Isolation of Dendritic Cells from the Human Female Reproductive Tract for Phenotypical and Functional Studies

Studies involving human subjects were conducted according to the principles expressed in the Declaration of Helsinki. Studies were approved by Dartmouth College Institutional Review Board and the Committee for the Protection of Human Subjects (CPHS). Written informed consent was obtained before surgery from HIV-negative women undergoing hysterectomies at Dartmouth-Hitchcock Medical Center (Lebanon, NH). Trained pathologists selected tissue samples from the EM, CX, and ECX, free of pathological lesions and distant from the sites of pathology. Blood samples were obtained from volunteer healthy donors recruited at Dartmouth Hitchcock Medical Center. Blood donors were anonymous. Freshly isolated EM, CX, and ECX tissues from the operating room were transferred to Pathology for isolation and classification prior to transfer to our laboratory in separate, sterile polypropylene tubes.

1. Enzymatic Digestion of Tissues

NOTE: Use sterile materials and work in a biological safety cabinet.

1. Prepare a modified Hank's Balanced Salt Solution (HBSS) using 1x HBSS with phenol red, 100 U/mL penicillin-streptomycin, 0.35 g/L NaHCO₃, and 20 mM HEPES.
2. Prepare the enzymatic cocktail (modified HBSS, 450 U/mL collagenase IV, 0.01% [wt/vol] deoxyribonuclease I, and 2 mg/mL D-glucose). Pass the solution through a sterile 0.22 µm filter. Ideally, prepare the digestion enzyme on the day of use, but it can be frozen at -20 °C for storage. Avoid repeated freezing and thawing cycles.
3. Rinse the tissue with modified HBSS and place in a 100 x 15 mm² Petri dish. Adjust the Petri dish size based on the amount of tissue available and the volume of enzyme cocktail used (see step 1.4.2 below).
   NOTE: Tissues vary in size depending on the surgical sample obtained from the pathology, ranging from 1-10 g.
4. Physical processing with disposable scalpel and forceps:
   1. Add approximately 3 mL of digestion enzyme cocktail to the tissue to prevent drying while cutting.
   2. Use forceps to stabilize the tissue and mince with a scalpel to increase the surface area of the tissue exposed to the digestion enzyme cocktail. Cut the tissue into small pieces (< 2 mm on a side). Add sufficient digestion enzyme cocktail to cover all tissue pieces (5 mL/g of tissue). Place the lid on the Petri dish.
   3. Incubate at 37 °C with 5% CO₂ on a rotator at approximately 80 rpm for 45 min. Ensure that the rotator speed thoroughly mixes the digestion enzyme cocktail without spilling or producing bubbles.
   4. Visualize the tissue preparation with an inverted light microscope with the 4X and 10X objectives.
      NOTE: After digestion, small tissue fragments and a mixed cell suspension including visible epithelial glands should be present (see Figure 1A).

2. Physical Separation of Single Cells

1. In a sterile environment, place a taut 250 µm mesh with holder into a 150 x 15 mm² Petri dish. Ensure that the mesh is about 0.5 cm above the surface of the Petri dish.
2. Wet the mesh with 2 mL of modified HBSS and transfer the digested tissue onto the mesh.
3. Using the flat surface of a 10-mL syringe plunger, gently but firmly, grind the digested minced tissue through the mesh.
   Caution: Grinding too aggressively will damage the mesh and increase cell mortality. This step will separate the epithelial cells and stromal cells (including immune cells) from the connective tissue and mucus.
4. Once the tissue fragments have been thoroughly dispersed, elevate the mesh and the holder 2-3 cm above the Petri dish and rinse with 3 mL of modified HBSS to recover additional cells.
5. Collect the cell suspension. Place the 20 µm mesh with holder in a Petri dish and wet with 2 mL of modified HBSS. Pour the cell suspension through the mesh holding the mesh 2-3 cm above the Petri dish, and rinse with 6 mL of modified HBSS.
   NOTE: This step will separate the epithelial sheets, which are retained on the top, from single cells that pass through the mesh. The flow-through is a mixed cell suspension, which includes immune cells and stromal fibroblasts.
6. Collect the mixed cell suspension and centrifuge at 500 x g for 10 min. Aspirate the liquid and resuspend the pellet in 4 mL of modified HBSS for density gradient centrifugation.
7. Layer the mixed cell suspension over 3 mL of polysucrose solution and centrifuge at room temperature at 500 x g for 30 min with no brake. Collect the white band from the top of the polysucrose solution and wash with PBS.
   NOTE: This fraction is the enriched mixed cell suspension, which is rich in immune cells and contains some stromal fibroblasts.

3. Dead Cell Removal

NOTE: If a large percentage of dead cells (> 20%) are present in the enriched mixed cell suspension after density gradient centrifugation, dead cell removal with magnetic beads is recommended. Dead cells need to be removed since they may interfere with the positive magnetic bead selection process.

1. Count the cells using a hemocytometer on a light microscope with the 10X objective. Spin down all the cells in the enriched mixed cell suspension (500 x g, 10 min, 4 °C). Completely aspirate and discard the supernatant. Add 100 µL of dead cell removal beads per 1 x 10⁷ total cells (live and dead), according to the manufacturer's instructions. Incubate at room temperature for 15 min.
2. Place a 30 µm filter on the magnetic column, apply the cell suspension and allow the bead-labeled cells to fall through the magnetic column.
3. Rinse the column with dead cell removal buffer as recommended (3 mL). Collect the flow-through containing the live cells. The range of cell recovery after dead cell removal is presented in Figure 1B.
   NOTE: These cells can be used to perform flow cytometry staining for phenotypical studies (section 5) or continue with DC isolation (section 4).

4. DC Purification by Positive Magnetic Bead Selection

1. Following dead cell removal (step 3.2), spin the cells down, remove the supernatant, and resuspend the cell pellet in 80 µL magnetic selection buffer per 1 x 10⁷ cells (PBS, 0.5% heat-inactivated human AB serum (HS), 2 mM EDTA).
2. For positive selection of DC populations, add CD1a or CD14 magnetic beads according to the manufacturer's instructions (20 µL magnetic beads per 1 x 10⁷ cells). Incubate for 15 min at 4^°C.
3. Wash the cells with the magnetic selection buffer, spin down, and remove buffer completely. Resuspend the cells in a minimum of 500 µL of magnetic selection buffer.
4. Attach the column to the magnet and add a 30 µm filter on the top of the column to retain any remaining cell clumps. Rinse the filter and column with magnetic selection buffer as recommended.
5. Apply the cell suspension to the column. Rinse 3 times with magnetic selection buffer.
6. Transfer the column to a 15-mL tube, add 5 mL of magnetic selection buffer, and apply the plunger to release the selected cells from the column.
7. To increase the purity, repeat steps 4.4-4.6 with the recovered cells using a new column. The range of cell recovery after magnetic bead selection is presented in Figure 1C.

5. Assessment of Cell Purity: Flow Cytometry and Microscopy

1. Antibody staining for flow cytometry:
   1. Resuspend up to 1 x 10⁶ cells in 100 µL of PBS containing 20% heat-inactivated HS to block Fc receptors, and incubate for 10 min on ice.
   2. Add the desired fluorescently labeled antibodies and the reagent for identification of dead cells (for an example, see section 6.5). Set fluorescence minus one (FMO) controls to establish the gates, and use compensation beads to prepare single stain controls for compensation. Examples of antibodies used are given in Table 1. Incubate for 20 min at 4 °C, protected from light.
   3. Wash the cells with 2 mL of magnetic selection buffer.
      NOTE: Presence of EDTA is important to avoid the formation of cell clumping for flow cytometric analysis. Spin down and discard the supernatant.
   4. Fix the cells by adding 100 µL of 2% paraformaldehyde solution per tube.
   5. Analyze the samples in by a flow cytometer^10,11.
2. Spin the column (e.g., cytospin) and use Giemsa staining to analyze morphology:
   1. Resuspend the cells in 200 µL of magnetic selection buffer.
   2. Load in a spin column and spin for 5 min. Let the slide dry completely.
   3. Fix the slide by immersion in 100% methanol for 2 min. Rinse the slide with deionized water and allow it to air dry.
   4. Perform a standard Wrights-Giemsa staining. Cover the cells with Eosin and incubate for 10 min, rinse with deionized water, and let it dry. Cover the cells with Wrights-Geimsa stain, incubate for 1 min, rinse with deionized water, and air dry.
   5. Examine the preparations with an inverted microscope (Figure 2A, 40X objective).

6. Allogeneic Stimulation Assay to Assess DC Cell Function

1. Thaw the frozen peripheral blood mononuclear cells (PBMCs)
   1. Warm the cell culture media with 10% HS to 37 °C.
   2. Thaw the PBMCs by placing the cryotube in a 37 °C water bath, until a small amount of frozen cell media remains in the cryotube. In a sterile environment, transfer the contents of the cryotube to 10 mL of pre-warmed cell culture media with 10% HS in a 15-mL tube.
   3. Centrifuge at ~ 500 x g for 7 min. Remove the media, disperse the pellet, resuspend the PBMCs in 10 mL of cell culture media with 10% HS, and centrifuge the cells.
   4. Repeat the wash step a third time. Count the cells.
2. Naïve T-cell selection:
   1. Use a negative selection magnetic antibody cell selection kit to isolate naïve T-cells according to the manufacturer's protocol.
   2. After isolation, check the purity using CD45RA and CCR7 antibodies.
      NOTE: Typical purity is > 99% naïve T-cells.
3. Naïve T-cell proliferation dye staining:
   1. Prepare 600 µL of 2x solution of cell proliferation dye, protecting the dye from exposure to light.
   2. Wash the naïve T-cells 2x with PBS. Resuspend cells in 500 µL PBS.
   3. While gently vortexing the cells in the biosafety cabinet, add 500 µL of proliferation dye to the naïve T-cells.
   4. Incubate the cells for 10 min at 37^°C.
   5. Stop the cell labeling by adding 5 mL of cell culture media with 10% HS to the cells. Incubate on ice for 5 min, protected from light.
   6. Centrifuge the cells at ~ 500 x g for 7 min, aspirate the media, and resuspend the cells in 4 °C of cell culture media with 10% HS. Repeat two more times for a total of 3 washes. Take an aliquot of cells to count prior to the last centrifugation.
      NOTE: Typically, final naïve T-cell numbers will range between 5% and 15% of the initial PBMC cell count.
   7. Resuspend the cells with the cell culture media with 10% HS at the desired cell concentration.
4. DC-allogeneic naïve T-cell co-culture:
   1. Plate the isolated DCs and naïve T-cells in a 96-well, round-bottom plate, at a 1:15 ratio of DCs to T-cells. Ensure that the optimal number for DCs ranges between 3,000 and 5,000 cells/well.
   2. Centrifuge the plate at ~ 500 x g for 3 min to ensure the cells are located at the center of the well. Cell-to-cell contact is important for proper signaling to occur.
   3. Incubate at 37 °C for 6 days. Monitor the cluster of cells, which appear in the wells as proliferation occurs, with microscopy (Figure 3A).
5. Flow cytometry staining to assess proliferation:
   1. After 6 days, cells can be examined by flow cytometry. Stain the co-culture for flow cytometry.
   2. Prepare a 2x solution of yellow viability dye (maximum emission 572 nm) in PBS.
      NOTE: The use of PBS without serum is important, as serum proteins interfere with the dye).
   3. Centrifuge the plate with cells at ~ 500 x g for 5 min.
   4. Remove 100 µL of supernatant.
      NOTE: At this point collect and store the supernatant for separate analysis of soluble factors.
   5. Wash the cells two times with PBS: add 150 µL/well of PBS, centrifuge at ~ 500 x g for 5 min, remove 150 µL of supernatant with a pipette. Ensure that there is 50 µL of supernatant remaining in each well.
   6. Add 50 µL of 2x viability dye to each well. Incubate at room temperature for 10 min, protected from light.
   7. During the incubation, prepare a master mixture of desired antibody markers. For example: CD3 APC/Cy7, CD4 PE, CD8 FITC, etc.
   8. Add the antibody mixture to each well. Incubate at room temperature for 10 min, protected from light.
   9. Wash the cells 2x with PBS + 2% HS: add 150 µL/well of PBS + 2% HS. Centrifuge at ~ 500 x g for 5 min. Remove 150 µL of supernatant with a pipette.
   10. Fix the cells by adding 100 µL/well of 2% para-formaldehyde. If needed: transfer the cells and media to polypropylene flow cytometry tubes. Tubes can be kept at 4 °C and protected from the light, until the flow cytometric analysis.
   11. Read tubes in a flow cytometer.