An Ex Vivo Explant Model for Studying Glial Interactions in the Mouse Retina

We want to understand neuroinflammation in glaucoma, the leading cause of irreversible blindness worldwide. Specifically, we're studying how glial support cells in the retina influence neuronal loss during disease progression.

Glia such as astrocytes and microglia are thought to influence glaucoma progression, but many of the tools used to study neuronal function in live animal models are ill suited for these cells. Retinal explants are used to study neuroinflammation and glial function, but the learning curve is steep. Our protocol makes it more approachable and hopefully enables wider adoption of the technique.

Compared to traditional in vitro cell culture, our explant approach better preserves the natural environment for cells in the inner retina, enabling more accurate investigation into their physiological functions.

[Narrator] To begin, place the extracted mouse eye in a dissection dish filled with sterile, room temperature PBS. Identify an appropriate holding point and grasp it with angled forceps, then gently position the eye on the submerged lab wipe while ensuring the anterior-posterior axis from the cornea to the optic nerve is positioned horizontally. While maintaining a firm hold with angled forceps, use the tip of a number 11 scalpel to make an incision parallel and approximately 0.5 millimeters posterior to the limbus where the cornea transitions to the sclera. Insert one blade of the spring scissors inside the globe and cut circumlimbally around the eye, repositioning as needed with forceps. After completing the circumlimbal cut, remove the anterior segment and lens with forceps. If a long piece of optic nerve remains, trim it to a length of one to two millimeters using fine scissors. Then rotate the eye cup so that it faces upward to enable visual inspection and facilitate vitreous removal. Continue using angled forceps to immobilize the eye cup and inspect the retina for visible damage and examine the vitreous chamber for pigmented cell debris from the retinal pigment epithelium or choroid. Using a modified transfer pipette, flush the vitreous chamber with PBS, keeping the pipette tip submerged to prevent air bubbles. Then, with a fine watercolor brush, gently remove larger debris while minimizing contact with the retina. For persistent debris, use fine-tipped forceps carefully, avoiding direct metal contact with the retina. After clearing visible debris, flush the vitreous chamber with PBS from the transfer pipette three to five times and use the fine brush to probe near the periphery for residual ciliary body elements, detecting vitreous by the drag on brush fibers. If pockets of vitreous remain, sweep outward toward the periphery with the brush, keeping fibers trailing at a shallow angle to prevent retinal damage. Hold a pair of forceps in a closed position with the tips touching and gently insert them between the retina and choroid using any natural gaps formed during handling. Use the flat arms of the forceps to gradually enlarge the space between the retina and choroid until full separation is achieved. Continue stabilizing the sample with one pair of forceps while using a second pair to gently pull the eye cup, including the sclera and the choroid, downward. If the retina descends with the eye cup, use the forceps to gently probe and detach any remaining points of connection, avoiding the optic nerve head. Then with the retina still anchored at the optic nerve head continue holding the tissue steady and use the second pair of forceps to bunch up the eye cup below the optic nerve head. Inspect to confirm that the retinal periphery is not folded due to residual vitreous, particularly at sites where ciliary body remains. If needed, flush the chamber with PBS using a transfer pipette and use the brush to gently uncurl any folded retina and remove excess vitreous. After the retina is exposed from both sides, use spring scissors to make a series of relieving cuts approximately 90 degrees apart from the retinal periphery toward the optic nerve head. While holding the submerged tissue, use forceps to laterally pull the lab wipe away from the sample and remove it from the dish without touching the retina. While holding the retina in place, use the spring scissors to sever the optic nerve just beneath the retina. Then carefully lift and remove the remaining eye cup tissue from the dish. Now fill a 35-millimeter Petri dish with PBS and use forceps to place a filter square at the bottom with the rough, matte side facing upward, avoiding any creases. Then, using the transfer pipette, gently aspirate the retina and transfer it into the Petri dish. Use the brush to orient the retina with the inner surface facing upward and position it directly above the filter square. Slowly aspirate PBS to lower the retina onto the filter. Once the retina is seated on the filter, use the brush to gently unroll any peripheral folds. Adjust the PBS level to balance retinal stability and hydration, allowing smooth brush movement without drying the tissue. Now use the transfer pipette to drip PBS from about one centimeter above to rinse the surface and inspect the retina again for debris. Close the lid of the 35-millimeter dish and carry it to the biosafety cabinet. Place the closed dish inside the biosafety cabinet without contacting any interior surfaces or equipment. Sterilize or replace gloves when transitioning to aseptic work and transfer the six-well plate preloaded with explant media from the incubator into the biosafety cabinet. Within the cabinet remove the lid of the 35-millimeter dish and use angled forceps to lift the filter square without touching the retina. Then open the six-well plate and gently lower the filter onto the center of the insert in one well, immersing it slowly into the media. Once the retina separate from the filter, slowly move the filter aside and remove it from the well using angled forceps. Use a one-milliliter pipette to aspirate 500 microliters of media from the insert, trapping the retina between the insert and the air-liquid interface. Finally, replace the lid on the six-well plate and return it to the incubator, ensuring the retina remains centered within the well. To examine large-scale changes in retinal glia, three-day explanted retinas were compared with sham explants fixed immediately after isolation instead of being cultured. Microglia in sham retinas showed a regular, non-overlapping distribution pattern, whereas by day three the explanted retinas' organization became irregular with cells appearing clustered, implying migration. Retinal astrocytes in sham retinas displayed close alignment with the vasculature, which diminished significantly after three days in vitro. GFAP expression in Mueller cells was faint or absent in sham retinas but became distinctly visible by day three in vitro, especially near tissue edges. After one day in vitro, microglia exhibited process retraction and early signs of activation which progressed to a compact amoeboid morphology by day three. At the 24-hour mark, retinal ganglion cell density quantified using Brn3a showed a modest but marked decline in culture explants over sham ones. TMEM119, a homeostatic microglial marker, was highly expressed in sham retinas but was nearly undetectable after three days in vitro. CD206 expression, marking hyalocytes, remained stable after three days of in vitro culturing. GFAP staining revealed astrocyte and Mueller cell reactivity around sites of mechanical injury sustained during dissection and handling.