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Golgi-Cox Staining Protocol for Neurons and Processes
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Golgi-Cox Staining Protocol for Neurons and Processes

Contributed by Tracey Wheeler

George Masson University

(Tracey’s way…there are others as well, so don’t be nervous if you come across them!)

Step 1: Prepare the Golgi-Cox solution

            Solution A: 5% Potassium Dichromate in distilled H2O

                        200ml distilled H2O + 10 grams Potassium Dichromate

                        (Mix in a glass beaker using a glass rod – best to do under fume hood)

            Solution B: 5% Mercuric Chloride (sublimate) in distilled H2O

                        200ml distilled H2O + 10 grams Mercuric Chloride

                        (Mix in a glass beaker using a glass rod)

                        (Mix solution on top of hotplate (on 5), stirring until dissolved)

                        (Must be done under fume hood)

            Solution C: 5% Solution of Potassium Chromate in distilled H2O

                        160ml distilled H2O + 8 grams Potassium Chromate

                        (Mix in a glass beaker using a glass rod – best to do under fume hood)

            Mix Solution A and Solution B into a 500ml glass beaker.

            Mix Solution C and 400ml of distilled H2O into a 1,000ml + glass beaker.

            Slowly pour the AB Solution into the C Solution while stirring continuously

            with a glass rod.

            Store in a glass stoppered bottle for 5 days in the dark.

            Note:  You can easily manipulate the quantity of solution.  Just make sure to

            follow these ratio’s

                        5 Volume parts of 5% Potassium Dichromate solution

                        5 Volume parts of 5% Mercuric Chloride solution

                        4 Volume parts of 5% Potassium Chromate solution

                        10 Volume parts of H20 (to add to PC solution)

Step 2:  Transfer Golgi-Cox solution into small glass bottles.

            Use a plastic syringe to remove the GC solution from the large glass bottle(s). 

            Be sure to avoid the reddish precipitate that formed on the top and bottom of

            the bottle.

            Glass bottles should be filled about ¾ full (to save room for 1 rat brain).

Step 3: Sacrifice Rats using Saline Perfusion Technique.

            Deeply anaesthetize the animal.

            Place on an empty breeder box with wire top. (to allow blood to drain into


            Open chest cavity to expose heart

            Insert 60 ml syringe filled with 9% saline into bottom right chamber of the heart.

            (This would be the animal’s left chamber)

            Using scissors, cut the bottom left chamber of the heart open.

            (This would be the animal’s right chamber)

            Begin to slowly push saline through the animal’s system until the blood leaving the left

            chamber is clear. (This may take 3 syringes of saline.)

            When fluids are clear, decapitate and remove brain.

            Drop whole brain into prepared bottle(s) of Golgi Cox solution. 

            Place in the dark for 14 days, refresh solution after 2 days.

Step 4:  Transfer Brains into Sucrose Solution.

            Mix 300 grams of Sucrose into 1000ml of distilled H2O.

            Place Beaker over hotplate and stir (using stir bar) until dissolved.

            Cool in refrigerator.  (Once cool, ready to use.)

            Empty GC solution from jar and place brain on chem. wipe paper.

            Slightly blot.

            Rinse jar in distilled H2O, and refill with Sucrose Solution ¾ full.

            (In order to save room for brain.)

            Place brain in jar with Sucrose Solution.  Brain will Float.

            Place jar(s) in refrigerator. 

            (Once brains sink, they are ready to be sectioned.)

            (Brains should be sectioned within 2 months of transfer into sucrose.)

Step 5: Section using a Vibratome.

            Prepare razor blade by immersion in xylene for 5 minutes to remove any traces of

oil. (This should be done under the fume hood.)  Wipe blade dry.

            Prepare a 6% sucrose solution. (6 grams sucrose in 100ml distilled H2O.)

            Mix well and make sure it is at room temperature or below before using.

            Fill the vibratome reservoir with the 6% Sucrose solution until the blade is covered.

            Mount brain section (up to ½ a full brain) onto vibratome platform using

superglue.  (Make sure tissue is adhered well before sectioning, 5-7 minutes

or more.)

            Insert platform (with adhered brain section) into reservoir.

            Set the vibratome speed and amplitude around the midpoint for sectioning

            (adjust  as necessary for your specific machine and comfort level.)

            Section at 200 micro meters or desired thickness.  (Sections over 400 may be

difficult to analyze.)

            Using a small paintbrush coax the section onto a gelatinized slide.

            Cover tissue with Parafilm.  Place slide on flat surface covered with bibulous

paper.  Place another sheet of bibulous paper over paraflim.  Place your

palm over the section an press down slightly, being careful not alter

movement.  (Your goal is to press the section into the gelatin on the slide so it

 will adhere to slide during staining.)

            Remove bibulous paper and place in humidity chamber.

            Note:  We used a water sleeved incubator.  In this apparatus it is necessary to

keep the parafilm on the section.  We also placed the slides on plastic trays

which also held capfuls of water to be sure the slides did not dry out.

            Do not keep the slides in storage (humidity chamber or water incubator) for

more than 4 days.)

Step 6: Staining

Prepare fresh solutions (enough to cover all slides):

         Distilled H2O (3 total)

         Ammonium Hydroxide (1 total - keep under fume hood!)

         Kodak Fix (1 total – keep under fume hood, mix in dark)

         50% alcohol (1 total)

         70% alcohol (1 total)

         95% alcohol (1 total)

         100% alcohol (3 total)

         CXA solution (1 total – keep under fume hood!)

         Kodak Fix Solution:

               Prepare all ingredients in beakers 

               Mix in order: (do not mix in light)

               1010 ml distilled H2O (add)

               251 ml Kodak Fix solution A (add)

               28 ml Kodak Fix solution B (add)

               2020 ml distilled H2O

               (you can manipulate amounts, just use one half or 1/3 each)

        CXA Solution:

               1000 ml Chloroform

               1000 ml Xylene

               1000 ml 100% Alcohol

               (you can manipulate amounts, just use one half or 1/3 each)

        Remove parafilm from slides if necessary and place in slide

        rack.  Dip according to process below:

        1.  Distilled H2O for 1 minute

        2.  Ammonium Hydroxide for 30 minutes (IN THE DARK)

            (Using a darkroom light mix the Kodak Fix solution now.)

        3.  Distilled H2O for 1 minute (Best to just keep lights off)

        4.  Kodak Fix solution for 30 minutes (IN THE DARK)

        5. Distilled H2O for 1 minute (once in H2O you can turn on


        6.  50% alcohol 1 minute

        7.  70% alcohol 1 minute

        8.  95% alcohol 1 minute

        9.  100% alcohol 5 minutes

       10. 100% alcohol 5 minutes

       11. 100% alcohol 5 minutes

       12. CXA 15 minutes (Keep under fume hood)

             (Keep slides in CXA under fume hood while cover slipping,

             pull one slide out at a time.)  Note: change gloves often – they

             will disintegrate in CXA.

Step 7:  Coverslip with permount and lie out to dry.

            If possible all cover slipping should be done under fume hood.  Slides should

be allowed to remain under fume hood for 24 hours – lying flat.

            Pull one slide out of CXA at a time.

            Using a glass dropper, place 2 drops of permount on top of tissue.

            (Sections dry quickly, do not remove slide from CXA and allow to sit in air

for more than 20 seconds.)

            Place glass coverslip over section, being careful to avoid trapping air

bubbles.  Note:  Too little permount could allow tissue to dry out, too much

will cause coverslip to slide off – monitor your work for these problems.

            Place slide on absorbent paper (we use white tray liners – usually under rat


            Allow slides to lie flat for 24 hours.

            Slides can now be moved into slide boxes for storage.  KEEP BOXES

            OPEN!!!  Slides need to continue to dry for 6 months before analysis should

be attempted.


1. Gibb R and Kolb B (1998) A method for vibratome sectioning of Golgi-Cox stained whole rat brain. J Neurosci Methods. 79(1):1-4. PubMed Abstract

2. Castano P, Gioia M, Barajon I, Rumio C and Miani A (1995) A comparision between rapid Golgi and Golgi-Cox impregnation methods for 3-D reconstruction of neurons at the confocal scanning laser microscope. Ital J Anat Embryol. 100 Suppl 1:613-22. PubMed Abstract

3. Buller JR and Rossi ML (1993) Immunocytochemistry on paraffin wax Golgi-Cox impregnated central nervous tissue. Funct Neurol. 8(2):135-51.  PubMed Abstract

4. Pugh BC and Rossi ML (1993) A paraffin wax technique of Golgi-Cox impregnated CNS that permits the joint application of other histological and immunocytochemical techniques. J Neural Transm Suppl. 39:97-105. PubMed Abstract

5. Grandin T, Demotte OD, Greenough WT, Curtis SE (1988) Perfusion method for preparing pig brain cortex for Golgi-Cox impregnation. Stain Technol. 63(3):177-81. PubMed Abstract

6. Buell SJ (1982) Golgi-Cox and rapid golgi methods as applied to autopsied human brain tissue: widely disparate results. J Neuropathol Exp Neurol. 41(5):500-7. PubMed Abstract

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