Human corneas with donor consent for research were obtained from the Queensland Eye Bank and used with ethics approval from the Metro South Hospital and Health Service's Human Research Ethics Committee (HREC/07/QPAH/048). Sheep corneas were obtained from euthanized animals at the Herston Medical Research Facility of the University of Queensland under a tissue sharing agreement.

1. Preparation of dissection tools

1. Sterilize two pairs of number 4 watchmaker forceps by either soaking them in a solution of 70% ethanol for 5 min or by autoclaving them.

2. Preparation of culture medium and tissue culture plates

1. Prepare 100 mL of culture medium containing Opti-MEM 1 (1x) + GlutaMAX-1, 5% fetal bovine serum and 100 U/mL Pen/Strep. This culture medium is adequate for sheep corneal endothelial cell cultures, however, for human corneal endothelial cell cultures the medium should be supplemented with 50 µg/mL bovine pituitary extract, 0.08% chondroitin sulphate, 200 µg/mL calcium chloride and 0.3 mM L-ascorbic acid 2-phosphate. Culture medium can be stored in the dark at 4 °C for two weeks.
2. Coat the wells of a 6-well tissue culture plate with Attachment Factor using the manufacturer's instructions and then add 1 mL of culture medium to each coated well. Prepare one well for each cornea.

3. Explant dissection and cell culture procedure

1. Place the cornea, endothelium side up, into a sterile Petri dish on the stage of a dissecting microscope. Adjust the illumination so that the corneal surface is well-lit with sufficient contrast to highlight the endothelial layer. Adjust the zoom so that the edge and some central corneal endothelium is in view.
2. Use sterilized watchmaker forceps to gently lift and tear Descemet's membrane away from the underlying stroma (Figure 1). The membrane will detach from the stroma as a strip that immediately curls up. Place the strip into one well of a 6-well tissue culture plate that has been coated with Attachment Factor and contains 1 mL of culture medium. The lid of the tissue culture plate should be kept on at all times, except when explants are being added to it, to reduce the risk of contamination.
3. Remove strips of Descemet's membrane from the extreme periphery of the endothelium first and then from central regions later. Place all strips from one cornea into a single well within the 6-well plate.
4. Incubate the explants in a humidified cell culture incubator set at 5% CO₂ and 37 °C. Leave the culture undisturbed for 2 days to allow the explants to settle and attach to the plate surface. Typically, up to one third of explants fail to attach to the plate.
5. Remove the medium and any unattached explants from the culture after 4 days and gently add 2 mL of fresh culture medium taking care not to disturb the attached explants. Culture medium should be changed twice per week.

4. Continuous production of corneal endothelial cells by serial explant culture

NOTE: Explants can be transferred to fresh tissue culture plates after 10 days to establish additional corneal endothelial cell cultures.

1. Place the explant culture onto the stage of a dissecting microscope and adjust the illumination and zoom so that the explants are visible.
2. Using sterilized watchmaker forceps, gently pluck each explant from its culture plate and transfer it to a fresh well of a 6-well tissue culture plate that has been coated with Attachment Factor and contains 1 mL of culture medium.
3. Allow the explants to settle onto the surface of their new plate for 4 days before replacing the culture medium with 2 mL of fresh culture medium. Change culture medium changed twice per week.

5. Growing corneal endothelial cells on glass coverslips for immunofluorescence analyses

NOTE: Cell cultures that are destined to be analyzed using immunofluorescence should be established on glass coverslips that can be mounted onto glass microscope slides following the staining procedure.

1. Sterilize a pack of round glass coverslips of 13 mm diameter in an autoclave or by soaking in 70% ethanol for 15 min followed by three rinses in phosphate-buffered saline (PBS).
2. Place individual coverslips into the wells of a 24-well tissue culture plate, coat them with Attachment Factor, and then add 0.5 mL of culture medium to each well.
3. Using sterilized watchmaker forceps remove explants from their tissue culture plates and transfer them to wells containing coverslips. Allow the explants to settle onto the coverslips for 4 days before changing the medium.
4. After the required incubation period, fix and stain the coverslip cultures within their culture wells. Mount the stained coverslips onto glass microscope slides for analysis under a fluorescence microscope.

6. Subculture of corneal endothelial cells using Dispase II

NOTE: Large fibroblastic cells can be selectively removed from explant cultures in 6-well plates before subculturing using this procedure. If all cells are to be subcultured, do not perform steps 6.2 to 6.4. The aim of this procedure is to transfer the cells to fresh plates while maintaining their cell-to-cell contacts as much as possible. The cells should be handled gently. Completely confluent wells should be passaged at a ratio of 1:2, while subconfluent wells should be passaged at a ratio of 1:1 or less.

1. Remove the medium from the culture and briefly rinse with DPBS.
2. Add 1 mL of Versene. Incubate at room temperature for 30 s.
3. Remove the Versene and add 1 mL of TrypLE. Incubate at 37 °C in the tissue culture incubator for 3 min.
4. Observe the culture using a phase contrast microscope. Gently tap the culture to dislodge cells from the plate. As soon as the large fibroblastic cells have detached from the plate remove them and the TrypLE using a 1 mL pipette. Wash residual TrypLE off the remaining cells by rinsing twice with 2 mL of DPBS.
5. Add 1 mL of 1 mg/mL Dispase II to the culture and incubate in the tissue culture incubator for 1 to 2 h or until all cells have detached from the plate. The cells should gradually float away from the plate in clumps.
6. Collect the cells using a 1 mL pipette and transfer to 20 mL of DPBS in a 50 mL centrifuge tube. Centrifuge for 5 min at 300 x g at room temperature.
7. Remove the supernatant and gently resuspend the cell pellet by trituration with a 1 mL pipette in 1 mL of culture medium. Transfer the cell suspension to either one or two wells of a 6-well plate, to passage at a ratio of either 1:1 or 1:2 respectively.
8. Top up the medium in each well to make 2 mL and place the cultures into the tissue culture incubator. Replace the medium with 2 mL of fresh medium twice per week.

7. Growth of corneal endothelial cell layers on biomaterial membranes

NOTE: The following procedure describes the steps involved in mounting a membranous biomaterial in a custom-made mounting device—called a micro-Boyden chamber—for cell culture. Please refer to our recent publication⁶ for further information about the device and for purchasing details.

1. Assemble the upper chamber of the micro-Boyden chamber by placing a red O-ring into its center.
2. Use a trephine of 18 mm diameter to punch out a disc from a biomaterial sheet on a polytetrafluoroethylene (PTFE) cutting board. Place this disc over the red O-ring in the micro-Boyden chamber's upper chamber.
3. Screw the lower chamber onto the upper chamber, securing the biomaterial disc in between.
4. Soak the assembled micro-Boyden chamber in 70% ethanol for 1 h to sterilize it.
5. Completely immerse the assembled micro-Boyden chamber in sterile HBSS for 10 min to remove the ethanol. Repeat this wash step twice. Perform a final wash step for 10 min in unsupplemented culture medium.
6. Transfer the sterilized micro-Boyden chamber to culture medium in the well of a 6-well plate ensuring that the upper chamber is uppermost. Adjust the level of culture medium so that it contacts the lower surface of the biomaterial membrane but does not flow into the upper chamber.
7. Prepare a suspension of corneal endothelial cells using the procedure outlined in section 6. Sufficient cell density in the suspension should be achieved to allow a seeding density of at least 100,000 cells per cm² on the membrane in the micro-Boyden chamber. The culture surface area within the micro-Boyden chamber is 0.5 cm² and it can hold a volume of 100 µL. Therefore, a suspension containing 500,000 cells/mL should be prepared.
8. Pipette 100 µL of cell suspension (50,000 cells) onto the membrane in the micro-Boyden chamber. Incubate in the tissue culture incubator for 4 h before topping up the medium to completely submerge the chamber. Incubate the culture for the required period of time, replacing the medium twice per week.
   NOTE: Once the corneal endothelial cells have attached to the upper surface of the biomaterial membrane the micro-Boyden chamber can be flipped over within the culture well so that the lower chamber is uppermost. More cells can then be added to the chamber to initiate cell cultures on the other surface of the membrane. For example, corneal stromal cells may be readily applied to the alternate surface thus mimicking the relative location of corneal cell types as seen within the posterior cornea.