Place 5 mL of the 10% silver nitrate in a 100 mL flask.
Using a pasteur pipette, add a drop of strong ammonium hydroxide then swirl the solution for a few seconds.
A precipitate will form at first.
Continue adding ammonium hydroxide drop by drop and swirling until the precipitate is just redissolved.
Add 5 mL of 3% sodium hydroxide and mix well.
A precipitate will again form.
Add drops of ammonium hydroxide until the precipitate is just redissolved, leaving a faint opalescence.
Dilute to 50 mL with distilled water.
5µ paraffin sections of neutral buffered formalin fixed tissue are suitable.
Other fixatives are likely to be satisfactory. A section adhesive is recommended.
Dehydrate with ethanol, clear with xylene and mount with a resinous medium.
Reticulin fibres – black
Nuclei – red
Background – grey
Gordon & Sweets also suggested a variant that used one of Foot's ammoniacal
silver solutions. Apart from that, the method is the same as given above.
Foot's ammoniacal silver
To 10 mL of 1% silver nitrate add 0.1 mL of 40% potassium hydroxide.
Add strong ammonium hydroxide by drops until the precipitate just redissolves.
Make up the volume to 100mL with distilled water.
Ensure that both the ammonium hydroxide and sodium hydroxide are fresh and full strength.
Keep both well stoppered when not in use. For the ammonium hydroxide, pour sufficient for use from the
stock bottle into a beaker, then immediately restopper the stock bottle. Do not return excess ammonium
hydroxide to the stock bottle.
After making the ammoniacal silver solution, smell the solution to ensure it has only
a faint smell of ammonia. If the smell of ammonia is strong it indicates that too much ammonium hydroxide
has been added. If so, it is preferable to make the solution again.
Improperly made ammoniacal silver solutions can affect the quality of the impregnation.
Most references to the Gordon & Sweets' reticulin stain specify that the
ammoniacal silver solution should be made with 10.2% aqueous silver nitrate and 3.1% aqueous sodium hydroxide.
No explanation is given. The 10% and 3% solutions respectively, as given by Gray, work satisfactorily.
Iron alum is ferric ammonium sulphate. For routine formalin fixed tissue 15 minutes
in the iron alum is usually sufficient. If necessary the time may be extended up to 2 hours.
10% formalin is made by diluting strong formalin 1:10 with tap water
(10 mL strong formalin, 90 mL tap water).
Toning is a variable step. Untoned sections give dark brown reticulin fibres on a paler
brown background. Many microscopists prefer to tone for about 15 seconds to produce brown-black reticulin
fibres on a pale grey-brown background. Others tone longer (a few minutes) to produce black reticulin
fibres on a grey background. Longer toning produces purple tones. Tone according to the personal preference
of the microscopist reviewing the slides.
Reference Drury, R A, and Wallington, E A, (1967). Carleton's histological technique., Ed. 5.
Oxford University Press, London, England.
Culling, C F A, Allison, R T, Barr, W T, (1985). Cellular pathology technique., Ed. 4.
Butterworths, London, England.
Gray, Peter. (1954) The Microtomist's Formulary and Guide. Originally published by:– The Blakiston Co. Republished by:– Robert E. Krieger Publishing Co.