足月胎盘胎膜中原发性人蜕细胞的分离

The overall goal of this cell isolation protocol is to develop a method for isolating primary decidual cells with high yield end cell viability. This method can help answer key questions in the field of human reproduction, such as the deciduous role in blastocyst implantation and labor initiation. It can also provide insight into various pregnancy related pathologies.

The main advantage of this technique is that it provides a time and cost effective method for the isolation of primary decidual cells with high viability end yields. To begin, leave five 50 milliliter tubes on a rack and two bottles of HBSS and HBSS in the fume hood. Then pipette 25 milliliters of HBSS in the appropriately labeled tube.

Next, lay overlapping diapers in the fume hood. Then, take the term placenta out of the container and place it in the fume hood. Position the placenta on the diaper pad with the maternal side up.

If the placenta is maternal side down, use an additional diaper to flip the placenta. Look for the point of membrane rupture and make incisions using scissors and forceps to allow the membrane to unfold and lay flat on the fume hood surface. Then use a cell scraper to gently scrape the decidual tissue off of the chorion.

Transfer the tissue to a 50 milliliter tube containing 25 milliliters of HBSS Remember to also collect small blood clots, as these also contain some decidual cells. Gently shake the 50 milliliter tube to wash the collected tissue, then pass the tissue through a 250 micrometer metal sieve. Repeat the washing process twice with HBSS followed by two washes with HBSS in fresh tubes.

Chorion contamination is easily visible due to its threadlike shape and lighter color and can be removed after washing. After washing, place the tissue in a sterile 50 milliliter tube containing 20 milliliters of enzymatic digestion solution to digest the entire decidua obtained from the whole term fetal membrane. Use paraffin film to seal the cap of the tube.

Then incubate the tube at 37 degrees Celsius in a shaking water bath. Set the timer for 20 minutes. Once the incubation is over, remove the paraffin film.

Use 70%ethanol to sterilize the surface of the tube containing the digested tissue and place it under the fume hood. Shake the tube briefly. Next, pass the cell suspension through a metal sieve into a new sterile specimen container.

Dilute the cell suspension with an equal volume of culture medium to stop the enzymatic reaction. Transfer the cell suspension to a new 50 milliliter tube. Then centrifuge the cells at 420 G for 11 minutes at four degrees Celsius.

If required, place the remaining undigested tissue into a new 50 milliliter tube containing 20 milliliters of fresh enzymatic digestion solution and repeat the digestion process. If the digestion is performed a second time, leave the first tube with the cell suspension on ice. Once the second digestion is complete, combine both cell suspensions.

After centrifugation, carefully aspirate the supernatant. A manual pipette may be required to remove the supernatant. Then, re-suspend the cell pellet in wash buffer.

After washing the cells, repeat the centrifugation process. At the end of the centrifugation process, aspirate the supernatant and re-suspend the cell pellet in five milliliters of wash buffer and 35 milliliters of erythrocyte lysis buffer. In order to lyse the red blood cells, briefly vortex the tube at the beginning and at the end of the incubation.

Then, incubate the cell suspension on ice for 20 minutes. After incubation, repeat the centrifugation process. Post-centrifugation, aspirate the supernatant and re-suspend the pellet in 40 milliliters of wash buffer, then pass the cells through a 70 micrometer nylon filter to exclude cell clumps.

Repeat the centrifugation process. Aspirate the supernatant once centrifugation is done. Then re-suspend the cell pellet in 10 milliliters of culture medium.

Count the number of cells with a hemocytometer using the Trypan blue dye exclusion method. In order to determine the cell type composition of the decidual cell population, a multi-color imaging flow cytometry gating strategy has been developed. The analysis shows exclusion of cell doublets and cell debris using serial gating.

Once a debris-free single cell population is obtained, cell viability is determined using a fixable viability dye. The analysis demonstrates 80%viability of the total decidual cell population, indicated by the blue gating. Next, the isolated decidual cells are stained with fluorophore conjugated mouse monoclonal antibodies.

This is to analyze the presence of stromal, immune, epithelial and trophoblast cell markers and the viability dye. These markers will determine the decidual cell types. Flow cytometry fluorescent images demonstrate a positively stained cell for all four markers and viability dye.

These markers are then used to determine the percentage of each cell type in the decidual cell population. Interestingly, the Vimentin positive stromal cells constitute the highest population of cells, while there are virtually no trophoblast cells. These results demonstrate the purity and high viability of the isolated primary human decidual cells.

Here, the dot plots obtained show the gating for the positive cell type populations. Fluorescent minus one controls are used to accurately gate the true positive decidual cell populations. Next, captured spinning disk confocal microscopy images are used to validate the flow cytometric results by showing that the decidual cell population is primarily composed of Vimentin positive stromal cells with no trophoblast cell contamination.

Once mastered, this technique can be completed in three hours. Following this procedure, the decidual cells can be analyzed in a variety of ways to examine gene and protein expression, which can better help us understand the cellular changes occurring in pregnancy complications.