Placentae were collected from the Labor and Delivery Unit at University Hospital under a protocol approved by the Institutional Review Board of Oregon Health and Science University in Portland, Oregon, with informed consent from the patients.

1. Collection of Placental Tissue

1. Preparation
   NOTE: All equipment that encounters tissue must be sterile.
   1. Sterilize the dissecting equipment by autoclaving for 60 min at 121 °C.
   2. Use personal protective equipment (PPE): lab coats/gowns/scrubs, gloves, and mask with a face shield or goggles.
   3. Turn on water bath and set to 37 °C.
   4. Warm two 50 mL conical tubes, each containing 25 mL of complete media (Iscove's Modified Dulbecco's Medium supplemented with 10% FBS and 1% Penicillin/Streptomycin, Table 3) in a 37 °C water bath.
   5. With informed patient consent, obtain a placenta immediately following delivery by cesarean section.
   6. Become familiar with the placenta. The umbilical cord is inserted on the fetal side (chorionic plate) and blood vessels can be seen radiating out from the umbilical cord insertion site. The opposite side is the maternal side, or basal plate. It includes the decidua and structures that contain vessel trees, called cotyledons.
2. Sampling of Villous Tissue
   NOTE: Villous tissue should be sampled from the placenta as soon as possible following delivery, preferably in 30 min or less.
   1. With the chorionic plate facing upwards, excise 2-3 full-thickness sections (approximately 2.5 cm x 2.5 cm in size) 2-3 cm in from the periphery of the placenta at random, using forceps and scissors (Figure 1A).
      NOTE: Avoid parts of the placenta that appear abnormal (i.e. white calcifications).
   2. Trim away the chorionic and basal plates and any large blood vessels.
   3. Place the resulting villous tissue (Figure 1B, approximately 80 – 120 g total) in warm complete media and begin trophoblast isolation within 30 min of sampling.
      NOTE: Some downstream assays and applications are affected by the length of latency period between delivery, sampling, and trophoblast isolation. It is best to keep these periods as short as possible and consistent between isolations.
   4. Using the techniques described in steps 1.2.1 – 1.2.2, sample 5 random 1 cm x 1 cm sections of villous tissue from across the placenta (including the center).
   5. Cut each 1 cm x 1 cm villous tissue sample into 4-5 smaller pieces (approximately 30 mg each), place in 2 mL microcentrifuge tubes, and flash freeze in liquid N₂. Store at -80 °C for use in later analyses.

2. Isolation of Trophoblasts from Villous Tissue

1. Preparation
   NOTE: Use sterile equipment and perform procedures involving cells in a laminar flow hood.
   1. Thaw four frozen density gradients (Table 1, see table of the essential supplies, reagents, and equipment, supplementary material) at 4 °C the night before the placenta arrives.
      NOTE: Alternatively, density gradients can be made on the day of the isolation.
   2. Turn on centrifuge and set to 20 °C.
      NOTE: All centrifugation steps are at maximum acceleration and deceleration unless specified otherwise.
   3. Prepare 1x HEPES-buffered salt solution (HBSS) supplemented with Ca⁺² and Mg⁺² (Table 3).
   4. Warm trypsin to 37 °C.
   5. Dilute DNase to approximately 2720 kilounits/mL in sterile supplemented HBSS.
   6. Warm 50 mL of Newborn Calf Serum (NCS) to 37 °C.
   7. Warm complete media to 37 °C.
   8. Warm freezing media (90% FBS, 10% DMSO, Table 3) to 37 °C.
      CAUTION: DMSO is toxic and must be handled with gloves.
   9. Prepare digestion buffer (Table 2 and 3) by mixing 308 mL of supplemented HBSS, 50 mL of Trypsin (3751.7 BAEE units/mL), and 0.5 mL of DNase (379.4 kilounits/mL) in a sterile bottle.
      NOTE: The approximate time required to isolate cells from villous tissue is 7 h.
2. Processing the Villous Tissue
   1. Rinse each piece of villous tissue in a 50 mL conical tube filled with room temperature phosphate buffered saline (PBS). Repeat and replace the PBS as necessary until the excess blood is removed (PBS rinse will be light red or pink when the tissue is thoroughly rinsed).
   2. Place the villous tissue in a sterile Petri dish and remove as many blood vessels as possible by gently scraping off the soft villous tissue from the vessels using a microscope slide.
   3. Finely mince the resulting villous tissue using scissors.
3. Villous Tissue Digestion and Crude Isolation of Trophoblasts
   1. Transfer the minced villous tissue to a sterile bottle with digestion solution according to the calculated volumes based on the specific activity of trypsin and DNase (165 mL, Table 2).
   2. After 35 min of incubation in a 37 °C water bath with shaking at 70 revolutions per minute (rpm), tilt the digestion bottle on its side and allow the undigested pieces of tissue to settle at the bottom of the bottle. Carefully draw up the supernatant with a serological pipette, avoiding the settled tissue.
   3. Dispense the supernatant through a 100 µm cell strainer equally between 50 mL conical tubes.
      NOTE: To save time, it is advisable to begin the second digestion by adding digestion solution (110 mL, Table 2) to the remaining settled tissue and resuming incubation as described in step 2.3.2.
   4. Gently layer 3 – 5 mL of NCS underneath the strained supernatant by slowly dispensing from a serological pipette at the bottom of the tube. A meniscus between the strained supernatant (containing trophoblasts) and NCS should be visible (Figure 2A).
   5. Centrifuge the supernatants over NCS at 1,250 x g for 15 min at 20 °C. The resulting pellet will include red blood cells in the lower-most layer followed by a white layer containing the trophoblast cells (Figure 2B).
   6. Repeat steps 2.3.2 – 2.3.5 for each of the second and third digestions (adding 110 mL and 83.5 m, respectively, of digestion solution to the bottle of tissue, Table 2).
   7. Once all supernatants have been centrifuged, resuspend each pellet in 5 mL of warm complete media, and then pool the suspensions together.
   8. Split the cell suspension equally between two 50 mL conical tubes and centrifuge at 1,250 x g for 15 min at 20 °C.
   9. Gently remove the supernatant and resuspend each of the cell pellets in 6 mL of warm complete media.
4. Density Centrifugation
   1. Divide the cell suspension equally between four density gradients (3 mL each) by slowly and carefully layering the cell suspension on the top of the density gradients with a transfer pipet.
   2. Centrifuge the density gradients at 1,250 x g for 20 min at 20 °C with minimum acceleration and deceleration. This should produce distinguishable bands of sedimented cells (Table 4).
   3. Slowly and carefully remove the top layers of density gradient media (DGM) until the opaque band(s) containing trophoblast cells (between 35 - 50% DGM) is reached (Table 4).
   4. Transfer the trophoblast bands into two 50 mL conical tubes and fill with warm complete media.
   5. Gently invert the tubes 3 – 6 times to mix and centrifuge at 1,250 x g for 15 min at 20 °C.
   6. Remove the supernatant and resuspend each cell pellet in 5 mL of warm complete media. Combine the cell suspensions and count viable cells using a hemocytometer and Trypan blue (or preferred cell counting method).
5. Counting Cells with a Hemocytometer
   1. Mix the cell suspension by pipetting up and down with a serological pipet or by gently inverting the tube several times.
   2. Combine equal parts of cell suspension and Trypan blue (i.e. 20 µL each) in a separate tube and mix gently.
   3. Incubate for 1 – 2 min at room temperature.
   4. Gently dispense 15 – 20 µL of the cell-Trypan blue mixture in between the coverslip and the hemocytometer and allow the cells to diffuse across the grid by capillary action.
   5. Count viable cells (dead cells will be stained deep blue) in each of the four 4 x 4 quadrants with a tally counter. Employ a system of counting to ensure cells are not counted more than once (i.e. do not count cells that touch the bottom or left boundaries).
      NOTE: Each 4 x 4 quadrant should contain between 50 – 150 cells. Too few or too many cells can lead to an over or underestimation of cell number.
   6. Average the total cell counts from each of the 4 x 4 quadrants, multiply by 10⁴, and then multiply by the dilution factor (cell suspension to Trypan blue) to calculate number of cells per mL.
   7. Multiply the number of cells per mL by the total volume of cell suspension to calculate the total cell yield.
      NOTE: Approximately 100 million cells are expected from isolations starting with 80-120 g of villous tissue.
6. Plating Cells
   1. Plate 3 million cells/well (3.3 x 10⁵ cells per cm²) in a 6-well plate (2 mL of suspension per well) and gently rock back and forth and side to side to evenly distribute the cells.
      NOTE: Trophoblasts require tissue culture treated plates to adhere properly. A monolayer of cells is required to promote syncytialization.
   2. Leave the plated cells in the laminar flow hood for approximately 30 min to allow cells to evenly distribute, settle, and begin to adhere to the bottom of the wells before placing in the incubator.
   3. Culture cells for up to 72 h (with daily media changes) in a 37 °C incubator with 5% CO₂ and 95% humidity.
      NOTE: Examine the trophoblasts under a microscope at 10 – 20x every 24 h of culture. Trophoblasts do not proliferate and cannot be passaged. Over the course of 72 h of culture, the round individual trophoblasts fuse to form a syncytium (Figure 3A and B).
7. Freezing Cells
   1. Pellet unused cells by centrifugation at 1,250 x g for 10 min at 20 °C.
   2. Aspirate as much media as possible from the pellet.
   3. Resuspend the pellet in freezing media (Table 3).
   4. Freeze the aliquots at -80 °C in a freezing container filled with 100% isopropanol. Transfer the frozen aliquots to liquid N₂ the following day for long term storage.
      NOTE: Cells can be cultured after freezing.
8. Thawing Cells
   1. Remove an aliquot of frozen cells and thaw in a 37 °C water bath while swirling. Remove the aliquot from the water bath just before it has fully thawed to a liquid.
   2. Immediately transfer the thawed cells to a 15-ml conical tube. Beginning slowing at first, add 10 mL of complete media with intermittent mixing.
   3. Invert tube several times to mix.
   4. Centrifuge at 200 x g for 10 min at 20 °C.
   5. Aspirate the supernatant and resuspend the pellet in 2 – 5 mL of warm complete media.
   6. Count the cells and plate as previously described.

3. Treatment of Primary Trophoblasts with TNFα, Collection of Cell Lysates, and Western Blotting

1. Preparation
   1. Warm complete media to 37 °C.
   2. Make a 10 µg/mL stock of TNFα in complete media and store at -20 °C until ready for use.
   3. Make a 1 µg/mL working stock of TNFα in complete media when ready to treat the cells. Serial dilute the TNFα working stock to 10⁴ pg/mL, 10³ pg/mL, 500 pg/mL, 250 pg/mL, and 125 pg/mL in complete media.
      NOTE: The TNFα concentration of 10⁴pg/mL is moderately cytotoxic. Adjustment of the TNFα concentrations tested will depend on the specifics of the desired downstream applications.
2. Treating Cells with TNFα
   1. After 24 h of culture, aspirate the culture media from the cells and replace with 2 mL of TNFα supplemented medias per well on 6-well plates (at least two wells per treatment and vehicle control).
   2. After 24 h of TNFα-exposure (48 h of culture), replace the TNFα supplemented medias with complete media.
3. Harvesting Cells and Total Protein
   1. After 72 h of culture (24 h past removal of TNFα), aspirate media, gently rinse cells with room temperature PBS, and add 80 µL of ice cold Radioimmunoprecipitation Assay Buffer (RIPA) containing freshly added protease and phosphatase inhibitors (Table 3) directly to each well.
   2. Remove the cells from the plate with a cell scraper. Transfer the cells to a 1.5 mL microcentrifuge tube, pooling wells within treatment groups.
   3. Lyse the cells by vortexing the tubes on high for at least three intervals of 15 s.
   4. Incubate the cells at 4 °C with rocking for 15 min.
      NOTE: Alternatively, after step 3.3.3., the cells can be incubated on ice for 15 min with intermittent agitation by flicking the tube, vortexing, or pipetting up and down.
   5. Repeat step 3.3.3.
   6. Centrifuge the tubes at 10,000 x g for 5 min at 4 °C to pellet cellular debris.
   7. Transfer the supernatant (containing cellular protein) to a new 1.5 mL microcentrifuge tube and store at -80 °C.
      NOTE: The protocol can be stopped here and resumed at a later time. It is advisable to make several aliquots of cellular protein samples to avoid multiple freeze-thaw cycles.
   8. Determine total protein concentration by a preferred method, such as a bicinchoninic acid assay (BCA, see table of the essential supplies, reagents, and equipment, supplementary material).
4. SDS-PAGE and Western Blotting for Rubicon or Protein of Interest
   NOTE: Follow Western blotting protocols according to manufacturer's instructions using a preferred laboratory system.
   1. Load between 20-40 µg of total protein in sample buffer (Table 3) per well on a 12% acrylamide gel. Separate proteins by SDS-PAGE in running buffer (Table 3).
   2. Wet-transfer the proteins from the gel to a polyvinylidene difluoride (PVDF) membrane according to manufacturer's instructions in transfer buffer (Table 3).
   3. Incubate the membrane in 5% milk powder in Tris-Buffered Saline with 0.1% Tween 20 (TBST, Table 3) for at least 1 h at room temperature with rocking.
   4. Incubate the membrane in a Rubicon primary antibody (see table of the essential supplies, reagents, and equipment, supplementary material) at 1:500 in 1% milk powder in TBST overnight at 4 °C with rocking.
   5. Gently wash the membrane 3 x 5 min in TBST and 1 x 5 min in TBS at room temperature with rocking.
   6. Incubate the membrane in 1:2000 – 1:5000 secondary antibody (HRP-linked or preferred visible conjugate, see table of the essential supplies, reagents, and equipment, supplementary material) in 5% milk powder in TBST for at least 1 h at room temperature with rocking.
   7. Repeat the washing procedure outlined in step 3.4.5 and visualize the blot with an appropriate visualization (i.e. chemiluminescent) substrate on an imaging system.
   8. Repeat the washing procedure outlined in step 3.4.5 and probe the membrane for β-actin or a preferred loading control according to manufacturer's instructions.
   9. On preferred image analysis software, analyze the expression of Rubicon by manual quantification of absorbance (band intensities), subtraction of background absorbance, and normalization to the corresponding loading control band intensities. Perform statistical analyses as appropriate to test for statistically significant changes in protein levels.