The Warthin-Starry stain

INTRODUCTION

The Warthin-Starry is a silver stain primarily used to detect spirochaetes and some small bacilli such as:

• Helicobacter pylori (linked to gastric ulcers, gastric cancer)
• Bartonella henselae, afipia felis (cat-scratch disease)
• Treponema pallidum (syphilis)
• Legionella (legionellosis)
• Leptospira (Weil’s disease)
• Borrelia (Lyme disease)

While these can also be demonstrated via immunohistochemistry, which have a high specificity, however, they require expensive antibodies. Similarly, bacteria such as Borrelia and H. pylori can be stained with dyes, notably the Giemsa stain.

The bacteria are argyrophilic, that is, they have the ability to bind silver ions from a solution, but they cannot reduce that silver to a visible metallic form themselves. Therefore, a reducer must be used when using silver to stain them.

The Warthin-Starry stain was developed by Aldred Scott Warthin and Allen Chronister Starry for the demonstration of Spirochaeta pallida, now known as Treponema pallidum, the causative agent of syphilis. They published their paper “A more rapid and improved method of demonstrating spirochaetes in tissues (Warthin and Starry’s cover-glass method)” in the American Journal of Syphilis in 1920.

Their method was an improvement from the Levaditi method, which took place over several days and did not provide consistently good results. Methods utilised nowadays are variations of the Warthin-Starry method from 1920, using a reducing solution consisting of hydroquinone, silver nitrate and gelatin, instead of the pyrogallic acid used in the original method. Coincidentally, the use of pyrogallic acid as a developer in photography also fell out in favour of hydroquinone in the 1920s due to its erratic and unreliable behaviour at the time.

Since the publication of the original method, other variations have been published with variations in reagents and incubation times. These methods will be presented later in the Methods section.
As well as these published variations, different companies have developed commercial kits that can be purchased and used for the stain.

PRINCIPLE

The Warthin-Starry stain is a silver stain that can be divided in two stages:
First, the formation of submicroscopic clusters of metallic silver on the bacteria of interest by using a buffered silver nitrate solution.
Second, the further reduction of more silver ions on the bacteria using a developer solution, which makes the bacteria microscopically visible.

The stain is similar to the silver stains used to demonstrate axons in nerve fibres, with an important difference being the pH of the buffered solution. Whilst a pH of 8.0 to 8.5 is optimal for the demonstration of axons, a lower pH is required for the demonstration of spirochaetes without the staining of other tissue components. The higher the pH, the more silver that will be deposited and reduced in the surrounding tissue, thus creating background staining that will obscure the bacteria.

First, the sections are incubated in a heated buffered silver nitrate solution. The pH for this solution should be between 3.5 and 4. This low pH allows for the silver ions to bind more to the bacteria than to the rest of the tissue components, thus creating less background staining once developed. Some of these ions are reduced on the bacteria at this point, forming submicroscopic deposits of metallic silver which are too small to be visible under the microscope at this stage.

In the second stage, the sections are developed in a reducing solution. This solution generally contains hydroquinone, gelatine, and buffered silver nitrate solution. Concentrations of these reagents will vary depending on the protocol.
The hydroquinone reduces the silver ions to metallic silver, while the gelatine sequesters these silver ions thus slowing down their reduction. The silver already reduced on the bacteria catalyses the reduction of more silver ions (that is, it speeds up the reaction). This creates bigger deposits of metallic silver on the bacteria which are now microscopically visible.
Silver ions on the rest of the tissue are reduced at a much slower rate, resulting in a yellow/light brown background colour that does not obscure the dark grey/brown bacteria. If left in the developer solution for too long, the tissue will eventually stain as dark as the bacteria, therefore obscuring its visualisation.

As a result of the stain with adequate development, spirochaetes are clearly stained allowing for the visualisation of individual bacterium which are not obscured by the golden/yellow background staining of the tissue.

METHODS

All the following methods are applied to formalin fixed, paraffin embedded (FFPE) tissue sections.

My preferred method for the Warthin-Starry stain is a variation of the method published by Kerr in 1938. It is as follows:

Reagents:

• 1% citric acid in distilled water.
• Buffer solution: To 125 mL of distilled water, add 1 mL of 1% citric acid. Check that the pH is approximately 4. pH indicator paper can be satisfactorily used for this.
• 2% silver nitrate in buffer.
• 1% silver nitrate in buffer. This can be made by diluting part of the 2% silver nitrate 1:1 in buffer solution.
• 3% hydroquinone in buffer.
• 5% gelatine in buffer.
• For the developing solution, mix the following solutions immediately before use:
• 3 mL of 2% silver nitrate in buffer.
• 12.5 mL of 5% gelatine in buffer.
• 1 mL of 3% hydroquinone in buffer.
  If a larger volume of developer is required, the same 2:12.5:1 proportion of reagents applies.

Procedure:
For staining up to five slides, the incubations in different reagents can be carried out in slide mailers (unused and clean) which can hold up to approximately 25 mL of solution, instead of using 50 mL Coplin jars. If slide mailers are used, the recommended volumes of reagents to make up are as follows:

• 126 mL of citric acid buffer (1 mL of 1% citric acid in 125 mL of distilled water).
• 20 mL of 2% silver nitrate in buffer.
• 20 mL of 1% silver nitrate in buffer (to 10 mL of 2% silver nitrate in buffer, add 10 mL of buffer solution).
• 10 mL of 3% hydroquinone.
• 20 mL of 5% gelatine.
• 33 mL of developer (6 mL of 2% silver nitrate, 25 mL of 5% gelatine in buffer, 2 mL of 3% hydroquinone in buffer).

If Coplin jars are used, adjust volumes as required.

If possible, it is recommended to stain more than one slide of the same tissue, each with an accompanying control, in order to obtain different development times.

The staining procedure is then as follows:

1. Dewax and hydrate the sections through xylene and graded alcohols, bringing them to distilled water as per laboratory’s standard procedure.
2. Place slides in buffer solution for five minutes.
3. While the slides are in the buffer solution, pre-heat the 1% buffered silver nitrate solution for five minutes in a water bath at 65-70°C.
4. Place the slides in the heated 1% buffered silver nitrate solution, outside of the water bath, for ten minutes with or without agitation.
5. While the slides are in 1% buffered silver nitrate, prepare the developer solution.
6. Transfer the slides directly from the 1% buffered silver nitrate into the freshly made developer solution and keep in the water bath with agitation for 1 to 5 minutes, until the tissue sections appear macroscopically yellow/golden. Please note different tissue types will develop at different rates and thickness of the section will affect staining intensity. When checked microscopically at this step, the control must show dark brown spirochaetes and a yellow/golden background which does not obscure the spirochaetes.
   If more than one slide of the same tissue is used, different development times can be achieved (e.g. 1 minute, 2.5 minutes, 5 minutes).
7. Rinse slides with buffer solution.
8. Wash slides in tap water.
9. Dehydrate, clear, and mount slides as per laboratory’s standard procedure.

This method provides clear demonstration of the spirochaetes in dark brown with a golden/lighter brown background staining which does not obscure the bacteria, as seen in the pictures above.

Other methods include:

From Bancroft’s
Reagents:

• Acetate buffer pH 3.6 (sodium acetate 4.1g, acetic acid 6.25mL, distilled water 500mL)
• 1% silver nitrate in pH 3.6 acetate buffer
• Developer. Dissolve 3g of hydroquinone in 10mL pH 3.6 buffer, and mix 1mL of this solution and 15mL of warmed 5% Scotch glue or gelatin; keep at 40°C
• Take 3mL of 2% silver nitrate in pH 3.6 buffer solution and keep at 55°C. Mix these two solutions immediately before use.

Procedure:

1. Deparaffinize and rehydrate through graded alcohols to distilled water.
2. Celloidinize in 0.5% celloidin, drain, and harden in distilled water, 1 minute.
3. Impregnate in preheated 55-60°C 1% silver solution, 90-105 minutes.
4. Prepare and preheat developer in a water bath.
5. Treat with developer for 3.5 minutes at 55°C. Sections should be golden brown at this point.
6. Remove from developer and rinse in tap water for several minutes at 55-60°C, then in buffer at room temperature.
7. Tone in 0.2% gold chloride.
8. Dehydrate, clear, and mount.

Notes: It is wise to take a few slides through at various incubation times to ensure optimum impregnation.

From Kiernan’s Histological and Histochemical Methods

Reagents:All solutions are used hot (55°C). The bottles of stock solutions should be stood in a suitable water bath or oven for an hour or two before using.

• Acetate buffer, pH 3.6. 0.1M acetic acid-sodium acetate buffer. Can be kept for several weeks, however, do not use if it is cloudy or contains any kind of deposit.
• Silver solution 2%. Buffer 50mL, silver nitrate 1.0g.
• 5% gelatin in distilled water. Warm to about 40°C to melt solution before using. The solution must not be turbid.
• Reducer. This is a physical developer. Make up the three solutions while the sections are in the silver solution, and combine them immediately before using.
• Hydroquinone, 30 mg. Dissolve in 20 mL water and heat to 55°C.
• Silver solution 2%, 15mL. Heat to 55°C.
• 5% gelatin, 37.5mL. Heat to 55°C.
• Combine the three hot solutions in a staining jar immediately before carrying out Step 3 of the procedure.

Procedure:

1. Dewax and hydrate paraffin sections.
2. Dilute silver solution 2% with an equal volume of acetate buffer and immerse slides in the diluted solution for 60 minutes at 55°C. Prepare the components of the reducer.
3. Combine the three components of the reducer. Leep at 55°C until the sections are a gold-brown colour (3-5 minutes).
4. Pour off the reducer and wash at 55°C (running hot tap water is satisfactory) for 5 minutes.
5. Rinse in distilled water, dehydrate, clear, and mount in a resinous medium.

From Histotechnology: A Self-Instructional Text

Reagents:

• Citric acid, 1% solution.
• Acidulated water. To triple-distilled water, add enough 1% citric acid to bring the water to pH 4.0.
• Silver nitrate (for developer), 2% solution in acidulated water.
• Silver nitrate (for impregnation), 1% solution in acidulated water. Do not preheat this solution.
• Gelatin, 5% solution in acidulated water.
• Hydroquinone, 0.15% solution in acidulated water.
• Developer solution.
• Silver nitrate, 2% solution, 12mL
• Gelatin, 5% solution, 30mL
• Hydroquinone, 0.15% solution, 16mL 

• Use a prewarmed graduated cylinder and combine reagents in the order given making certain the solutions are mixed well after each addition. Prepare immediately before use.

Procedure:

1. Place the 2% silver nitrate, 5% gelatin, and hydroquinone solutions in separate 50mL plastic centrifuge tubes; heat in a water bath at 54°C for at least 1 hour.
2. Place a 100mL graduated cylinder and a chemically clean Coplin jar in the oven for at least 1 hour (for developer).
3. Deparaffinize and hydrate sections to acidulated water.
4. Place slides in the 1% silver nitrate impregnating solution in a water bath at 43°C for 30 minutes; do not preheat the solution.
5. Just before the slides are due out of the impregnating solution, prepare the developer (place in the warm Coplin jar) and place in the 54°C water bath.
6. Put slides in the developer for 3-4 minutes; check after 2 minutes and continue checking frequently until they are ready.
7. Wash slides quickly and thoroughly in distilled water.
8. Dehydrate sections in 95% and absolute alcohols and clear in xylene (2 changes each).
9. Mount sections with synthetic resin. 

Technical notes:

• If the sections have been overdeveloped, they may be treated with iodine and sodium thiosulfate for colour removal and then restained.
• These are three textbook methods, however, there are other published variations of the original method as well as microwave methods and commercial kits available. The Sources section lists published papers with different methods that have not been listed above and that are worth reading.

TROUBLESHOOTING

As with any silver stain, it is important to use clean glassware and slides to prevent artifacts from the contamination of reagents.

The main issues that can occur in the Warthin-Starry stain, as listed on “Silver staining for spirochaetes in tissue: rationale, difficulties, and troubleshooting” (Kiernan 2002, Laboratory Medicine), are as follows:

ISSUE	CAUSE	SOLUTION
Weak staining	-Insufficient silver nuclei formed in the first stage. -Inadequate development.	-Longer incubation in hot buffered silver nitrate. -Check development time at different points up to 12 minutes.
Staining of unexpected structures: components of the tissue	-Use of fixatives containing mercuric chloride or potassium dichromate. -Excessive exposure to either the buffered silver nitrate or the developer solution.  -Excessive temperature of the reagent -Presence of argentaffin components such as melanin, cytoplasmic granules in certain neuroendocrine cells, granules within some cell nuclei.	-Avoid use of fixatives containing those chemicals. -Reduce time in buffered silver nitrate and/or developer solution. -Ensure the temperature of the reagents is the right temperature for the protocol followed. Use thermometer in the solution. -The presence of argentaffin components in the tissue cannot be avoided.
Granular deposits: formalin pigment. Appears as black granules mostly in and near blood vessels.	Storage of specimen in acidic solution of formaldehyde which causes granules to form as a result of the degradation of haemoglobin.	Use neutral buffered formalin.
Silver precipitates on the tissue section or on the slide.	-Presence of dirt on the glassware (bottles, slides, staining jars, etc) such as metallic silver remaining from previous uses. -Presence of chloride ions (e.g. from tap water). -Impurities in the gelatin.	-Thoroughly wash and rinse all glassware, including slides. Glassware that contained silver solutions can be cleaned with nitric acid and rinsed with distilled water. -Use distilled water for the last rinse of glassware. -Gelatin from chemical suppliers is unlikely to cause issues. Avoid contamination of stock bottles of chemicals by using different spatulas.

Sources and other articles of interest

Reminder: sci-hub (currently at https://sci-hub.tw/) can be used to access texts behind a paywall. One can also use the hashtag #canihaspdf on Twitter to ask people to share the full text with you if they have it.

• Warthin, AS; Chronister, AC (1920). "A more rapid and improved method of demonstrating spirochetes in tissues (Warthin and Starry's cover-glass method)". American Journal of Syphilis. 4: 97–103.
• https://en.wikipedia.org/wiki/Pyrogallo
• Silver Staining for Spirochetes in Tissues: Rationale, Difficulties, and Troubleshooting (Kiernan 2002, Laboratory Medicine)
• Silver staining of spirochaetes in single tissue sections (Faine 1964)
• A buffer modification of the Warthin-Starry silver method for spirochaetes in single paraffin sections (Faulkner 2945, Stain Technology)
• Improved Warthin-Starry method of staining spirochetes in tissue sections (Kerr 1938, American Journal of Clinical Pathology)
• A Warthin-Starry Method for Spirochetes and Bacteria Using a Microwave Oven (Churukian 1988, Journal of Histotechnology)
• Books:
• Bancroft’s Theory and Practice of Histological Techniques
• Histotechnology: A Self-Instructional Text
• Histological and Histochemical Methods: Theory and Practice