This protocol is the first to describe how to process and evaluate mouse size for RP pathologies using light transmission electron and confocal microscopy. The main advantage of this protocol is the inclusion of quantitative assessments of the RP pathologies, using these three imaging techniques for statistical comparisons. The techniques described in this protocol may also help to expand our knowledge of molecular pathways important for RPE health.
Individuals who have not previously dissected mouse size may struggle with this protocol. We suggest these individuals practice with some mice to master the techniques described in this protocol. To begin, prepare a gravity feed perfusion system on an absorbent underpad for cardiac perfusion in a fume hood.
Pour 40 milliliters of freshly prepared fixative buffer into the syringe barrel of the perfusion system. Turn the valve parallel to the tubing line to allow the buffer to flow through the tubing line. Flush the line until all air bubbles are removed from the line.
Then turn the valve perpendicular to the tubing line to stop the buffer from flowing into the tubing line. To perform cardiac perfusion, transfer the euthanized mouse to a shallow tray near the perfusion system. With the abdomen facing up.
Spray the abdomen with 70%ethanol, then create a five centimeter inferior cut using curved scissors and forceps through the skin and abdominal wall on the left side of the mouse below the rib cage. Proceed to make a three centimeter medial cut through the skin and abdominal wall at the top of the inferior cut. Make another five centimeter inferior cut at the end of the medial cut through the skin and abdominal wall on the furthest right side of the mouse below the rib cage.
Make another three centimeter medial incision to remove the abdominal skin flap with curved forceps. Next, cut through the diaphragm and sternum to expose the heart. Insert the gauge needle into the left ventricle of the heart and turn the valve.
Cut the right atrium with curved scissors to allow blood and fixative to exit the heart. Allow 10 milliliters of fixative buffer to perfuse the mouse for one to two minutes, or until the liver becomes pale in color and no blood flows out of the right atrium. Once the perfusion is complete, turn the valve perpendicular to the tubing line to stop the buffer flow.
To nucleate the eyes, remove the mouse from the shallow tray and place it on the absorbent underpad in a fume hood. Then orient the head of the mouse with the left eye facing the experimenter and the right eye out of view. Annotate the superior side of the eye with a tissue marking dye.
Gently push down around the eye socket with the thumb and index finger for the protrusion of the eye from the eye socket. Then using curved scissors, hold the eye with the blade at a 30 degree angle from the eye socket and cut around the eye. Remove the eyeball from the head with curved forceps and place it on an absorbent underpad.
Nick the cornea with a number 11 scalpel blade and place the eye with curved forceps into a two milliliter micro tube containing two milliliters of fixative buffer. After nucleating both eyes, incubate them in a fixative buffer overnight on a shaker in a four degree room at a speed of 75 rpm. The following day, replace the fixative buffer with two milliliters of PBS and incubate for 10 minutes on a shaker at room temperature and 75 rpm.
Next to generate posterior segments place the eye into a Petri dish filled with PBS under a dissecting microscope. Gently lift any fat and muscle away from the eyeball with fine tipped forceps. Carefully trim the fat and muscle with micro dissecting scissors in a parallel direction to the eyeball until the eyeball has a uniform bluish-black color Place fine tipped forceps at the corneal puncture site and cut around the perimeter of the cornea beginning at the puncture site with micro dissecting scissors to remove the cornea and iris from the eyeball.
Then using fine tipped forceps, gently remove the lens from the eyeball to yield the posterior segment. Place the posterior segment into a tube containing two milliliters PBS, and store at four degrees Celsius in a refrigerator until use. Transfer the eye to a Petri dish containing PBS and remove the cornea and iris as demonstrated.
Pull out the lens with fine tipped forceps. Then using two fine tipped forceps, gently separate the neural retina from the posterior segment. Carefully cut the neural retina at the optic nerve head for removal from the posterior eye cup.
Place the eye cup into a two milliliter micro tube containing five microliters of PBS. To fix posterior eye cups, add 500 microliters of methanol to the tissue and incubate on a shaker at 75 rpm, for five minutes. Repeat methanol fixation three times with final incubation for two hours.
After fixation, wash the tissue thrice with PBS, before proceeding to immunofluorescence staining and visualization through confocal microscopy. Four month old TMEM 135 TG mice, showed a significant reduction in retinal pigmented epithelium or RPE thickness at 600 and 900 micrometers away from the optic nerve, relative to age-matched wild type mice. Incidents of RPE pathologies including microvascuolization, macrovacuolization and migration of 325 day old WT and TMEM 135 TG mice were calculated.
The average frequency of RPE microvacuolization was lower in the wild type compared to the TMEM 135 TG mice. No RPE macrovacuolization and migration were detected in wild type compared to the average values observed in TMEM 135 TG mice. In transmission electron microscopy, basal laminar deposits were observed in 24 month old retinas that were absent in two month old retinas.
Analysis of cumulative frequencies of the basal laminar deposits heights, indicated a shift to the right of the line for the 24 month old, compared to the two month old, demonstrating an increase in deposits in 24 month old mice. This observation was supported by larger average heights in 24 month old retinas. RPE in four month old TMEM 135 TG mice was dysmorphic.
The RPE in TMEM 135 TG retinas are larger and less dense than age-matched wild type retinas. Also, more multinucleated RPE cells in the TG retinas were observed, compared to controls. Following this protocol researchers could evaluate specific pathways using molecular biology techniques in mice, which could contribute to our understanding of how these RP pathologies develop in mice.
This protocol will help to standardize RP phenotyping in mice, which will translate findings from mouse studies to other AMD models and aid in our understanding of AMD pathobiology.
