Rapid, Directed Differentiation of Retinal Pigment Epithelial Cells from Human Embryonic or Induced Pluripotent Stem Cells

The overall goal of this method is to effectively direct the differentiation of pluripotent stem cells into retinal pigment epithelial cells in a homogenous layer. This method helps answer questions in the field of retinal and eye research such as how the retina develops and how to establish a stem cell disease model. The main advantage of this technique is that it's faster and more efficient than spontaneous methods.

This method uses a combination of growth factors and small molecules to direct the differentiation of stem cells to become immature RPE in 14 days and mature functional RPE after three months. Demonstrating portions of the procedure will be Cassidy Arnold, the associate director of the laboratory for stem cell biology and engineering at UC Santa Barbara. To begin this procedure, grow stem cell colonies in feeder-free, serum-free conditions to approximately 80%confluence before passaging.

Then coat a 12-well plate with extracellular matrix-based hydrogel as per manufacturer recommendations and allow it to set for one hour at room temperature or overnight at four degrees Celsius. After that aliquot the volume of retinal differentiation media and PBS without calcium or magnesium needed for day zero and warm them in a water bath to 37 degrees Celsius. Next add the growth factors to the warmed RDM and bring EDTA to room temperature.

If necessary, remove all the differentiated colonies using a P10 pipette tip based on morphology from the stem cells that will be passaged for differentiation. Fibroblastic cells between colonies as well as the opaque cells within colonies indicate the differentiated cells to be removed. Manual removal of cells is one of the most critical steps of the protocol because starting with a completely undifferentiated stem cell population will allow for a more efficient enrichment.

Next passage a single well of a six-well plate into four wells of a 12-well plate by aspirating the medium from the stem cells and washing the wells once with two milliliters of prewarmed PBS. Afterward aspirate the PBS and rinse each well three times with one milliliter of EDTA per well. Then gently tilt the plate and aspirate the EDTA.

Do not agitate the plate in any way to avoid prematurely lifting the cells. After the third wash, add one milliliter of EDTA again and incubate the plate at room temperature in the hood for three to five minutes. Do not disturb the plate during this incubation.

Following that, aspirate the EDTA and add one milliliter of RDM per well that will be seated plus 0.5 milliliters of extra medium. For example, wash one well of a six-well plate with 4.5 milliliters of RDM to plate on four wells of a 12-well plate. Then, use a cell scraper to gently detach the cells.

Triturate the cells in RDM by pipetting up and down five times in the well and complete this step quickly to prevent reattachment to the plate. Place in a conical tube. Dissociate large clumps of cells but do not triturate to single-cell suspension.

Distribute the cells evenly in the pipette and complete this step quickly to prevent reattachment to the plate. If desired, use a light microscope to observe the size of the cell clumps. Subsequently, seed the cells on ECM-coated, 12-well plates.

Tilt the plate back and forth to distribute the cells evenly throughout the wells and place it in a cell culture incubator at 37 degrees Celsius and 5%CO2 until the next medium change. In this procedure, coat a six-well plate with growth factor reduced ECM as per manufacturer recommendations. Allow it to set for one hour at room temperature over overnight at four degrees Celsius.

Then aliquot the amount of DPBS needed and one milliliter of RDM per well of enrichment and warm them to 37 degrees Celsius in a water bath. Bring the trypsin-like enzyme to room temperature and warm the required amount of RPE supporting medium to 37 degrees Celsius. After that, add an antimicrobial reagent and ROCK inhibitor to the RSM to achieve a final concentration of 0.5x antimicrobial and 10 micromolar ROCK inhibitor.

Use this medium for the first four to seven days to improve attachment. Next, aspirate the spent medium from all the wells and add one milliliter per well of prewarmed RDM without growth factors. If necessary, dissect and scrape away all the non-RPE cells using a P10 pipette tip under a dissecting microscope.

After dissection, aspirate RDM and all the cell debris and wash them twice with one milliliter of prewarmed DPBS per well. Subsequently, add 0.5 milliliters of TDE per well of a 12-well plate and incubate at 37 degrees Celsius for five minutes. Afterward, use a cell scraper to gently remove cells from the plate.

Gently triturate the cell enzyme suspension by pipetting up and down three to four times to create a uniform suspension. Next, dilute the cell enzyme suspension at a ratio of one to 10 in the prewarmed RSM without ROCK inhibitor. Centrifuge the cell suspension at 173g for five minutes at room temperature.

After five minutes, aspirate the medium from the cell pellet and resuspend the cells in RSM with 10 micromolar ROCK inhibitor. Next, strain the cells using a nylon mesh cell strainer with 40 micrometer pores. Count the number of cells using a hemocytometer and calculate the concentration of cells in the strained solution.

Afterward, seed the cells on the growth factor reduced, ECM-coated plates at one times 10 to the five cells per square centimeter in four milliliters of RSM with 10 micromolar ROCK inhibitor. Replace the RSM with 10 micromolar ROCK inhibitor 48 hours after cell seeding and continue to replace the media every three to four days. Stop adding ROCK inhibitor to the RSM after four to seven days.

Allow the cells to mature for 28 to 35 days at 37 degrees Celsius and 5%CO2 and continue to replace the RSM every three to four days. Shown here are the induced pluripotent stem cells immediately before passaging for differentiation. The stem cells are small and densely packed within the colonies which do not appear to contain differentiated cells such as fibroblasts between or opaque passages within colonies.

Here are the immature RPE cells at day two. The cells at this stage should still be subconfluent and extending projections into the empty spaces between the cells. These cells are before pick to remove enrichment on day 14.

Non-RPE cells appear as patches or opaque ribbons. The round patch contains cells with a fibroblastic morphology whereas the opaque patches are thought to be neural retina. These images show RPE at passage zero, one and three on day 30.

At each passage, the cells should be confluent and appear to have the hallmarks of retinal pigmented epithelial morphology including phasebright borders and a polygonal shape. Once mastered, the passaging and enrichment techniques can be completed in one to two hours depending on the scale of the cell culture. Following this procedure, techniques such as gene or protein quantification and functional assays can be used in order to answer questions about RPE's ability to secrete growth factors or phagocytose outer segments.

Papers published by Dave Buchholz in 2013 and Lindsay Leach in 2015 detail the development and optimization of this method. We have provided a detailed and thorough demonstration of each step in the procedure to make it readily available to researchers in the field. Happy experimenting.