Primary Cell Cultures from the Mouse Retinal Pigment Epithelium

The overall goal of this procedure is to demonstrate a feasible way to obtain retinal pigment epithelium cells from mouse eyes, and to culture them in vitro while retaining their original properties. This method will help RPE researchers to obtain dissection skills that are required for establishing of mouse RPE cell cultures, and to demonstrate how to do that in an easy way. The main advantage of this procedure is that it is simple and feasible.

The only thing required is practice, and the visual demonstration of RPE isolation and the microscopy literally demonstrates how to obtain the required skills. To isolate the mouse eyeballs, first euthanize two mice using any approved method, and wearing gloves place it on an absorbent pad. Position a thumb and index finger around the eye and gently push the skin to bulge out the eyeball.

Once the eyeball is protruded, tuck the tip of a pair of angled scissors between the skin and the eyeball and carefully cut the tissue around the eye until it is released. Then, briefly dip the isolated eye in the 70%ethanol and submerge it in PBS in a Petri dish. After collecting both eyes from each of the mice, proceed with the dissection.

Under the dissecting microscope, carefully remove the majority of connective tissue from around the eye. It is not necessary to keep the eye submerged during this step. To do so, use suturing forceps to lift the connective tissues from the eyeball, and cut them out using Vannas scissors.

Do not cut the sclera because it is practically impossible to obtain the intact posterior eye cups if the sclera gets cut. After cleaning off the connective tissue, submerge the eyeballs in PBS, and switch to working in a laminar flow cabinet under sterile conditions. Now, use forceps to transfer the eyes into a dish of warm DMEM containing a high concentration of glucose.

After 20 to 30 seconds, replace the wash solution using aspiration for a second wash. Transfer eyeballs to warmed 2%Dispase II working solution in 15 mL tubes and let the eyes incubate at 37 degrees celsius for 45 minutes to dissociate. After the incubation, the eyeballs should be soft.

Next, wash the eyes two times in the same vessel with warmed growth medium. After the second wash, replace the growth medium with fresh medium, and transfer the eyeballs and the medium to a new dish to continue the dissection. Continue working in a clean bench with the help of a dissection microscope.

Start the dissection by securing the eye with forceps and make an incision around the ora serrata using Vannas scissors. Keeping the eyeball in the solution, carefully remove the anterior cornea by extending the cut around the ora serrata circumference. After completing the cut, pull off the anterior cornea.

Next, remove the lens capsule and associated iris pigmented epithelium by gently pulling it out of the eye cup using toothed forceps. Then, repeat the dissection on the remaining eyes and incubate the posterior eye cups in growth medium for 20 minutes at 37 degrees celsius to facilitate separation of the neural retina from the RPE. After 20 minutes, cut the posterior eye cups into four petals.

Make the cuts long enough to flatten the eye cup, but short enough to keep the petals connected. Next, flatten the eye cup to remove the neural retina. Secure the remaining RPE choroid sclera complex with the non-dominant hand and very gently tug away the retina from the edges.

Once the neural retina is removed, transfer the quartered RPE choroid sclera complex into a new sterile culture dish filled with warm growth medium. In the Petri dish, secure one petal of the quartered RPE choroid sclera complex with super fine forceps, and using a microsurgical crescent knife, gently peel off the intact sheets of RPE from the underlying basement membrane. The brownish RPE should be clearly distinguishable from the dark, sticky, fluffy choroid.

Then, wet a p200 micropipette tip with growth medium and use the tip to transfer the RPE sheets into the new dish containing warmed growth medium. Now, wash the RPE sheets three times in warmed growth medium. After each wash step, carefully collect the RPE sheets while avoiding other tissues, such as the debris from the retina or choroid.

After the last wash, transfer the RPE sheets to a 1.5 mL tube, and allow them to sediment by gravity. Then, in the laminar flow cabinet remove the extra growth medium and re-suspend the cells in freshly made warm growth medium. Now, gently triturate the tissue using a p200 micropipette without making bubbles.

Triturate between 40 and 50 times, just enough to make a single cell suspension. Be careful as too much trituration can be lethal. Now, plate the RPE cells collected from both mice in one 12 mL polyester membrane insert in the well of a 12 well culture plate.

A high cell density is desired so that the RPE cells retain their hexagonal shape and pigmentation. A mouse primary RPE culture established from two mice in a 12 mm polyester membrane insert reached 90 to 95%confluency after one week in culture. After two weeks in culture, the cells were 100%confluent and started to form a mosaic of hexagonal pigmented and bi-nucleated cells.

By week three, the cells continued to change in shape and pigmentation. However, after four weeks a portion of the cells became hyperpigmented. The purity of the primary RPE cell culture was assessed using the RPE65 antibody, which labels retinoid isomerohydrolase, an enzyme critical to the vertebrate visual cycle.

The integrity of the cell to cell junctions and hexagonal cell shape was demonstrated by staining with phalloidin. Tight junctions were visualized by analysis of ZO-1 protein expression, which could be visualized using an amino stain and validated by western blotting. Overall, the cultures are able to proliferate, retain pigmentation and their hexagonal shape while forming tight junctions and expressing functional markers, such as RPE65.

After watching this video, you should have a good understanding of how to isolate RPE cells from the mouse eyes and culture them properly in vitro. Once mastered, this technique can be done in three hours if it is performed properly. While attempting this procedure, it is important to remember to always keep the eyes wet and try to operate as fast as you can.