The overall goal of this methodology is to properly Golgi stain brain tissue in order to examine and quantify dendritic morphology and complexity. This method can help answer key questions in neuroscience. Since Golgi staining only stains a small subset of cells, this technique could be used to trace the structure of neurons.
The main advantage of this technique is that brain sections can be easily stained without risk to them drying out or falling off during any of the washing steps. Demonstrating the procedure will be Fred Kiffer, Tyler Alexander and Julie Anderson, graduate students in Dr.Allen's laboratory. Begin by immersing freshly harvested, unfixed rodent half brains in five milliliters or mercuric chloride-based solution.
As mercuric chloride solution is light sensitive, cover the tube with foil. Store the samples in the dark at room temperature. The following day, slowly decant the solution into a temporary container before properly disposing of it in a biohazard waste container.
Add five milliliters of fresh mercuric chloride solution and ensure that the brains are immersed. Return the samples to the dark at room temperature for 13 more days. After 14 days, prepare post-impregnation buffer by adding 30 grams of powered post-impregnation buffer to 90 milliliters of distilled or deionized water, DH2O.
Fill each well of the six-well plate with 6.5 milliliters of the post-impregnation buffer, one well per brain. Then, remove the mercuric chloride solution from the brain tissue as before. Rinse the tissue with DH2O, then transfer the tissue to the six-well plates with post-impregnation buffer.
Cover the plates with foil and store at room temperature in the dark. The next day, aspirate and replace the post-impregnation buffer. Re-cover the plate and store in a four degree Celsius fridge.
Prepare two or three 12-well plates per brain by first labeling the plates on their lids with numbers in ascending order from left to right and up to down. Add the identification number of the brain sectioned. Pipette two milliliters of 1X PBS into each well of each plate, then set up the stage and turn on the vibratome.
Then, put tape on the tissue block of the specimen holder and add a little cyanoacrylate glue on top. Remove a brain from the six-well plate and place it on a plastic or glass dish. Use a stainless steel double edged blade to cut the cerebellum caudally, making sure that the portion of the cerebellum remaining on the brain is flat.
Then, place the flat surface of the remaining cerebellum on the glue. After the glue is dry, place the specimen holder with the tissue into the specimen bath. Place the specimen bath on the vibratome and cover the tissue sample in PBS.
Then, attach the blade to the blade holder of the vibratome and slowly lower the blade into the specimen bath until it is fully submerged and slightly below the top of the tissue. Next, set the vibratome speed to seven, the amplitude to six and the cutting angle to 12 degrees. Start the vibration and cut 200-micron sections.
Use a large paint brush to move the tissue sections from the specimen bath into each designated well of the 12-well plates. Continue to cut the tissue until the desired number of tissue sections is reached. After sectioning, use a transfer pipette to aspirate the PBS from the wells of the 12-well plate, then add two milliliters of PBS-T and wash for 30 minutes with agitation.
Close to the end of the wash period, dilute three parts ammonium hydroxide solution with five parts DH2O. Then, after removing the PBS-T, add the diluted ammonium hydroxide solution. Protect from light using foil and stain the sections for 19 to 21 minutes.
Then, after safely removing the ammonium hydroxide solution, add post-staining buffer. Protect with foil and stain for another 19 to 21 minutes. Finally, rinse the sections in PBS-T three times for five to 10 minutes each time.
Mount the sections on 1%gelatin coated slides with the large paint brush. Allow them to dry for 20 to 30 minutes at room temperature, then place them in a Coplin Jar and leave overnight. Remove the slides from the Coplin Jar and place them in a plastic rack in the fume hood.
Place in a staining dish containing 100%ethanol for five minutes. Dehydrate the sections through two further five-minute ethanol washes. Next, place the slides in a glass rack and use a metal handle to lower the rack into a glass staining dish containing 99%xylene.
Wash the slides in two changes of xylene for five minutes each. After the time for the second wash has elapsed, remove the slides from the xylene one at a time and quickly cover the tissue with approximately 0.25 milliliters of mounting media. Next, take a slide cover and lay it over the media.
Push any trapped air bubbles underneath the slide covers to the edge with a blunt object, such as the end of paint brush. Place the slides in an area out of direct sunlight and allow them to dry for two to three days. After drying, proceed to image acquisition using a microscope and analysis with the appropriate software.
This bright field image shows a Golgi-stained neuron that meets the criteria for imaging and further analysis. All three spine types can easily be seen. Staining is consistent across the entire length of the dendrite and the dendrite is isolated from other neurons.
This figure shows that Golgi stained neurons within the Dentate Gyrus showed decreased length at the fourth and sixth order following 5-fluorouracil treatment compared to the saline-treated group. Golgi staining can be used to detect significant differences in spine density. In the CA1 apical pyramidal dendrites, 5-fluorouracil administration to live mice significantly decreased spine density.
In the basal dendrites, there were no significant changes in the overall density of spines. Sholl Analysis of CA1 apical pyramidal dendrites revealed that 5-fluorouracil treatment significantly decreased dendritic arborization at distances between 40 to 160 microns away from the soma. In the basal dendrites, the 5-fluorouracil treatment decreased the dendritic arborization at 30 to 110 microns away from the soma.
Once mastered, this technique, accounting for both the Golgi staining process and mounting, cleaning and covering can be done in three hours per brain if performed properly. After its development, this technique paved the way for researchers in the field of neurobiology to explore neuroanatomical connections in a murine model. While attempting this procedure, it's important to remember to keep the sections in ammonium hydroxide solution for only 19 to 21 minutes as to not overstain the neurons.
After watching this video, you should have a good understanding of how to properly stain, section and rinse brain tissue. Don't forget that working with the mercuric chloride-based solution, ammonium hydroxide solution and xylene can be extremely hazardous, so take precautions and wear lab gloves and work in a fume hood when performing this procedure.
