Begin by positioning a euthanized mouse for eye enucleation using curved forceps to press the tissue around the eye to displace the eye out of the socket. Then lift and remove the eye, and transfer it to fresh PBS in the dissection tray. Using ultra fine scissors, cut the optic nerve as close as possible to the eyeball, and carefully insert fine tip straight tweezers into the eyeball through the optic nerve's exit at the posterior of the eye.
Now, insert scissors at the posterior of the eye and begin making an incision from the posterior toward the corneal scleral junction, and continue the incision until half of the junction has been separated. Gently push on the cornea so that the lens can exit through the incision. Using fine tip straight tweezers, carefully remove any large pieces of tissue from the lens.
After finding the equatorial region, shallowly pierce the lens, and then remove the lens capsule. Transfer the lens fiber cells to a 60 millimeter dish with 1%paraformaldehyde. Using a sharp scalpel, split the ball of the fiber cells in half along its anterior posterior axis.
And then further cut the halves along the same axis to produce quarters. Use the straight tweezers to remove the nucleus region from the lens fiber cell quarter. Next, add 200 microliters of 1%paraformaldehyde to a 48-well plate.
Transfer the lens cortex to the plate and incubate for 15 minutes at room temperature with gentle shaking at 300 RPM. After blocking, add appropriate primary antibodies to the 48-well plate and transfer samples to the primary antibody solution. Incubate the samples overnight at 4 degrees Celsius with gentle shaking or mutation.
The next day, wash the samples with PTX, and similarly, incubate them with secondary antibodies. After washing the samples, add one drop or 50 microliters of mounting media onto a plus charged microscope slide. Place the tissue in the mounting media.
And use tweezers to gently separate the fiber cells from each other. Then, gently place a cover slip on top of the separated cells and mounting media. After removing excess media, use nail polish to seal the edges of the cover slip on the slide before confocal microscopy.
After the lens dissection and lens capsule removal, transfer the fiber cell mass to wet gloved fingertips and gently roll in all directions to separate the lens nucleus. Then, transfer the lens nucleus to freshly made 1%paraformaldehyde solution in a 48-well plate and incubate overnight at 4 degrees Celsius with gentle shaking or mutation. The next day, transfer the sample to a 60 millimeter dish with 1%paraformaldehyde and use a sharp scalpel to split the lens nucleus along the anterior posterior axis in half, then into quarters.
Then, postfix, block, stain, and mount the sample as previously demonstrated In these preparations, bundles of lens fiber cells with unique morphologies are found from different lens regions. The staining of the F-actin network shows enrichment at the cell membrane in differentiating and mature fibers while F-actin signals are present in the cytoplasm of nuclear fibers. Scanning electron microscopy and confocal images of differentiating fiber cells show ball and socket interdigitations with small interlocking protrusions, whereas mature fibers have paddles decorated by small protrusions.
Scanning electron microscopy and confocal microscope images of nuclear lens fiber cells reveal infrequent and larger interlocking protrusions on the short sides of the cells where the cell membrane is rough and has tongue and groove interdigitations in these cells from the center of the lens.
