Generation of ESC-derived Mouse Airway Epithelial Cells Using Decellularized Lung Scaffolds

The overall goal of this procedure, is to direct the differentiation of mouse embryonic stem cells into mature airway epithelial cells, using a 3D culture set up with decellularized lung scaffolds.This method can help answer key questions in the area of lung development, such as the role of cell matrix contractions and growth factor signalling during lung lineage differentiation.The main advantage of this technique is that it uses a 3D culture system in a defined serum free setting, and achieves robust airway epithelial differentiation with limited contamination from other endoderm lineages.After euthanizing an adult wistar rat, according to the text protocol, secure the animal to a dissecting surface by fixing forepaws and hindlegs, and use 70%ethanol to spray down the chest and abdomen.Create an incision just below the rib cage.Cut through the skin and expose the diaphragm.Then, make small incisions through the diaphragm, to cause the lungs to retract, reducing the chance of puncturing the lungs.Next, create a vertical incision along the sternum, and open the thoracic cavity to access the heart and lungs.Then with a suture, ligate the inferior vena cava, and use small dissecting scissors to place a small incision in the left atrium.Using a prepared 10ml syringe with a 25 gauge needle filled with heparinized Hank's Balance salt solution, start lung perfusion by inserting the needle into the right ventricle, and push the buffer through the pulmonary circulation.Continue the perfusion until the lungs turn white, and the fluid flowing from the left atrium runs clear.Following perfusion, expose the trachea, and make a small incision near the thyroid cartilage for canulation.Then, with a plastic catheter, canulate the trachea and use a suture to secure it in place.Now, set up a gravity perfusion system by fixing a 10ml syringe to a retort stand and clamp.Then, remove and discard the syringe plunger.Secure the maximum filling point on the syringe barrel at 20 cm above the lungs, then attach a two way stopcock to the end of the syringe, and a long plastic tubing to the other end of the stopcock.Gently pull out the catheter from the trachea, and attach to the end of the plastic tubing.Then, pour decellularization solution into the syringe and allow the solution to completely fill the attached plastic tubing and catheter.Place the solution filled catheter back into the trachea and lavage the lungs by filling to total lung capacity for one minute.Then remove the catheter from the trachea to allow the fluid to flow out of the lungs.Repeat the lavage of the lungs eight times with decellularization solution, followed by ten rinses with PBS.The lungs will appear white with the completion of the lavage steps.Then dissect the trachea and lungs from the neck and chest cavity.Remove excess tissue from the dissected lungs, including the esophagus and heart, then transfer the lungs into cold PBS until preparation for vibratome sectioning.To prepare thick sections, make approximately 15ml of 2%and 4%low melting point agarose in PBS, and enough 6%agarose for embedding all lobes into small rectangular blocks.After transferring the agarose to 15ml tubes, place the tubes on a heat block and maintain the temperature above 40 degrees celsius to avoid gelling.Using small scissors, dissect the decellularized lung at the end of each lobar bronchus, and detach each lobe.Then use absorbent sheets to remove excess PBS, and transfer the lobes to 2%agarose on the heating block for five minutes to coat the tissue.After transferring each lobe to a petri dish, place the dish on a cold plate and allow the surface to gel for one minute.Gently place the lobes into 4%agarose, and after incubating for five minutes and cooling, repeat the coating with 6%agarose.After coating, use metal base molds to embed each lobe separately in 6%agarose, with at least 3ml of agarose surrounding the tissue from the edges.Next, with forceps, orient each lobe by positioning the tissue's largest flat edge at the surface of the metal mold facing the experimentor, and complete the embedding using plastic cassettes.Allow the blocks to gel on a cold plate for at least 30 minutes before vibratome sectioning.Alternatively, store the blocks in a humidified chamber for up to 12 hours at four degrees celsius prior to vibratome sectioning.To prepare sections, set up the vibratome by filling the sectioning chamber with cold PBS.Set up a surrounding ice bath to maintain a cold temperature throughout sectioning.Now, remove the blocks from the metal molds, and use a razor blade to trim down the excess agarose surrounding the lobes, while keeping approximately three millimetres from the edge of the tissue.Using adhesive, fix the tissue to the center of the specimen plate, and submerge the plate into the PBS filled sectioning chamber.Load the sectioning blade on the vibratome.Set the sectioning boundaries on the vibratome, by selecting the following speed, amplitude, and thickness values respectively.Section each lobe completely, and if sections do not fully separate from each other, use small scissors to manually cut the sections free.Then, gently collect scaffold sections, and transfer into cold PBS.Using forceps, gently free each section from the surrounding agarose.To decontaminate scaffold sections, after transferring them from PBS to microcentrifuge tubes, add 90 units per milliliter of nuclease in PBS, and incubate on a rotator at room temperature for 12 to 24 hours.After transferring sections to new microcentrifuge tubes, according to the text protocol, under sterile conditions add antimicrobial solution, and incubate on a rotator at room temperature for six hours.Then under sterile conditions, use PBS to rinse the scaffolds twice and transfer them into serum free medium, or SFDM, prior to seeding with cells.To set up an air liquid interface culture, use sterile forceps to place hydrophobic membranes on a petri dish.Then, transfer each decellularized scaffold section from SFDM onto a membrane, ensuring the sections are spread evenly on the membrane.Prepare six or twelve well plates by filling wells with 1ml or 0.5ml of SFDM respectively.Then, gently place the membranes into the wells, allowing the membrane to float on top of the medium creating an air liquid culture set up.To seed 3D scaffolds, following the preparation of endodermal cells and fluorescence activated cell sorting, to enrich for a definitive endoderm, use a haemocytometer to count the sorted cells.Then spin down the cells at 400 Gs for five minutes.Using SFDM, re-suspend the pelleted cells to approximately 100, 000 cells in 10