Cultivate Primary Nasal Epithelial Cells from Children and Reprogram into Induced Pluripotent Stem Cells

The following protocol follows the guidelines of the institutions human research ethics committee.

1. Sampling the Nasal Mucosa

NOTE: Obtain samples from subjects who are free of signs of respiratory viral infection.

1. Prepare a 15 ml conical fresh before the participant visit and add 2 ml BEGM (Bronchial Epithelial Cell Growth Medium) plus 20 µl sterile Penn/Strep/Fungizone (P/S/F) (0.01%).
2. Open cytology brush just before taking the sample, and be sure to keep brush sterile and do not touch it to any surfaces besides the nasal surfaces.
3. Ask participant to sit still and tilt head up towards ceiling. Have smaller or more restless children sit on their hands and/or lying down to avoid swatting brush away.
4. Aim brush at the back of the nose where the passage narrows (Looks like a small black hole). Slide brush down and twist wrists as the brush is removed from nostril.
5. Place brush in conical and submerge in BEGM. Cut off excess brush and replace cap on tube. DO NOT VORTEX.
   NOTE: If the participant is willing, obtain a second brush in the same manner but in the second nostril. Obtaining a brushing of both nostrils will result in a higher likelihood that the sampling will be successful. Sampling the same nostril may result in bleeding and will not likely improve the sample.
6. Keep samples as close to 37 °C as possible during transport using a beaker with warm water.

2. Cell Count and Cytospin

1. Gently agitate the brush in BEGM and take 10 µl of the sample for a cell count using a hemacytometer. Add 10 µl of a live/dead stain and count only live cells under a microscope.
2. Take 50 µl of the sample and dilute to 200 µl with 1x PBS. Add to cytospin funnel attached to glass slide and spin at 300 rpm for 2 min. Let slide dry O/N and stain slide following manufacturer's instructions. Allow to dry O/N, preferably in the dark.
3. Stain Slide with Hema 3 Stain
   1. Transfer each solution (Hema fixative, light blue; Hema 3 Solution I, pink; and Hema 3 Solution II, deep blue) into a staining dish; Keep covered when not in use. Insert slides into slide staining boat
   2. Dip slides continuously in fixative for 30 sec, then allow excess to drain. Dip slides continuously in Solution I for 30 sec, then allow excess to drain. Dip slides continuously in Solution II for 30 sec, then allow excess to drain
   3. Rinse with deionized water until water runs clear. Allow to dry O/N, preferably in the dark
4. Look at slide under microscope and count cells, determine percentage of epithelial cells. Should be 90% or greater if sampling is good.

3. Seeding Cells

NOTE: Carry out all cell culture procedures in a proper and certified tissue culture hood using sterile technique.

1. Coat plates with Bovine Dermal Collagen (BDC), type 1. Add 0.5 ml of 3 mg/ml BDC diluted in 49.5 ml 1x PBS. Prepare less if only a few plates will be used. Coat a T25 flask with 2 ml BDC and incubate in sterile hood O/N or at 37 °C for 2 hr, if necessary.
2. After incubation, aspirate any remaining liquid. Expose flasks to UV light for 30min in sterile hood. Store flasks for up to one month at 4 °C, but bring them to RT or 37 °C before seeding cells.
3. Keep brush and cells in BEGM at 37 °C until ready to seed. Gently swish brush inside conical, but do not vortex.
   1. Plate 7 x 10⁵ cells in a T25 flask. If there is less, use a smaller container, such as one well of a 6-well plate. This size plate requires about 3 x 10⁵ cells.
      NOTE: If there are not this many cells, it is not advisable to continue.
4. Under a sterile hood, take brush out of tube and gently rotate brush on the surface of the coated flask, being careful not to scratch the coating. Discard brush in an appropriate biohazard container.
5. Slowly pipette the remaining cells in BEGM out of the conical and into the T25 flask. Observe the cells under microscope. A number of floating cells will be observed and there may be some debris from the nose, which is acceptable at this stage (Figure 3A).

4. Cell Culture

1. After seeding cells, let them settle for 48 hr without disruption (Figure 3). After 48 hr, slowly and gently add 2 ml warm BEGM and 20 µl sterile Penicilin/Streptomycin/Fungizone (P/S/F).
   NOTE: DO NOT aspirate the original 2 ml of media.
2. On the 4^th day after seeding, carefully aspirate all liquid, and replace with 4 ml fresh, warmed BEGM plus 1x Pen/Strep (fungizone is no longer necessary). Change media every two days and assess cells for attachment, growth, and confluence. When cells reach ~80% confluence, usually in 2-3 weeks, they are ready to be passaged.
3. One T25 flask (about 2.5 x 10⁶ cells when confluent) can be passaged into three T25 flasks or one T75 flask. After the first passage, cells should be robust and healthy, reaching confluence about every three days.

5. Passaging Cells

1. Gently aspirate the media from the plate. Add 1 ml 0.05% Trypsin solution per T25 flask to the plate and place in the incubator for about 4min, or until cells detach from plate. Check the level of detachment of cells every 2 min and do not leave Trypsin on longer than 10 min.
2. Add 5 ml Trypsin Neutralizing Solution (TNS), or serum-containing media to neutralize the enzyme. Remove all the liquid and cells from the flask and place in a 15 ml conical tube.
3. Centrifuge at 200 x g (acceleration 0 deceleration 0) for 5 min to obtain a cell pellet. Aspirate supernatant and resuspend cells in BEGM + P/S in the volume required for the new flasks (4 ml for a T25 flask, 10 ml for a T75 flask)
4. Perform cell count with a live/dead stain as described above (2.1). Add appropriate amount of BEGM and cells to the coated flasks and return to the incubator. Maintain as detailed above or cryopreserve in freezing medium (70% BEGM, 20% FBS and 10% DMSO) at a concentration of 0.5-2 x 10⁶ cells per ml.

6. Reprogramming to iPSCs

NOTE: Before generating iPSCs, ensure adherence to all institutional regulations governing the generation and use of human iPSCs. Sterile technique is especially important for iPSC cultures, as culture medium does not routinely contain antibiotics. It is critical that NECs appear healthy and are robustly proliferative for successful reprogramming.

1. Plate 5 x 10⁵ NECs per well of a 6 well tissue culture plate. After 24 hr, add polybrene to BEGM to a final concentration of 8 µg/ml and transduce NECs by adding lentiviral particles expressing Oct4, Sox2, Klf4, c-Myc to the media. Aim to achieve a multiplicity of infection (MOI) of ~5²⁷.
   1. To determining the MOI, prepare serial dilutions of the concentrated-lentiviral stock and transduce HT1080 cells with these dilutions. After first unequivocally detection of dTomato expression in the HT1080 cells, calculate the number of "transducing units/ml" by scoring the number of dTomato-positive cells/small clumps of cells in each dilution.
      NOTE: Typically, there will be dilutions that are too high (everything is dTomato positive) and too low (none or only a couple of positive cells).
   2. Perform scoring from the dilutions in which discrete positive cells/small clumps of cells is identified. Determine the titer by correcting the number of transducing units/ml with the appropriate dilution factor. MOI is then the ratio of the number of transduced cells to the number of virus particles ²⁷.
      NOTE: Each dTomato positive cell/small clump is assumed to arise from a single virus particle.
2. About 3 hr post-transduction, remove media and replace with fresh BEGM. Discard lentivirus-containing media using institutionally-approved procedures. Return transduced NECs to the incubator for 3 days.
3. On day 3, replace media with fresh BEGM, and then incubate for another 3 days. Aspirate spent media, wash cells with 1x PBS and add 1 ml 0.05% Trypsin solution to the well. Passage cells with Trypsin as written above (section 5). Carefully aspirate supernatant and resuspend cells in 4 ml BEGM + P/S. Cells can be passaged to a new well of a 6 well plate coated with BDC, or added to MEF cultures, if ready (see next step).
4. Prepare MEF coated plates. On day 4 add 0.1% gelatin solution (1 ml/well) to 2 wells of a 6 well (to perform +/- Thiazovivin (SPT) see step 6.6). On the next day, thaw a vial of inactivated MEFs ²⁸.
5. Place MEFs into 10 ml of MEF media (DMEM + 1x NEAA + 10% dFCS). Spin at 200 x g for 5 min to pellet. Resuspend in MEF media. Aspirate gelatin from 6 well plate and add 1.87 x 10⁵ MEFs per well of the gelatin-coated 6 well plate. Incubate O/N at 37 °C, or for a minimum of 24 hr.
6. Transfer 2 ml of BEGM containing transduced cells per well into 2 wells of a previously coated 6 well plate and incubate O/N. On the next day, replace BEGM with 2.5 ml per well of standard hESC media containing 4 ng/ml basic fibroblast growth factor. Feed cells with fresh hESC media +/- SPT daily for 10 days
   NOTE: It is possible to add a cocktail of small molecules (SB431542, PD0325901, and Thiazovivin (SPT) cocktail) to enhance the efficiency and kinetics of reprogramming for NECs ²³.
7. After ten days, continue to feed daily using hESC media without SPT. Feed cultures daily until hESC-like colonies appear (Figure 5A, P0 colony). Identify colonies harboring putative iPSCs by their high nucleus:cytoplasmic ratio and prominent nucleoli.
8. Manually excise and transfer colonies with hESC-like morphology to separate wells for culture and expansion as separate lines in a feeder-free system. Excised colonies should adapt rapidly to these conditions.
   1. After plating excised colonies these discrete putative iPSC lines are now considered passage 1 (p1). Feed cultures daily with mTeSR1. Passage and expand iPSC lines using standard procedures. It may take several days to a week for p1 colonies to reach the size at which they should be passaged.

7. Maintaining iPSCs in Culture

1. Feed and evaluate cultures daily. Manually excise areas exhibiting overt differentiation by scraping with a sterile glass pipette or micro pipette tip. Change media after daily picking.
2. Passage cells after approximately every four days: Gently aspirate media, add 1 ml warm Dispase (1 mg/ml) per well of a 6-well plate (half the feeding volume), then incubate at 37 °C for about 4 min. The edges of the colonies will begin to lift, which looks like a white outline around the colony. Gently aspirate the Dispase and wash 3x with warmed DMEM/F12.
3. Add 2 ml mTeSR1 and use a cell lifter to lift the colonies from the plate. Use a 5 ml serological pipette or P1000 micropipette to gently triturate the cells until the colonies are broken into smaller clusters.
   1. Avoid producing very small clusters or single cells, as survival and subsequent proliferation will be sub-optimal. When re-plating colonies, a 1:4 to 1:6 split will maintain optimal colony density. Gently remove colonies and media from the plate and add to new, matrigel-coated wells.