Neuroscience
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Preparation of Newborn Rat Brain Tissue for Ultrastructural Morphometric Analysis of Synaptic Vesicle Distribution at Nerve Terminals
Chapters
Summary June 7th, 2019
We describe procedures for processing newborn rat brain tissue to obtain high-resolution electron micrographs for morphometric analysis of synaptic vesicle distribution at nerve terminals. The micrographs obtained with these methods can also be used to study the morphology of a number of other cellular components and their dimensional structural relationships.
Transcript
Among several existing protocols for tissue preparation for transmission electron microscopy, we selected methods that yield optimal structural preservation and high resolution micrographs to perform morphometric analysis of synaptic vesicles distribution at presynaptic terminals. The high contrast images obtained with these methods allow to quantify parameters such as distance of synaptic vesicles from the presynaptic membrane number of vesicles docked at the presynaptic membrane and inter vesicle distances. All of these parameters are indicative of synaptic function.
Since alterations in the spatial organization of synaptic vesicles may be associated with the disruption of synaptic function, these methods have been used to examine the effects of general anesthetics on synaptic development. These methods could also be used to examine the effects of any drug or treatment on the detailed morphology of synapses to provide insight into their function. Begin by preparing osmium tetroxide for post fixation of rat brain sections previously fixed with paraformaldehyde and glutaraldehyde via vascular perfusion.
Sections become rigid after treatment with osmium tetroxide. Therefore tissue handling requires attention from then on. Also, before adding osmium, make sure that your sections are flattened as any fold will likely result in tissue fracture.
Open one five milliliter glass ampule of 4%osmium tetroxide and water and drop it inside a brown glass bottle. Add five milliliters of 0.2 molar phosphate buffer and then add an additional 10 milliliters of 0.1 molar phosphate buffer to achieve a 1%final solution. Use a 20 milliliter syringe fitted with a long needle to draw up the 1%osmium tetroxide and then add the osmium tetroxide to a brown glass jar covered with aluminum foil.
After making sure that the specimen has been unfolded and flattened, add 1%osmium tetroxide and 0.1 molar PB to the specimen vial. Incubate the specimen in osmium tetroxide for one hour at room temperature. Extract the osmium tetroxide with a micropipet after incubation is finished and rinse the sections as described in the protocol.
To prepare uranyl acetate, place a 200 milliliter volumetric glass flask containing 100 milliliters of 70%ethanol on a stirring plate. Add four grams of uranyl acetate to the flask, wrap an aluminum foil to prevent precipitation by light and stir continuously. Dehydrate the sections in 50%ethanol.
Stain with the prepared the 4%uranyl acetate for one hour and follow with serial dehydration and rinsing as described in the manuscript. Remove the syringe plunger from a 60 milliliter gavage syringe and cap it with a safety needle. Position the syringe with the open side up and add each ingredient of the resin mix one by one.
Finish by adding 525 microliters of DMP30 with a pipet. Moving the plunger of the pipet very slowly as DMP is very viscous. Put the plunger back in the syringe.
Invert the syringe a few times to mix the contents. The color will change from yellow to amber. Evacuate the air inside the syringe.
Then place it on a rocker with continuous shaking for at least 30 minutes. Add one volume of epoxy resin and one volume of acetone to a scintillation vial and shake to mix. Apply this one to one resin to acetone mixture on the tissue section after the last acetone rinse keeping in covered to avoid acetone evaporation.
Remove the one to one mixture of resin and acetone after two to four hours and replace it with full resin and incubate for four hours. Put all resin waste in a collection container under the hood to polymerize and to be disposed of later. Cut two rectangular pieces of clear polychlorotrifluoroethylene film and wipe them with 70%ethanol.
Trim them so that there is at least 1.5 centimeter of tissue-free plastic on every side of the section, making sure that they have the same width but the height of the sheet on top is approximately 2/3 of the bottom sheet. Slowly tilt the vial and use a fine camel paintbrush and a flexible micro-spatula to very gently lift the specimen from the bottom. Then carefully move the section along the vial walls and transfer onto the bottom PCTFE sheet.
Remove excess resin with a micro-spatula and gently cover the specimen with the top sheet. Gently push out any trapped air bubbles without applying direct pressure onto the tissue section and wipe out the excess resin. Use a solvent resistant pen to label the sheets and place in an oven at 60 degrees celsius for two to three days to polymerize.
Gently pull apart the sandwiched PCTFE sheets to open them. Use a solvent resistant pen to mark the side of the sheet that contains the adhering section marking it near the tissue. Use a disc punch to obtain a circle sample of the section making sure that the pen mark is part of the punched out tissue.
The pen mark will gleam when light is shined on the specimen. Place the cap of an embedding capsule side up on a capsule holder. Place a drop of resin on the cap and insert the punched disk inside the cap with the tissue section facing up.
Insert a capsule into the cap using a gentle corkscrew movement. Use fine tweezers to roll and lower a printed two centimeter long piece of paper label into the capsule, making sure it fits to the curvature of the side walls of the capsule. Pour the embedding resin inside the capsule, fill up to edge of the capsule and use a pointed wooden stick to push the tissue specimen down to the bottom.
Place the capsule in the oven at 60 degree celsius for two to three days for polymerization. And then continue with sectioning and staining as described in the protocol. Electron micrograph of satisfactory preservation of neuronal cell structure shows layered cell membranes without breaks.
The cytoplasm is finally granular and without empty spaces. Mitochondria are neither swollen nor shrunken with a conserved outer double membrane and intact internal cristae. Optimal preservation of neuronal ultrastructure and fine morphological detail is essential in order to visualize synaptic vesicles and measure their distance from the presynaptic membrane.
Synaptic vesicles should be distinct and lined by an unbroken single membrane. In order to facilitate morphometric analysis of synaptic vesicle distribution, presynaptic and postsynaptic membranes should be parallel in their continuity preserved. Electron micrograph of defective preservation of tissue structure shows the distortion and breakage of neuronal cell membranes and the presence of markedly enlarged extracellular spaces.
Mitochondria appear distended and have swollen cristae. In another example of defective tissue structure preservation, there were large white empty spaces within the cytoplasm in place of finely granular cytoplasmic substance with enlarged extracellular spaces. An artificial membrane as forall likely resulted from mobilization of lipids during fixation with glutaraldehyde.
When working young brain tissue use 1 molar phosphate buffer as a solvent for all your solution so that you keep tonicity as close as possible to that of newborn rat brain and minimize artefactual tissue shrinkage and or swelling. The micrographs obtained with these methods can be used to study the detailed morphology and dimensional structural relations of a number of cellular components other than synaptic vesicles. For example, mitochondria, golgi apparatus and nuclear chromatin.
Several steps in this protocol require chemicals that can be toxic. Always work under a fume hood and wear personal protective equipment when handling aldehydes, osmium, uranium and lead compounds.
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