Reprogramming Primary Amniotic Fluid and Membrane Cells to Pluripotency in Xeno-free Conditions

The protocol follows institutional guidelines of the ethics committee for human research. Written consent of the patient was obtained for using the amniotic fluid for research.

This protocol follows the policies of the Institutional Animal Care and Use Committee of the University of South Alabama.

1. Isolation and Culture of Primary Amniotic Mesenchymal Stem Cells

1. Plating of amniotic fluid cells
   1. Obtain a minimum of 2.5 mL of amniotic fluid harvested in the process of amniocentesis by a physician.
      NOTE: All handling of live cells and tissue must be performed in a sterile tissue-culture cabinet and proper personal protective equipment must be used. Familiarity with basic cell culture and sterile technique is required.
   2. Prepare the amniotic fluid and membrane cell (AFMC) culture medium: EBM-2 basal medium, 15% fetal bovine serum (FBS), 20 ng/mL of bFGF, 25 ng/mL of EGF, 10 ng/mL of IGF. For culture of primary amniotic fluid as well as amniotic membrane stem cells, the medium should be supplemented with antibiotic-antimycotic solution.
   3. On day 0, mix 2.5 mL of amniotic fluid with 3.5 mL of AFMC culture medium and plate into a T25 flask. Incubate at 37 °C and 5% CO₂ for at least 48 h undisturbed before checking for the presence of colonies.
   4. On day 5, colonies of adherent cells should be present. Gently rock the flask to dislodge cells that did not fully adhere to the bottom and debris and vacuum-aspirate the spent medium/amniotic fluid mixture using a Pasteur pipette. Replace with 5 mL of fresh AFMC medium.
   5. Culture for another 5 days. Change medium every other day as described in step 1.1.4.
2. Isolation of primary mesenchymal stem cells from human amnion
   1. Obtain placentas as soon as possible following birth, within 24 h at the latest, to maximize cellular integrity. Cut a 9 cm² segment of the amnion, remove blood clots and wash in a 50 mL centrifuge tube with 30 mL of PBS supplemented with antibiotic-antimycotic solution.
   2. Mince the membranes using a pair of scalpels to fine pieces in a sterile 10 cm tissue culture dish. The digestion of the membranes and the extraction of the cells will be achieved using a tissue dissociation system. Follow the manufacturer's protocol.
      NOTE: the finer the pieces of the tissue after mincing, the higher the cell numbers recovered after digestion.
   3. Transfer the minced membrane tissue mass using the scalpel blades into one tissue dissociation tube and mix with 4.7 mL of RPMI 1640 medium. Mix in the dissociation enzymes (see Table of Materials).
   4. Mount the tubes onto the tissue dissociator and run program "h_tumor_01". Incubate the tubes at 37 °C on a rocking platform for 30 min.
   5. Further dilute the suspensions with 35 mL of RPMI 1640 and apply to a 70 µm strainer placed over a 50 mL collection centrifuge tube.
   6. Centrifuge for 5 min at 200 x g at room temperature, discard supernatant, resuspend the pellet in 5 mL of RPMI 1640, count the cells using a hemacytometer, and plate at a density of 10,000 cells/cm² into tissue culture-treated vessels with freshly prepared AFMC medium supplemented with antibiotic-antimycotic solution.
      NOTE: In case of incomplete digestion, small pieces of tissue will be present and single cells will be scarce. Spin down again and plate the whole pellet into one T75 flask.
3. Culture of primary AFSC and AMSC
   1. Passage colonies of AFSC/AMSC by vacuum-aspirating spent medium using a Pasteur pipette and adding 2 mL of cell detachment enzyme into the flask. Incubate at 37 °C for 5-8 min.
   2. Tap on the flask to help dislodge the cells and mix the suspension with an equal volume of AFMC medium (antibiotic-antimycotic supplement should not be necessary from this point on). Centrifuge at 200 g for 4-5 min. Remove the supernatant using either a glass Pasteur pipette or simply by means of inverting the tube and emptying it into a waste container.
   3. Flick the bottom of the centrifuge tube to break up the pellet into a single cell suspension in the remaining drop of liquid and mix with AFMC medium for plating. Plate into T-flasks at a density between 2,500 and 5,000 cells/cm².
   4. Change medium every other day. Do not culture the cell lines beyond passage 6. For reprogramming purposes, use as low a passage as possible.
   5. Prepare frozen stocks of AFSC and AMSC as back-ups using freezing medium. Harvest cultures using a cell detachment enzyme, centrifuge at 200 g for 4 °C for 5 min.
   6. Remove the supernatant using a Pasteur pipette and flick the bottom of the tube to singularize the cells in the pellet. Resuspend in complete freezing medium at a density of 1×10⁶/mL and aliquot into cryovials. Store in a freezing container overnight at -80 °C. Then move to liquid nitrogen for long-term storage.

2. Reprogramming into Pluripotency

1. Obtain the reprogramming plasmids
   1. Purchase the reprogramming plasmids through a non-profit plasmid repository. A Material Transfer Agreement is needed.
   2. Transform the plasmids into E. Coli competent cells and isolate the plasmids using a commercial plasmid extraction kit. Follow the manufacturer's instructions.
   3. Measure the concentration of plasmid DNA using a spectrophotometer. Aim for a high resulting plasmid concentration, ideally around 1 µg/µL to avoid dilution of the sample during transfection.
   4. Measure the concentrations of the individual plasmids using a UV-spectrophotometer and aliquot them individually.
   5. Mix together 3 µg, 3 µg, and 2 µg of EN2K, ET2K and M2L plasmids, respectively. This is the reprogramming plasmid solution. The amount of plasmid solution is enough to transfect 1 x 10⁶ cells. Prepare several such aliquots.
   6. Store all aliquots at -80 °C.
2. Prepare target culture plates
   1. Coat one 6-well plate with vitronectin – add 1 mL of vitronectin dilution buffer into each well and mix in 40 µL of VTN stock solution (1 µg/cm²). Leave at room temperature (RT) or in the incubator at 37 °C for 1 h.
   2. Vacuum-aspirate the solution using a Pasteur pipette and replace with 2 mL of AFMC medium in each well. Store at 37 °C until the cells are to be plated.
      NOTE: Important: The AFMC medium used in this step should not contain any antibiotic or antimycotic solutions.
3. Harvest cultured primary AFSC/AMSC
   NOTE: Expand AFSC/AMSC in culture enough to make frozen stocks at a low passage number and dedicate one T-75 flask for reprogramming. Since as few as 100,000 cells are sufficient for an experiment, aim at harvesting around 500,000 cells to compensate for losses and if optimization of transfection parameters or different culture conditions are to be tested.
   1. At the earliest convenience at a low passage, harvest the AFSC/AMSC using cell detachment enzyme mix as described in step 1.3.1 to 1.3.2. After the cells have been centrifuged, proceed to the next step.
   2. Resuspend the pellet in 1 mL of PBS and mix well to wash serum components. Count the cells using a hemacytometer. Adjust the cell density to 100,000/mL of PBS and aliquot into 1.5 mL microcentrifuge tubes. This ensures only a minimal contact time between the cells and the buffer used for transfection.
   3. Place the microcentrifuge tube on top of 5 mL polystyrene tubes (as adaptors, will allow centrifugation in a regular swing rotor) and centrifuge at 200 x g for 4 min at room temperature. Invert the tubes and discard the supernatant into a waste container. Do not use a fixed-angle rotor.
   4. Perform an additional centrifugation step at 200 x g for 3 min at room temperature. This will allow the remaining liquid from the walls of the tube to collect at the bottom. Carefully aspirate all of it using a 200 µL pipette.
4. Transfection with reprogramming plasmids
   1. For reprogramming experiments, a transfection system (see Table of Materials) will be used to deliver reprogramming plasmids into the cells. Place transfection tips, transfection tubes, resuspension buffer, and electrolytic buffer into the tissue-culture cabinet. The kit reagents are kept at RT until they are opened, then they are stored at 4 °C.
      NOTE: We use the 10 µL version of the kit.
   2. Move the transfection device close so that its tube station can be placed directly into the cabinet. Fill one transfection tube with 3 mL of electrolytic buffer and mount the tube into the station by pushing it all the way inside the slot.
   3. Take the reprogramming plasmid solution aliquots prepared in step 2.1.5 out of -80 °C storage and allow them to thaw at RT in the culture cabinet.
   4. On the transfection device, select the following transfection parameters: 950 V, 40 ms, and 1 pulse.
   5. Resuspend the pellet containing 100,000 cells in 10 µL resuspension buffer. Work quickly from this point on since resuspension buffer is slightly toxic and an increased exposure time results in a noticeably lower cell viability.
   6. Mix in 1/10 of the reprogramming plasmid solution (the solution aliquot was prepared for a total of 1 x 10⁶ cells).
   7. Mount a transfection tip onto the transfection pipette.
   8. Aspirate the cell suspension into the transfection tip carefully, avoiding formation of air bubbles. If bubbles are observed, expel the suspension and repeat the aspiration. Air bubbles will impede transfection.
   9. Insert the transfection pipette into the transfection tube and press the "START" button on the screen of the transfection device. Wait for the screen message informing about the success of the transfection and remove the pipette from the tube immediately.
   10. Expel the suspension into 1 well of the target 6-well plate prepared in section 2.2. Mix in the medium from a neighboring well and distribute the suspension equally into both wells (the resulting cell density will be 50,000/well).
   11. Repeat the transfection for all microcentrifuge tubes containing AFSC/AMSC individually. Place the plate into the incubator at 37 °C and 5% CO₂.
5. Culture of transfected AFSC/AMSC
   1. Culture the transfected cells for 2-5 days. Then switch to reprogramming medium consisting of E8 supplemented with 100 µM of sodium butyrate on day 3.
      NOTE: Secondary passage can be performed to avoid overgrowth of the source AFSC/AMSC. However, the passaging will disable the option to calculate the reprogramming efficiency correctly if this parameter is of interest.
   2. Change the reprogramming medium every day to every other day for 10 days. Change the medium every day from day 10 on.
6. Manual picking of fully reprogramed colonies for clonal expansion
   1. Fully reprogrammed colonies appear around day 14. Allow colonies to expand in size and become compact. They can be manually picked and transferred to fresh plates as early as day 15-16.
   2. 1 h before the picking procedure, coat 24-well plates with 8 µL of VTN in 300 µL of vitronectin dilution buffer per well (1 µg/cm²) and incubate at RT or 37 °C. Replace the solution with E8 medium without sodium butyrate.
   3. Select colonies of a sufficient size (ideally over 400 µm in diameter) in a sterile culture cabinet. A phase contrast microscope or a stereomicroscope can be used.
   4. For picking, an LCD imaging microscope placed in the cabinet will be used since its monitor eliminates the need for oculars. Sterilize the microscope stage with 70% ethanol.
   5. Using a regular phase-contrast cell culture microscope, select, mark, and note the number of colonies to be picked. This is important to make sure time is not wasted for this process during the actual picking.
   6. Fill a number of PCR tubes that is equal to or greater than the number of colonies to be picked with 30 µL of 0.5 mM ethylenediaminetetraacetic acid (EDTA) in PBS. Colonies will be placed into these tubes for partial dissociation before plating.
   7. Plan to pick 5 colonies at a time from the plates using a 10 µL pipette set to 2 µL. Hold the pipette tip at an angle at the colony edge and carefully and gradually scrape the whole colony off the surface. Immediately aspirate the whole colony into the pipette tip and transfer it into one of the prepared PCR tubes with EDTA.
   8. Repeat with the remaining 4 colonies. Incubate at RT for 4-6 min.
   9. Pipet the suspension up and down gently using a larger pipette tip to break the colony down into smaller clumps. Avoid creating a single cell suspension.
   10. Plate the suspension directly into a target well of a 24-well plate prepared in step 2.6.2. Repeat with the remaining colonies.
   11. Repeat steps 2.6.6 through 2.6.10 if more than 5 colonies are to be picked but do not pick more than 5 colonies at a time. Incubate at 37 °C and 5% CO₂.
7. Clonal expansion and maturation of iPSC
   1. Allow colonies to grow and become compact. 3-6 days are sufficient. Change the culture medium daily. Use between 400 µL and 1 mL of E8 medium based on the cell density.
   2. Wells of the 24-well plate with a sufficient colony density will be expanded into 6-well plates. 1 h before passaging, coat the 6-well plates with VTN (as in step 2.2.1). Then replace the solution in the well with 2 mL of E8 medium per well.
   3. Aspirate the spent medium from the source wells using a 1 mL pipette and replace with 300 µL of 0.5 mM EDTA to wash. Aspirate immediately using the same pipette tip and replace with 300 µL of 0.5 mM EDTA again, then incubate at RT for 5 min. Aspirate all liquid using a 1 mL pipette.
   4. Set the 1 mL pipette to capacity, mount a 1 mL wide-bore tip on it, and aspirate the E8 medium from the target well into the tip. Wash the source iPSC culture off with a stream of the medium.
   5. Transfer the suspension into the target well and pipet up and down several times to break up colonies into clumps of 20-50 cells. Make sure they get distributed evenly in the well by gentle rocking and shaking the plate and incubate at 37 °C and 5% CO₂.
   6. Change the medium daily and passage every 3 to 4 days. Any differentiating colonies can be marked under the phase-contrast microscope and removed using a pipette tip in the culture cabinet. This allows for selective propagation of high-quality pure iPSC culture.
   7. 1 h before passaging, coat the wells of a 6-well plate with VTN as in step 2.2.1. Then replace the solution with 2 ml of E8 medium per well.
   8. Routine passaging is similar to the initial passaging done using 0.5 mM EDTA (steps 2.7.2 to 2.7.5). Replace spent medium in one well of a 6-well plate with 1 mL of EDTA to wash and discard.
   9. Add 1 mL of EDTA and incubate at RT for 5-7 min for partial dissociation. Optimize the incubation time if necessary, making sure a suspension of around 20- to 50-cell clumps is produced. Avoid dissociation into single cells.
   10. Discard the EDTA solution and aspirate 1 mL of the E8 medium using a 1 mL pipette from the target well into a wide-bore pipette tip.
   11. Wash off the source iPSC culture with a stream of the E8 medium repeatedly until a desired portion of it was released from the surface and transfer into the target well. This portion represents the split ratio (e.g., 1/8 of the culture can be transferred for a 1:8 ratio.)
   12. Passage every 3-4 days. Allow iPSC lines to mature by culturing them for at least 15 passages before using them in downstream experiments

3. Characterization and Confirmation of Pluripotency

NOTE: Refer to the supplementary files for details on flow cytometry and confocal microscopy.

1. Teratoma formation assay
   1. To determine the capacity of iPSC to differentiate into tissues representative of all three germ layers by teratoma formation assay, follow the institutional policies regarding animal care and use, planning ahead to allow time for filing the appropriate protocol documents. The teratoma formation in mice will take between 6-10 weeks.
   2. Culture 4 wells of a 6-well plate per iPSC line for 4 days. The approximate number of cells injected into one flank of a mouse is 0.5 to 1 x 10⁶ cells. Optional: an extra well can be dedicated to determining a representative cell number in a well by means of single cell dissociation with a cell detachment enzyme and counting.
   3. Calculate the volume of the E8/BMM mixture the iPSC clumps will be suspended in: Both flanks of a mouse are injected, each with 150 µL of clump suspension. Three mice are sufficient to test teratoma formation of one iPSC line. Include an extra 150 µL per needle in the resulting volume to compensate for dead volume loss. 1 needle per mouse is used. The total volume is therefore 3 * (150 µL * 2 + 150 µL) = 1350 µL
   4. Partially dissociate the iPSC colonies with EDTA as if they were to be passaged, as described in the steps 2.7.8 to 2.7.10. It is important that the colonies be dissociated into clumps and not separated into single cells.
   5. Wash off the iPSC colonies with 675 µL of E8 medium (half of the calculated volume from step 3.3.3), using a wide-bore tip. Transfer the clump suspension into a 5 mL polystyrene tube. Place on ice.
   6. Combine with 675 µL of BMM. Keep the resulting suspension on ice until the injection.
   7. Anesthetize the mice to immobilize them using isoflurane. This should be performed or guided by skilled personnel of the vivarium.
   8. Vortex the 5 mL polystyrene tube briefly and aspirate the clump suspension (450 µL per mouse) into one insulin syringe equipped with a 22 G needle. Inject 150 µL of the cell suspension subcutaneously. This volume contains approximately 1 x 10⁶ cells (see step 3.1.2).
   9. Inject 3 mice per iPSC line. Follow proper animal care practice for all procedures.
   10. Monitor the health of the mice daily.
   11. When the teratomas reach an endpoint diameter of 1.5 to 2 cm, euthanize the mice, explant the teratomas, and store them in formalin solution for tissue fixation for 24 h.
   12. Bring the fixed teratomas to a histology core facility for hematoxylin eosin (H&E) staining. A pathologist will grade the presence of tissues of all three germ layers.
       Note for Karyotyping: Live iPSC cultures should be shipped to specialized cytogenetic laboratories for testing of the integrity of the karyotype. It is recommended that this testing be performed every 5 passages.
2. Transcriptional profiling
   1. Culture 2 wells of a 6-well plate for 3-4 days to obtain one RNA sample. Only use high-quality cultures, minor contamination with differentiating cells can be addressed by scraping them using a pipet tip.
   2. Isolate RNA from iPSC cultures using a commercially available kit following the manufacturer's protocol. Ship samples to a specialized genomic core facility.
   3. Obtain the global transcriptional profiles of the iPSC lines using microarrays (see Table of Materials for supported choices) or RNA sequencing.
   4. Submit "*.idat" files for bioinformatic assessment of pluripotency through an online interface at the Coriell Institute. Alternatively, submit "*.cel" files for bioinformatic identification of cell type, including pluripotent stem cells, through an online interface at Johns Hopkins University. See Table of Materials for details on the types of data the individual bioinformatic assays accept.