Histological Staining Techniques
Holmes’ method for axons
Use 10-20 micron paraffin wax sections.
- Dewax sections, rinse in alcohol, rinse in distilled water.
- Place in 20% silver nitrate in closed coplin jar - 2
hrs in the dark.
- In the meantime prepare the impregnating solution.
- Remove slides from the 20% silver nitrate solution and
wash in 3 changes of distilled water over 10 minutes.
- Place the slides overnight in the impregnating solution
at 37ºC in a sealed coplin jar.
- Shake off superfluous fluid and place the slides directly
in the reducing solution - 2 minutes minimum. (10-15 mins recommended.)
- Wash in running tap water for 3 minutes then rinse in
- Check slides microscopically for good contrast,
return to reducing solution if insufficient.
- Tone in 0.2% gold chloride for 3 minutes.
- Rinse briefly in distilled water.
- Place in 2% oxalic acid for 2-10 minutes,
examine at intervals until the axons are blue-black colour. (4-5 mins
recommended, too long may result in lower contrast.)
- Rinse in distilled water and fix with 5% sodium
thiosulphate - 5 minutes.
- Wash in tap water - 10 minutes.
- Dehydrate, clear and mount.
- 1.24% aqueous boric acid - 55ml
- 1.9% aqueous borax (Na2B4O7.10H2O)
solution - 45ml
- Distilled water - 394ml
- 1% silver nitrate - 1ml
- 10% aqueous pyridine - 5ml
- Mix thoroughly
- Hydroquinone 1g
- *Sodium sulphite 10g
- Distilled water 100 ml
Axons - black
Background - greyish (turning purple with time)
Muscle striations - black
Coplin jars should be thoroughly cleaned before use.
The 20% silver nitrate can be re-used but the impregnating must only be used
This method is applicable for both CNS and peripheral nerve material, and also
demonstrates cross-striations of muscle fibres.
*When making up the reducing solution it is best to add the sodium sulphite
to the water while it is being rapidly stirred. If not it tends to form a hard
mass that is difficult to dissolve.
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