Measuring Alloreactivity in a Mixed Population of T Cells

1. Prepare Reagents and Materials Required

1. Prepare phosphate-buffered saline (PBS) containing 2% fetal bovine serum (FBS) ("wash buffer"). Prepare PBS with 2% fetal calf serum (FCS) containing 0.1% nonionic detergent ("perm-wash buffer"; see the Table of Materials). Prepare PBS with 1.5% formaldehyde.
   NOTE: Formaldehyde is corrosive and potentially carcinogenic and must be handled while wearing appropriate personal protective equipment.
2. Prepare PBS containing 2% FBS and 0.5 mM ethylenediaminetetraacetic acid (EDTA) for magnetic cell separation ("MCS buffer").
3. Prepare 50 µg/mL phalloidin-fluorescein isothiocyanate (Phalloidin-FITC) in dimethyl sulfoxide (DMSO). Prepare a 1 mg/mL nuclear stain (e.g. 1 mg/mL 7-amino actinomycin D (7-AAD) in DMSO or bis-benzimide dye; see the Table of Materials) in DMSO. Prepare fluorochrome-labeled antibodies appropriate for the cells of interest and the imaging flow cytometer.
4. Obtain animal tissues (e.g., lymph nodes and spleen) as a source of T cells and allogeneic animal tissues (e.g., spleen and bone marrow) as a source of antigen-presenting cells or progenitors.
5. Prepare cell culture medium (e.g., Roswell Park Memorial Institute medium (RPMI) 1640 or Dulbecco's Modified Eagle Medium (DMEM) supplemented with 10% FBS), 50 µM 2-mercaptoethanol, penicillin, and streptomycin and obtain 24- and/or 96-well cell culture plates.

2. Prepare Antigen-presenting Cells (APCs)

NOTE: In theory, any APC population could be examined with this method. Immature mouse bone marrow-derived dendritic cells (DCs) as APCs were sued in this case. Many protocols exist for generating these cells. Briefly, the following protocol was used.

1. Flush marrow from femurs and tibias into RPMI 1640 or DMEM.
2. Pass the suspended cells through a 70-µm cell strainer to remove small pieces of bone and debris.
3. Pellet the cells by centrifugation and then lyse red cells using an ammonium chloride buffer for 5 min at room temperature.
4. Pellet the cells by centrifugation (400 x g, 5 min) and resuspend the cell pellet.
5. Wash cells in 5 – 10 mL of wash buffer, pellet by centrifugation (400 x g, 5 min), and re-suspend the cell pellet.
6. Enrich hematopoietic precursors over a cell separation column by labeling the cells with biotinylated anti-CD3 (5 µg/mL), anti-B220 (5 µg/mL), anti-MHC class II (1 µg/mL), and anti-CD11b (5 µg/mL) antibodies.
7. Pellet the cells by centrifugation (400 x g, 5 min) and re-suspend the cell pellet.
8. Incubate the cells with anti-biotin magnetic microbeads (see the Table of Materials) at 4 °C for 10 min.
9. Wash and pellet the cells (400 x g, 5 min) and re-suspend them in 1 mL of MCS buffer before removing the labeled cells using a large positive selection (LS) magnetic cell separation column primed with 3 mL of MCS buffer and placed in its magnet. Wash the column 3 times with 3 mL of MCS buffer; the flow-through will contain the desired cells.
10. Culture the cells that pass through the column for 6 days in RPMI 1640 or DMEM supplemented with 2 ng/mL recombinant mouse granulocyte-macrophage colony-stimulating factor (GM-CSF) and 2 ng/mL of recombinant human transforming growth factor β1 (TGFβ1). Replace half the medium every 2 days with a fresh complete medium containing 2 ng/mL GM-CSF and TGFβ1. 
    NOTE: Human TGFβ1 has activity in mouse cells. Data have been generated using these immature DCs. Other cells (e.g., B cells and mature DCs) may be suitable as APCs but have not been tested in this assay.
11. Cryopreserve DCs in 90% serum/10% DMSO and store in liquid nitrogen; recover on the day of use. Prior to use, count the number of viable DCs in a hemocytometer using trypan blue exclusion. Re-suspend the cell pellet in culture medium at the appropriate density (see step 4.1) prior to use in section 4.

3. Prepare T Cells

1. Use negative selection methods to avoid inadvertently transmitting activating or inhibitory signals to the cells.
   NOTE: In this example, CD4⁺ T cells are prepared for analysis.
   1. To prepare CD4⁺ T cells from the mouse spleen, mash the spleen through a 70-µm cell strainer using the plunger of a syringe. Wash the cell strainer with wash buffer.
   2. Pellet the suspended cells by centrifugation (400 x g, 5 min) and then lyse the red cells by re-suspending the pellet in an ammonium chloride buffer for 5 min at room temperature.
   3. Pellet the cells by centrifugation (400 x g, 5 min) and re-suspend the pellet.
   4. Wash the cells in 5 – 10 mL of wash buffer and pellet by centrifugation (400 x g, 5 min). Re-suspend the pellet.
   5. Stain the cells with biotinylated antibodies to CD8, major histocompatibility complex class II (MHC II, 1 µg/mL), and CD19 (5 µg/mL). Incubate for 10 min at 4 °C.
   6. Wash the cells in 10 mL of wash buffer and pellet by centrifugation (400 x g, 5 min). Re-suspend the pellet.
   7. Incubate the cells with anti-biotin magnetic microbeads (see the Table of Materials) according to the manufacturer's directions.
   8. Wash the cells in 10 mL of wash buffer and pellet by centrifugation (400 x g, 5 min); re-suspend the pellet.
   9. Resuspend the cells in 1 mL of MCS buffer and enrich the CD4⁺ T cells over a magnetic cell separation column primed with 3 mL of MCS buffer on a magnet. Wash the column 3 times with 3 mL of MCS buffer. Column flow-through will contain the enriched T cells.
2. By standard flow cytometry, assess T cell purity using an aliquot of the negatively selected cells. Stain the cells using a fluorochrome-streptavidin conjugate (to identify any biotin-labeled cells that should have been removed on the column) and an antibody or antibodies to identify the T-cell population of interest (CD4 in this case); a purity of ≥85% is acceptable.
   1. Count the T cells in a hemocytometer by trypan blue exclusion (≥90% viability is acceptable).
      NOTE: MCS buffer contains EDTA, which must be removed prior to the assay. To accomplish this, pellet the cells by centrifugation (400 x g, 5 min) and wash them in 1 mL of wash buffer. Pellet the cells again (400 x g, 5 min) and re-suspend in culture medium at the appropriate density (see step 4.1).

4. Co-incubate T Cells and DCs

1. Seed T cells and DCs at a 2:1 T: DC ratio in a 24-well or 96-well cell culture plate. Ensure that the final culture volume is ≤500 µL for 24-well plates or ≤50 µL for 96-well plates.  
   NOTE: Smaller volumes encourage cell-cell interactions and allow space for subsequent fixation buffers.
   1. Adjust precise cell numbers empirically, but as a general guide, use 1 x 10⁶ T cells and 0.5 x 10⁶ DCs per well (96-well plate). These should be considered the minimum numbers of cells because using fewer cells makes the enumeration of immune synapses difficult.
      1. To increase cell numbers, set up replicate wells and pool after step 5 (fixation).
         NOTE: When setting up replicate wells, it is advisable to seed DCs in all of the wells first and then seed T cells in all of the wells; this minimizes discrepancies in incubation time between wells.
2. Incubate the plate for 4 h at 37 °C in a 5% CO₂ atmosphere.

5. Fix Cells in Plate

1. Add 3 times the culture volume of 1.5% formaldehyde in PBS to each well and incubate at room temperature for 30 min; it is important to fix cells prior to removing them from the plate to minimize the disruption of cell-cell interactions.
2. Transfer cells in the plate into tubes for subsequent washing and staining. At this stage, set aside additional cells for single-stain controls. Apart from these controls, the entire culture should be stained with the antibody cocktail (see step 4.1.1).

6. Stain Cells

1. Stain cells in 100 µL of wash buffer containing a cocktail of the desired fluorochrome-conjugated antibodies for 30 min at room temperature, protected from light.
   NOTE: The cocktail includes T cell-specific and APC-specific antibodies. Fluorochromes should be chosen such that they can be distinguished using the configuration of the imaging flow cytometer. In the experiments shown here, blue fluorophore-conjugated CD11b (5 µg/mL, see the Table of Materials) and APC-conjugated CD90.2 (5 µg/mL) were used.
2. Wash the cells in 1 mL of wash buffer and centrifuge for 5 min at 400 x g. Decant the supernatant. Resuspend the cells in perm-wash buffer containing phalloidin FITC at 0.05-0.5 µg/mL and incubate for 30 min at room temperature protected from light.         
   NOTE: Phalloidin FITC concentrations of about 0.1 µg/mL work well for mouse cells, but the appropriate concentration is expected to vary by supplier, cell type, and imaging flow cytometer.
3. Wash the cells in 1 mL of perm/wash buffer and centrifuge for 5 min at 400 x g. Decant the supernatant. Resuspend the cells in perm/wash buffer containing nuclear dye at the appropriate concentration (e.g., approximately 25 µg/mL 7-AAD) and incubate for 30 min at room temperature protected from light.
4. Wash the cells in 1 mL of perm/wash buffer and centrifuge for 5 min at 400 x g. Decant the supernatant. Wash the cells once in wash buffer, pellet, and re-suspend in 50 – 100 µL of wash buffer, transferring the cells to small, capped microcentrifuge tubes.
5. Proceed to data acquisition immediately or store the cells at 4 °C protected from light for up to several days prior to acquisition on the imaging flow cytometer.
   NOTE: Cells have been successfully stored in this way for up to 7 days. Longer storage may be possible but has not been tested.

7. Acquire Data

1. Initialize and configure the imaging flow cytometer according to the manufacturer's instructions. Ensure stability of the flow core prior to collecting any data.
2. Reserve one channel for brightfield image acquisition. Acquire single-stain control data with the brightfield channel turned off.
   1. Click the "Load" button and insert a tube containing a fully stained sample (a sample that has been stained with all required fluorochromes) into the holder.
   2. In the "workspace" window, select and create a new scatterplot with the aspect ratio over the area and gate singlets where the aspect ratio is close to 1. Create a new scatterplot for each channel used (intensity of the channel in the horizontal axis).
   3. For each fluorochrome, check the positive population and, if required, adjust the laser voltage in the "Illumination" box.
   4. Unload the tube and load the first single-stained tube.
   5. In the "Acquisition Settings" box, type the sample name and set the number of events that should be collected; if it is a single stain (for compensation controls), 1,000 – 2,000 events are sufficient.
   6. In the "Channels" box, select the channels that each sample has been stained with. For single-stain controls, all channels should be selected with brightfield and side scatter off. Click the "Record" button under the "Acquisition" box; when the number of events reaches the specified threshold, the acquisition will stop automatically.
   7. Click the "Return" button to unload the tube. Repeat steps 7.2.4 – 7.2.6 for each single-stain control. Depending on the cytometer and software, it may not be possible to set all of the gates shown in Figure 1 during acquisition (more precise gating is performed during analysis; see section 8).
3. Acquire samples as for single-stained samples (in step 7.2), but in the "Channels" box, select all channels that are required, including the brightfield channel.
   1. Before recording data, check to confirm that the channel intensity is appropriate for identifying the desired cell populations. If not, adjust the laser settings and re-record single-stain controls using the new settings, as described in step 7.2.
   2. For each sample, acquire several tens of thousands of events.
      NOTE: Under most conditions, cell-cell contact events are a small minority of the total cell number (most are single cells). In general, it is desirable to have at least 100 events in the final membrane contact gate.