The overall goal of this experiment is to develop a new approach to analyze membrane proteins at retinal ribbon synapses quantitatively. This method can help answer key questions in the retinal neurobiology field such as the precise location of proteins at synapses within the circuit. The main advantage of technique is that it can estimate the number, density, and rapidity of multiple proteins at retinal ribbon synapses.
Demonstrating the procedure of freeze-substitution and the ultra-thin sectioning we have docs Ron Petralia and Ya-Xian Wang from the NIDCD Advanced Imaging Core. To begin this procedure dissect an eyeball under the microscope. To do so, first remove the cornea.
Then remove the lens and vitreous from the inter-retinal surface with forceps. Afterward, peel the sclera until the retina is isolated from the eyecup. Cut the retina immediately into 100 200 micrometer thick strips with a razor.
Subsequently, mix the retina strips in 4%paraformaldehyde in 0.1 molar PB.Then, in 4%paraformaldehyde plus 0.01%glutaraldehyde at room temperature. After several washes in PB, with 0.15 millimolar calcium chloride, cryoprotect the retinal strips with glycerol in 0.1 molar PB, prior to freeze-substitution. Prior to placing the frozen tissue into the flat embedding holders, cut a thin circle from a clear plastic sheet and place it on the bottom of each holder to allow easier removal of the polymerized specimen box when finished.
Now, set the automatic sequence in the AFS. Fill the AFS with liquid nitrogen and place the flat embedding holders in the AFS. Fill them with 1.5%uranyl acetate in methanol and pre-cool them.
To plunge-freeze the retinal strips in liquid propane at 184 degrees Celsius, use a fine brush to place the samples on small pieces of double-stick tape attached to the metal stubs on the end of the plunging rods. After that, wick off extra buffer using the brush. Plunge the rods into the liquid propane for about five seconds.
Then, transfer them to the AFS using a small transport chamber that is filled with liquid nitrogen. After placing the frozen sample and instruments, forceps and scalpel, into the AFS, cool the instruments in the chamber for several minutes before touching the tissue. Or, cool them by placing the tips of the instruments for a few seconds into the small transport chamber filled with liquid nitrogen.
After that, remove the sample and tape from the stub using a fine scalpel. Next, place two pieces of frozen sections in each of the seven wells in the flat embedding aluminium holder with 1.5%uranyl acetate in methanol. Keep them at 90 degrees Celsius for at least 32 hours in the AFS.
Then, increase the temperature stepwise to 45 degrees Celsius in the automatic sequence. After that, wash the samples three times in pre-cooled methanol. Next, infiltrate the samples progressively with low-temperature embedding resins as an embedding medium.
On the next day, change the resin again and adjust the level of resin to reach the top edge of the wells. Set up the UV light on the AFS and polymerize the samples with UV light in the AFS until the end of the automatic sequence. Then, remove the samples from the AFS.
Typically, sample blocks still show some pinkness and color. Place the samples close to the fluorescent light in the chemical fume hood at room temperature overnight, or until they appear completely clear. In this step, cut 70 nonometer thick ultra-thin sections with an ultramicrotome, and collect them on the formvar carbon coated nickel grids.
Wash the grids in distilled water once, followed by three washes of TBS. Then, incubate the grids in 20 microliters of 5%BSA in TBS for 30 minutes, and then in 20 microliters of a mixture containing goat anti-CTB and an antibody to one NMDAR sub-unit overnight at room temperature. After that, wash the grids three times with 20 microliters of TBS each time at ph 7.6.
Subsequntly, wash the grids once with TBS at ph 8.2 for five minutes. Incubate the grids for two hours with 20 microliters of a mixture containing donkey anti-rabbit IGG coupled to 10 nanometer gold particles, and donkey anti-goat IGG coupled to 18 nanometer gold particles. After two hours, wash the grids in 20 microliters of TBS, at ph 7.6 three times, followed by a wash in the ultra-pure water and then air-dry the grids.
Subsequently, counterstain the grids with 5%uranyl acetate in distilled water for eight minutes, followed by 0.3%lead citrate in distilled water for five minutes in the dark. For triple-labeling experiments, incubate the grids overnight at room temperature with 20 microliters of a mixture of anti-goat CTB, anti-mouse PSD-95, and anti-rabbit UN2A. Then, incubate the grids for two hours with 20 microliters of a mixture of IGGs coupled to 18, 10, and five nanometer gold particles.
In this section, first set the scale bar. Then measure the length of the PSD of individual RGC dendrites with ImageJ software. Then, count the gold particles within 10 nanometers of the membrane based on an average thickness of seven to nine nanometers for the plasma membrane.
Calculate the particle density as the number of gold particles per linear micrometer. Then, measure the distance between the center of each gold particle and the middle of the PSD for synaptic location, or the edge of the PSD, for extra-synaptic location. This image shows the double immunogold labeling of GluA2/3 and CTB.
This is the pre-synaptic ribbon and the small gold particles are clustered in the PSD of the RGC processes. This image shows the double immunogold labeling of GluN2B and CTB. The small gold particles are on the extra-synaptic plasma membrane.
And this image shows the double immunogold labeling of GluN2A and CTB. Similar to GluA2/3, small particles are clustered within the PSD. The triple immunogold labeling of GluN2A, PSD-95, and CTB is shown here.
GluN2A gold particles and PSD-95 gold particles are co-localized within the the PSD on individual CTB-positive RGC dendrites. While attempting this procedure, it is important to remember to fix the tissue very gently, because some protein antigenicity, such as that of NMDA receptors decreases markedly with strong prolonged fixation. After watching this video, you should have a good understanding of how to detect and analyze membrane proteins with great precision at retinal ribbon synapses.
