Table of Materials.

1. CD8⁺ T cell purification and activation

1. Preparation of materials
   1. For the negative selection of CD8⁺ T cells: Prepare a negative selection medium from the supernatant of hybridoma cell lines GK1.5 and M5/114 antibodies, which produce anti-CD4 and anti-MHCII, respectively. Mix the supernatant at a 1:1 ratio, filter it, and store it at 4 °C for future use (>0.5 µg of each antibody/million total cells). Alternatively, commercial CD8⁺ T cell isolation kits are available.
   2. For activation of T cells: Add 500 µL of CD3 (10 µg/mL) and CD28 (16 µg/mL) antibodies in 1x DPBS (without calcium and magnesium) to a non-tissue culture (TC) treated 24-well plate and store overnight at 4 °C to ensure plate binding.
      NOTE: Antibodies and proteins coat non-TC-treated culture plates/dishes better than regular TC-treated ones, so non-TC-treated culture plates are used for T cell generation.
2. Prepare a 10 cm culture dish (or equivalent) by placing a 70 µm cell strainer into the dish and dispense 2 mL of R9 medium into the filter (R9 medium: RPMI 1640x supplemented with 10% FBS, 1% antibiotic-antimycotic, 20 mM HEPES buffer, 1% MEM non-essential amino acids, 50 µM β-mercaptoethanol).
3. Obtain a mouse with appropriate genetic background, sex, age, and weight as determined by the experimental design.
4. Euthanize the mouse using CO₂ followed by cervical dislocation, or appropriate euthanasia strategies as defined by local animal care and use protocols and approved by the institution.
5. Place the mouse in a supine posture, stretch all four limbs perpendicular to the body, and pin footpads to a dissection board using mouse dissection T pins, or equivalent. Make a superficial, central incision through the cutaneous layer on the ventral lower abdomen with mouse surgical dissection scissors and cut straight up to the chin (~8 cm). Separate the skin from the peritoneal lining to expose lymph nodes. Stretch the skin perpendicularly away from the trunk and pin it down⁵.
   1. Gently excise the cervical, axial, brachial, and inguinal lymph nodes bilaterally using blunt forceps (or the preferred type from a standard mouse surgical dissection kit). Transfer to the cell strainer prepared in step 1.2.
   2. Open the peritoneum and remove the spleen. Transfer to the cell strainer prepared in step 1.2.
   3. Dispose of the mouse carcass in the appropriate biohazard receptacle.
6. Prepare a single-cell suspension of the secondary lymphoid tissues by mechanical disruption over the 70 µm cell strainer with 2 mL of R9 medium from step 1.2 by twisting the tissue disruption tool one-half turn clockwise and counterclockwise repeatedly.
   NOTE: The plunger end of a luer-lock syringe, either 3 mL or 10 mL, works well to crush and homogenize the tissue. Alternative materials can be used at the user's discretion.
7. Wash the cells, syringe end, strainer, and culture dish with 7 mL of medium, ensuring the strainer remains upright to prevent the transfer of larger pieces of tissue or cellular aggregates that may be present into the single-cell suspension.
8. Transfer the cell suspension to a 15 mL conical tube.
9. Pellet the cells at 270 x g at room temperature (20 °C) for 5 min and decant the supernatant.
10. Add 500 µL of lysing buffer for 1 min to remove red blood cells. Dilute with 9.5 mL of R9 medium and spin at 270 x g (20 °C) for 5 min. Decant the supernatant.
11. Resuspend the pellet in 10 mL of the previously prepared negative selection medium containing anti-MHCII and anti-CD4 (step 1.1.1). Rock at room temperature (20 °C) for 30 min.
12. Pellet the cells at 270 x g for 5 min and decant the supernatant. Wash 3x with 10 mL of R9 medium.
13. Simultaneously, in a 15 mL conical tube, wash 200 µL of sheep anti-rat IgG beads in 7 mL of medium. Place the tube on a magnet, remove the medium, and repeat the wash step twice. Resuspend the beads in 7 mL of R9 medium.
14. Pellet the cells at 270 x g (20 °C) for 5 min, decant the supernatant, and then resuspend the pellet in 7 mL of bead suspension from step 1.13. Rock at room temperature (20 °C) for 45 min.
15. Enrich CD8⁺ T cells by magnetic bead depletion. Place the conical tube directly on the magnet without a lid for 3 min to remove antibody/bead-bound cells. CD8⁺ cells should be retained. With the tube still in the magnet, collect the cell suspension using a serological pipet or equivalent and transfer it to a new 15 mL tube. Centrifuge at 270 x g (20 °C) for 5 min and wash 3x in medium.
16. Wash the pre-coated activation plate (step 1.1.2) with 1x DPBS twice without directly pipetting on the bottom surface of the plate.
17. Optional: Perform a live/dead lymphocyte separation to remove dead cells.
    1. Transfer cells to a 15 mL conical tube. Gently add 2 mL of lymphocyte separation media to the bottom of the cell suspension. Centrifuge at 900 x g for 10 min (at room temperature) at low acceleration and deceleration.
       NOTE: Live cells will separate at the interface of medium and separation media (middle white layer). Dead cells will pellet at the bottom of the tube and may be discarded at the end.
    2. Transfer the middle layer containing live lymphocytes to a new tube and centrifuge at 270 x g for 5 min at room temperature. Decant the supernatant.
18. Resuspend the pellet in 12 mL of medium with 10 U/mL recombinant mouse IL-2. Plate 1 mL per well in a non-TC treated 24-well plate and transfer to 37 °C overnight.

2. Lifting the activated CD8⁺ T cells

1. Visualize cells using a standard benchtop light microscope, using the 4x or 10x objective.
   NOTE: Activated T cells will appear larger and round or elongated compared to inactivated T cells and should have dense clusters of cells throughout the well.
2. To lift the activated cells, disrupt the cells with a 1 mL pipette, making sure to mix well at the edges of the plate. Transfer cells to a new 15 mL conical tube.
   NOTE: Activated cells are stickier and require more vigorous pipetting.
3. Wash the wells with 1 mL of R9 medium to remove any remaining cells. Repeat once more if needed. Centrifuge the cells at 270 x g (20 °C) for 5 min. Discard the supernatant.
4. Perform a live/dead lymphocyte separation to remove dead cells as in steps 1.17-1.18.
5. Maintain T cells by live/dead cell removal every 3-4 days in R9 medium with IL-2 in a non-TC 6-well plate.
   NOTE: T cells proliferate better in wells of small dimension during early activation, possibly due to T cell-T cell contacts or T cell-derived soluble factors that help their activation. Rapid T cell proliferation causes the culture medium to yellow within 24-48 h post-culture. At this point, lift the cells and transfer them to a larger well (uncoated, non-TC 6-well plate) with sufficient fresh culture medium to feed the cells (typically 5 mL per well). Replace depleted medium with fresh whenever indicated by yellow medium color change, or every 3-4 days.

3. Preparation of glass dish

1. Prepare glass cell migration chambers or plates by coating with 300 µL/cm² Protein A (0.1 mg/mL) overnight at 4 °C, wash twice with 1x DPBS by dispensing onto the dish and decanting before moving to the next step.
2. Coat the migration chamber or plate with 300 µL/cm² recombinant mouse ICAM-1 or VCAM-1 Fc chimera (0.1-2.75 mg/mL) together with a recombinant mouse chemokine (0.1-10 µg/mL) for 2-3 h at room temperature. Other integrin ligands such as fibronectin, mouse collagen IV, and mouse E-cadherin are rendered to directly coat the chamber at 2.5-10 µg/mL overnight at 4 °C.
   1. Wash chambers twice with 1x DPBS by dispensing onto the dish and decanting immediately after before adding cells.
      NOTE: T cells can migrate differently in different concentrations of integrin ligands and chemokines depending on the status of activation, differentiation, sensitization, and priming. Assaying cell migration in multiple different concentrations is strongly suggested.
3. Plate the adherent cells on glass-coated plates 24 h prior to the start of imaging if co-culturing T cells with other cells, such as cancer cells, to measure T cell killing of cancer cells in a real-time manner.

4. Preparation of cells

1. Optional: Stain T cells and adherent cells with cell tracker dye of choice at 1 µg/mL per 1 x 10⁷ cells in 1x DPBS for 15 min at 37 °C, or according to the manufacturer's protocol.
2. Count cells using a hemocytometer or equivalent.
3. Plate 1 x 10⁵ CD8⁺ T cells (or desired amount) in Leibovitz's L-15 medium supplemented with 1-5 g/L glucose in a 37 °C chamber.
   NOTE: L-15 medium is formulated for use without a CO₂-sodium bicarbonate system. It is buffered using free basic amino acids, phosphate buffers, and galactose to maintain pH levels.
4. Optional: Perform integrin blocking.
   1. Preincubate T cells with a blocking antibody to an integrin (1-10 µg/mL, anti-mouse CD11a (M17/4), anti-mouse CD18 (M18/2), anti-mouse VLA-4 (9C10) for 10 min and allow to crawl in the presence of the same antibody.

5. Time-lapse microscopy

1. Perform video microscopy.
2. For T cells, acquire brightfield and fluorescent images every 10-60 s for 10-60 min, or desired acquisition settings.
   NOTE: T cell migration is temperature-sensitive, and the optimal temperature for mouse T cell migration is 37 °C. The temperature control system that this protocol uses consists of an enclosed and insulated microscope deck with an attached heating system. The imaging chamber sits on the imaging deck. The perimeter of this chamber has a well filled with distilled water that allows consistent heat and humidity during imaging. The microwell is positioned at the center of the imaging chamber.

6. Software-assisted analysis of T cell migration

1. Use the Volocity software to assess T cell migration.
   1. Open Volocity and create a new image sequence.
   2. Select and move all time-lapse video microscopy movies to the blue area in Volocity.
   3. Adjust brightness and contrast using the contrast enhancement tool to the researcher's desired settings.
   4. Check image sizes: 0.325 µm for 20x, 0.65 µm for 10x.
   5. Sequence: set timepoints (ex: 81 images if taking an image every 15 s for 20 min, 4 per min).
   6. Uncheck all timepoints in the measurement tab.
   7. Find objects using intensity-slide the bar so it is just right of the peak.
   8. Exclude objects by size: all cells that are smaller than 10 µm diameter and greater than 100 µm to exclude cell debris or non-cellular particles. Exclude T cells that are migrating for less than 20% of the recording time (which can vary depending on the definition by each investigator).
   9. Ignore static objects (check box).
   10. Automatically join broken tracts (check box).
   11. Set the shortest path to no more than 20 µm.
   12. The track of a cell migration is the line connecting the location of the same cell over time. Ensure that the tracking algorithm is tracking by detecting where individual objects are detected by segmentation in each frame, and the objects are matched across frames.
   13. Save a new protocol by choosing Measurements > Save Protocol > Name New Protocol.
   14. Click on measure all timepoints in the measurement tab.
   15. Sort tracks by timespan, high to low.
   16. Record track ID numbers for all cells with good tracks as defined by the investigator. After automatic cell tracking, manual evaluation of each cell track is necessary to exclude falsely identified or broken tracks. Exclude cells with bad tracks.
   17. Export the file as comma-separated text and transfer data to the desired analysis software.
       NOTE: Basic parameters: Velocity (µm/min), displacement (net movement, µm), track length (total path length, µm), and meandering index (MI; net displacement/track length. MI = 1 indicates a completely straight linear track).
2. Perform manual tracking using Image J.
   1. Open plugins and select tracking and manual tracking. Set the parameters – time intervals, x/y calibration (pixel size), and z calibration (in manual tracking mode). Start individual cell tracking by clicking on add track, selecting one cell at the first time point, and continuing it through all time points. Next, click on End track. Repeat it for all the cells of interest.
      NOTE: Trackmate, another tool for cell tracking, is an automatic tracking software available via the ImageJ plugin.