Isolation of Leukocytes from the Murine Tissues at the Maternal-Fetal Interface

Before working with the samples mentioned in this protocol, animal ethical approval must be given by the Local Research Ethics Committee and the Institutional Review Boards. When working with animal blood, cells, or hazardous agents as mentioned in this protocol, the proper biosafety and laboratory safety actions must be followed.

1. Mouse Handling and Tissue Collection

1. Prepare a sterile workstation and obtain sterile tools for tissue collection. These tools will include large and small surgical scissors, forceps, and fine-tip tweezers. Include small Petri dishes labeled with the appropriate tissue names, filled with 1x phosphate-buffered saline (PBS) solution (3 – 5 ml) equilibrated to RT (20 – 22 °C). Also include a large Petri dish, unlabeled, filled with 1x PBS solution (10 – 15 ml).
2. Euthanize a pregnant mouse (10.5 to 19.0 days post-coitum (dpc) or prior to delivery) using carbon dioxide (CO₂). To ensure that the mouse is euthanized, use the cervical dislocation technique.
   Note: Prior to 10.5 dpc, it is difficult to manually separate the decidual tissues from the uterine tissues. If the isolation of decidual leukocytes is not required, leukocytes from the uterine tissues of non-pregnant mice or those from mice at <10.5 dpc can also be performed by following this protocol.
3. Using a pair of sterile surgical scissors, completely remove the lower abdominal portion of the skin and muscular tissue. With forceps, move aside all other organs until the uterus and ovaries are visible. With the uterus still attached, use the forceps to move it outside of the peritoneal cavity (Figure 1A).
4. Locate the ovaries distal to the oviduct and uterotubal junction of each uterine horn. Using the small surgical scissors, make an incision at the uterotubal junction and separate the uterine horns from the mesentery containing the uterine vessels; then, excise at the cervix to detach the uterine horns from the peritoneal cavity (Figure 1B).
5. With small surgical scissors, make an incision between the cervix and the lower segment of the uterine horns. Leave attached a portion of the cervix during this process (Figure 1C) to avoid opening the uterine horn.
6. Immerse the uterine horns (including the cervix) in a large Petri dish filled with 1x PBS solution (10 – 15 ml) to hydrate the implantation sites.
7. While still immersed in 1x PBS solution, trim any fat from the uterine horns with the small surgical scissors. Keep the tissues immersed in 1x PBS solution throughout the dissection.
8. Holding the uterine horns in place with the forceps, remove one of the implantation sites from the uterus with a transverse cut through the inter-implantation region, using the small surgical scissors (Figure 1D). The implantation site contains the fetus, surrounded by the chorioallantoic membrane, and the uterine, decidual, and placental tissues.
9. Using the forceps, hold the site in place and make a small incision in the uterine wall adjacent to the placental and decidual tissues (Figure 1E). Trim around the perimeter of the placenta/decidua until these become separated from both the uterine wall and the chorioallantoic membrane that includes the fetus (Figure 1F).
10. Remove the placenta and attached decidual tissues and place in 1x PBS solution (3 – 5  ml; see step 1.12).
11. Place the fetus surrounded by the chorioallantoic membrane attached to the uterine tissues in a Petri dish filled with 1x PBS solution (3 – 5  ml). The hydration of these tissues will facilitate the separation of the chorioallantoic membrane from the uterine wall (see step 1.14).
12. Using a pair of fine-tip tweezers, gently peel the decidual tissue (white-grey layer) from the placental surface. Perform this step carefully to maintain the integrity of the placental and decidual tissues (Figure 1G).
13. Place the placenta and decidual tissues in two separately labeled Petri dishes filled with 1x PBS solution (3 – 5  ml each).
14. Gently remove the uterine wall from the chorioallantoic membrane with fine-tip tweezers. These are the uterine tissues.
15. Place the uterine tissues in a Petri dish filled with 1x PBS solution (3 – 5  ml).
16. Repeat steps 1.8 through 1.15 until all implantation sites have been processed.
17. Collect several uterine, decidual, and placental tissues to improve the yield of cells during enzymatic digestion. The amount of tissues depends on the litter size; however, leukocyte isolation requires >2 pieces of tissue.
    Note: For more detailed information regarding the dissection of the implantation sites on different gestational days, view the pictures found in Chapter Two of The Guide to Investigation of Mouse Pregnancy⁴².

2. Uterine, Decidual, and Placental Tissue Dissociation

1. Label several sterile 2 ml conical tubes with the appropriate tissue name, and place a vial with 1,000 µl of enzymatic solution on ice.
2. From the Petri dish filled with 1x PBS solution, transfer two pieces (100 – 150 mg) of the uterine, decidual, or placental tissues to a labeled 2 ml conical tube.
3. Add 500 µl of cold enzymatic solution into each 2 ml conical tube.
4. With fine sterile scissors, begin to dissociate the tissue immersed in the enzymatic solution until it is finely minced. Over time, the suspension will develop a milky appearance. This step should not exceed more than 2 min to prevent over-manipulation of the sample.
5. Once the tissue has been dissociated, add an extra 500 µl of cold enzymatic solution to the 2 ml conical tube.
6. Place the 2 ml conical tube on ice.
7. Repeat steps 2.2 through 2.6 with each individual tissue until all tissues have been processed.
8. Transfer all 2 ml conical tubes with homogenized tissue (minced tissue + enzymatic solution) samples from the ice into an incubator at 37 ºC and perform a gentle orbital agitation (80 rpm). Incubate for 30 – 35 min. Make sure the temperature of the incubator is stabilized before starting the incubation time.
9. Following incubation, remove the 2 ml conical tubes with dissociated tissues from the incubator and place them on ice to prevent any further digestion.
10. Label sterile 50 ml conical tubes according to tissue type. Remove the lids and replace them with a sterile cell strainer (100 µm pore size).
11. Pour the dissociated tissues from the 2 ml conical tubes through the cell strainer using ~20 ml of 1x PBS solution. The fragments of tissue will remain in the cell strainer and the cell suspension will pass through it.
12. Wash the dissociated tissues in the cell strainer 2 – 3 times with 20 ml of 1x PBS solution using a transfer pipette.
13. Rinse the empty 2 ml conical tubes with 1x PBS solution (1 ml) and pour the contents through the cell strainers.
14. Remove the cell strainers from the 50 ml conical tubes and replace the lids. Centrifuge the tubes at 1,250 x g for 10 min at 4 ºC.
15. Carefully, aspirate the supernatant without disturbing the cell pellet. The cell pellet contains the isolated leukocytes and stromal cells.
16. Re-suspend the cell pellet in 1 ml of RPMI culture medium without fetal bovine serum (FBS). Mix gently. Do not use vortex.
17. Add 500 µl of neat FBS to a 5 ml polystyrene plastic tube and slowly overlay the cell suspension.
18. Centrifuge for 10 min at 1,100 x g without the brake at RT to pellet viable cells while cellular debris is retained at the PBS/FBS interface.
19. Aspirate the supernatant without touching the cell pellet. The cell pellet contains mostly viable cells.
20. Optionally: To concentrate the mononuclear cells, use a 20% saturated polysaccharide solution, as described below.
    1. Resuspend the cell pellet in 1,000 µl of RPMI culture medium without FBS.
    2. Add 1 ml of 20% saturated polysaccharide solution to a 5 ml polystyrene plastic tube and slowly overlay the cell suspension on top of this solution, according to the manufacturer’s instructions. While layering the sample, avoid mixing the 20% saturated polysaccharide solution with the cell suspension.
    3. Centrifuge with a swing-out rotor at 500 x g for 30 min at RT without brake. It is very important to turn off the brake to avoid mixing the cells and losing the gradient. Mononuclear cells will be found in the interface between the 20% saturated polysaccharide solution and the 1x PBS solution.
    4. Aspirate and discard the supernatant. The cell pellet contains mostly mononuclear cells.
       Note: To perform viability staining and intracellular staining, see step 3. To perform immunophenotyping, see step 4. To perform viability staining and extracellular staining only, see step 5. To perform a cell culture of isolated leukocytes, see step 6. To perform magnetic cell sorting, see step 7.

3. Viability Staining for Fixable Cells

1. Resuspend the cell pellet in 1,100 µl of 1x PBS solution. Mix gently using a micro-pipette.
2. Transfer 100 µl of the cell suspension into a 5 ml polystyrene plastic tube. Add 400 µl of 1x PBS solution to the 5 ml polystyrene tube. This tube is a control for tissue auto-fluorescence. Store at 4 ºC until step 3.5.
3. Transfer the remaining 1,000 µl of the cell suspension to a new 5 ml polystyrene plastic tube. Add 1 µl of the viability dye and gently homogenize. Incubate for 30 min in darkness at 4 ºC.
4. Following incubation, add 1,000 µl of 1x PBS solution. Mix gently by hand.
5. Centrifuge both 5 ml polystyrene plastic tubes (steps 3.2 and 3.3) at 1250 x g for 10 min at 4 ºC. Aspirate and discard the supernatant.
   Note: The cell pellet (step 3.3) will be used in step 4. The cell pellet (step 3.2) will serve as a control for tissue auto-fluorescence.

4. Immunophenotyping

1. Resuspend the cell pellet in 50 µl of anti-mouse CD16/CD32 (diluted 1:100 in FACS buffer [0.1% BSA, 0.05% sodium azide, 1x PBS solution, ph 7.4]). Mix the cell suspension gently. Do not use the vortex.
2. Incubate the cell suspension in the 5 ml polystyrene tube in darkness at 4 °C for 10 min.
3. Following incubation, add 50 µl of each anti-leukocyte monoclonal antibody reacting with extracellular cell surface markers or isotype-matched control antibody (antibody dilutions must be validated). For example, to analyze leukocyte sub-populations, use anti-mouse antibodies reacting with the following leukocyte-cell surface markers: CD45, Ly6G, F4/80, CD11c, CD49b, B220, CD3, CD4, and CD8 (Table 1).
4. Once antibodies against extracellular markers have been added to the 5 ml polystyrene plastic tube, mix gently. Incubate in darkness for 30 min at 4 °C.
5. During this incubation, prepare and pre-warm the fixation buffer solution (diluted with deionized water, 1:5) to 37 ºC, if required. Keep the buffer at 37 ºC in darkness until its use is necessary.
6. After 30 min incubation, add 500 µl of FACS buffer in the 5 ml polystyrene plastic tube and gently mix. This step is needed to wash the cells and remove any excess of unbound antibody.
7. Centrifuge the samples at 1,250 x g at 4 °C for 10 min. Aspirate and discard the supernatant.
   Note: If intracellular staining is not required, see step 5. If intracellular staining is required, perform permeabilization and intracellular staining here, and then proceed with the fixation (step 4.8).
8. Add 500 µl of the pre-warmed fixation buffer solution to the cell pellet and incubate in darkness for 10 min at 37 ºC.
9. Dispense 500 µl of FACS buffer into the 5 ml polystyrene plastic tube. Mix gently.
10. Centrifuge the samples at 1,250 x g at 4 °C for 10 min. Aspirate and discard the supernatant.
11. Add 500 µl of FACS buffer to each sample. These samples can now be analyzed by flow cytometry. Acquire the samples immediately for best results.

5. Viability Staining for Unfixable Cells

1. After extracellular immunophenotyping, resuspend the cell pellet in 500 µl of FACS buffer.
2. Add 1 µl of 4’,6-diamidino-2-phenylindole dihydrochloride (DAPI, 200 µg/ml) just before acquiring the sample.
3. Visualize the samples within 1 – 2 min after adding DAPI using a flow cytometer.

6. Cell Culture

1. Prepare a workstation under a sterile fume hood. Pre-warm the RPMI culture medium to 37 °C using a water bath.
2. Resuspend the cell pellet in 1000 µl of FACS buffer for cell counting.
3. Count the number of viable cells using an automatic cell counter or hemocytometer and 0.4% trypan blue solution, following the manufacturer's instructions. Record the total live cell count.
4. Centrifuge the cells at 1,250 x g at 4 °C for 10 min to remove the FACS buffer.
5. Resuspend the cell pellet in 500 µl of pre-warmed RPMI culture medium by gently pipetting 2 to 3 times.
6. Plate the desired number of cells in a sterile 24-well plate and incubate at 37 °C. The incubation time will be determined by the research question.

7. Magnetic Cell Sorting

1. Set up a clean workstation with the following materials: MS columns, magnetic cell separator, 15 ml conical tubes, 30 µm pre-separation filters, and a multi-stand.
2. Resuspend the cell pellet in 500 µl of MACS buffer (0.5% BSA, 2mM EDTA, 1x PBS solution, pH 7.2).
3. Using an automatic cell counter or hemocytometer and 0.4% trypan blue solution, count and record the total live cell count.
4. Centrifuge the cells at 1,250 x g at 4 °C for 10 min. Aspirate and discard the supernatant. Add another 500 µl of MACS buffer.
5. Proceed to perform the magnetic separation following the manufacturer’s recommendations.
6. Determine purity by flow cytometry or microscopy.