Tag Archives: special stain

Gram-Twort Special Stain For Bacteria – Method and Tips

The Gram-Twort variation of the standard Gram stain is the most common variant used in histopathology laboratories for the demonstration of Gram-positive and Gram-negative bacteria in formalin-fixed sections.

The standard Gram stain was developed and reported in 1884 by Hans Christian Gram (therefore Gram stain should always be spelt with a capital). He developed the technique to distinguish a certain group of bacteria in lung tissue, but noted that Typhus Bacillus was not visualised. The standard Gram stain works on the premise that the peptidoglycan rich cell wall of Gram-positive bacteria is stained by the crystal violet and the Gram-negative bacteria take up the counterstain.

Below is the author’s method of choice.


– Crystal Violet – 0.5% crystal violet in 25% alcohol

– Gram’s iodine – 1g iodine + 2g potassium iodide. Dissolve in a few mls of distilled water. Make up to 300ml with distilled water.

– Acetone

– Stock neutral red-fast green – 90ml of 0.2% neutral red in ethanol + 10 ml of 0.2% fast green in ethanol.

– 2% acetic acid in alcohol.


1. Take sections to water.

2. Stain with filtered crystal violet for 2 minutes.

3. Wash well in tap water.

4. Treat sections with Gram’s iodine for 2 minutes.

5. Wash well in tap water.

6. Rinse sections with acetone until colour stops leeching from them (approx 5 seconds).

7. Counterstain with filtered neutral red-fast green (dilute stock 1 in 4 with distilled water) for 5 minutes.

8. Wash well in tap water.

9. Differentiate in 2% acetic acid in alcohol until red ceases to run from section (approx 5-10 seconds).

10. Rinse in alcohol.

11. Clear and mount in DPX-type mountant.


– Always run a positive control.

– If the Gram’s iodine step is missed it takes longer to differentiate and background staining is increased.

– If having problems with differentiation try completely drying section before differentiating. Blot dry and leave for 10 minutes to ensure complete dryness.

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Grocott Hexamine-Silver Special Stain For Fungus – Method and Tips

The Grocott Hexamine-Silver special stain is the method of choice for a large majority of histopathology laboratories for the demonstration of all fungi. The formalin-fixed sections are exposed to chromic acid which reacts with fungal cell wall polysaccharide components to form chromic acid-aldehydes. These then reduced by a hexamine-silver solution at an alkaline pH. This causes them to be selectively blackened.  It should be noted that this method is not specific for fungi but rarely fails to demonstrate any fungi within the test tissue.

Below is the author’s method of choice.


5% aqueous chromic acid (chromium trioxide)

1% aqueous sodium metabisulphite

Stock hexamine-silver solution = 100ml 3% aqueous hexamine + 5% aqueous silver nitrate.

Working hexamine-silver solution = 2ml 5% aqueous sodium tetraborate (borax) + 25ml distilled water. Mix, then add 25ml stock hexamine-silver solution.

0.1% aqueous gold chloride

5% sodium thiosulphate

0.2% light green in 0.2% acetic acid


1. Take sections to water.

2. Treat sections with chromic acid for 1 hour.

3. Wash thoroughly in tap water.

4. Treat with sodium metabisulphite solution for 1 minute.

5. Wash well in tap water.

6. Wash well in several changes of distilled water.

7. Treat sections with working hexamine solution (preheated in coplin jar at 56 degrees Celsius) at 56 degrees Celsius for 10-20 minutes. Check control sections to see if fungi are a dark brown colour, if not return to solution checking regularly at 3 minute intervals until correct colour achieved.

8. Wash in several changes of distilled water.

9. Treat sections with 0.1% aqueous gold chloride for 3 minutes.

10. Wash well in distilled water.

11. Treat sections with 5% sodium thiosulphate for 5 minutes.

12. Wash well with tap water.

13. Counterstain with 0.2% light green in 0.2% acetic acid for 1 minute.

14. Wash well in tap water.

15. Dehydrate, clear and mount.



– As with all other silver stains wash everything that you are going to use thoroughly with distilled water.

– Store the stock hexamine-silver solution at 4 degrees Celsius away from sunlight. It will keep for 1-2 months. If a white precipitate forms give it a good shake and it should redissolve.

– Do not extend the time in chromic acid too long as this can over oxidize the carbohydrates to carboxylic acid and therefore not take up the silver stain.

– Do not reduce the time in chromic acid as this will lead to under oxidation and therefore no take up of the silver stain.

– The chromic acid can be reused but its efficiency will decrease after each use.

– If the control sections are failing to stain even after extending the staining time in the heated hexamine-silver solution you have more than likely forgotten to add the borax. If so, you can just add it and continue with the stain.

– If you have a large amount of silver precipitate over your sections it is probably due to using low-grade silver.

– If you forget the sodium thiosulphate step you will not realise until after retrieving the slide from storage further in the future, as the remaining silver not removed will react with sunlight turning black.

– Try to keep your counterstain fairly light as dark counterstaining can mask fungal elements.


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Perls’ Technique For The Demonstration of Haemosiderin – Method and Tips

Iron is absorbed in the duodenum by cells called enterocytes. It is then stored or combined with a transport protein molecule. This iron-protein complex is then taken to the bone marrow where the iron is incorporated into the substance known as haemoglobin which is involved in oxygen transportation.

Iron can be stored in the bone marrow and spleen in its ferric state (Fe3+) as haemosiderin when combined with protein. When haemoglobin is broken down by tissues this results in the formation of haemosiderin.

When there is excess iron in the body haemosiderin can be found deposited in organs that are involved with iron storage such as the spleen, bone marrow and liver. This condition is known as haemosiderosis.

A condition called haemochromatosis exists where the body indiscriminately absorbs iron resulting in the deposition of copious amounts of haemosiderin in many tissues.

Haemosiderin founds in histology sections is usually derived from the breakdown of damaged erythrocytes and can also be found absorbed by macrophages (siderophages).

The method used by the wide majority of histology laboratories for the demonstration of haemosiderin is the Perls’ technique. This method works by the hydrochloric acid (HCL) splitting off the bound protein which then allows the potassium ferrocyanide to bind with the Fe3+ and form ferric ferrocyanide (Prussian blue).

Below is the author’s favoured method.


2% hydrochloric acid (HCL)

2% aqueous potassium ferrocyanide

1% aqueous neutral red


1. Take sections to water

2. Mix equals parts of HCL and potassium ferrocyanide and filter onto sections. Leave for 15 minutes.

3. Wash for 5 minutes in running tap water.

4. Counterstain with neutral red for 1 minute.

5. Dehydrate, clear and mount.


– Always include a positive control.

– Stronger staining results can be found by carrying out step 2 at higher temperatures (e.g. 37-56°C). This can result in false positive results. This author has found room temperature to suffice.

– The washing step (step 3) should not be decreased below 5 minutes as thorough washing is required to prevent a heavy dye precipitate resulting from the neutral red counterstain.

– The author has found neutral red to be the best counterstain. Do not use safranin as this can stain the Prussian blue granules a dark purple colour.

– Always mount in a DPX-type mountant as other mounting media results in fading of the stain.

– A common artefact is the presence of blue granules on and around the section. This can be due to expired HCL or potassium ferrocyanide. It can also be due to iron contaminants in the tap water. This can be fixed by replacing all steps from the cutting of the sections to the mounting of the stained slide that involve tap water with distilled water.

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Modified Ziehl–Neelsen Stain For Leprosy Bacilli – Method and Tips

Leprosy bacilli in comparison with tubercle bacilli are much less acid- and alcohol-fast. The leprosy bacilli’s lipid envelope is also much more affected by the fat solvents traditionally used to dewax sections (i.e. Xylene). Due to these factors a modification on the standard Ziehl-Neelsen technique is used for the demonstration of leprosy bacilli.

Below is the author’s preferred technique.


Dewaxing solution – equal parts of liquid paraffin and rectified turpentine

Carbol Fuchsin – as per standard Ziehl-Neelsen technique

Methylene blue counterstain – as per standard Ziehl-Neelsen technique

10% sulphuric acid


1. Dewax in ‘dewaxing solution’ described above for 30 minutes.

2. Blot dry and wash in running water for approximately 10 minutes.

3. Stain with filtered Carbol Fuchsin for 30 minutes at room temperature.

4. Wash well in tap water.

5. Differentiate in 10% sulphuric acid until section is pale pink.

6. Wash well in tap water.

7. Counterstain with Methylene Blue for 15 seconds.

8. Wash well in tap water.

9. Blot dry, clear and mount.


– This author always puts sections on ‘sticky’ slides to prevent any floating off.

– There are many variations on the ‘softer dewaxing solution’ for the modified Ziehl-Neelsen technique for leprosy bacilli including:

– Two parts xylene to one part vegetable oil / clove oil / groundnut oil / olive oil / cottonseed oil.

– Residual oil on the section after washing prevents shrinkage of the section.

– Place slides directly from heater into dewaxing solution as this helps quicken the dewaxing

– Some methods use a weaker acid-alcohol solution for differentiation, but this author prefers 10% sulphuric acid as it is quicker.

– Don’t be alarmed when the section is placed into the sulphuric acid as it will turn a black colour. It will return to a pink colour when placed back in water.

– Ensure the counterstain colour isn’t too intense as this can mask some leprosy bacilli and even turn them a purple colour.

– This is author lets the sections dry after washing after counterstaining and then directly mounts them.

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Verhoeff Van Gieson Elastin Special Stain – Method and Tips

Elastin is a connective tissue protein which allows the tissues of the body to return to their original shape after distortion or stretching. Elastin fibres can be of varying size and diameter and are particularly well seen histologically in sites such as the lung, heart, blood vessels and the dermis.

Histological demonstration of elastin fibres (or lack of them) are important in diagnostic pathology for conditions such as arteriosclerosis, temporal arteritis and elastosis. Fine elastic fibres are not so easily seen on standard haemtoxylin and eosin (H+E) staining therefore special stains which demonstrate elastin clearly are vital.

There are many elastin special stain techniques such as Weigert-Type, Orcein, Aldehyde-Fuchsin and Verhoeff’s. The most common is Verhoeff’s technique of staining elastin due to its quick method and strong elastin colour result. Below is the author’s favoured method for demonstrating elastin which is a version of the Verhoeff’s.


Verhoeff’s solution – (5ml 5% alcoholic haematoxylin) + (2ml 10% aqueous ferric cholride) + (2ml Lugol’s iodine) MAKE IMMEDIATELY PRIOR TO USE.

Note – Lugol’s iodine = 2g potassium iodine dissolved in ~4ml of distilled water, then dissolve 1g iodine, then make up to 100ml.

2 % aqueous ferric chloride

Van Gieson counterstain = (100ml saturated aqueous picric acid) + (1% aqueous acid fuchsin), boil for 3 mins then filter.


1. Take sections to water.

2. Stain with Verhoeff’s solution for 15-20 mins

3. Wash well in tap water

4. Differentiate in 2% aqueous ferric chloride until only elastin fibres remain darkly stained.

5. Wash in tap water for 5 mins.

6. Counterstain with Van Gieson for 3 mins.

7. Dehydrate, clear and mount.


– aim for slight under-differentiation as the Van Gieson stain will continue the differentiation though more slowly.

– dehydrate quickly as alcohol can leach some of the Van Gieson stain from the section. You can accelerate dehydration by blotting the section with filter paper.

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Periodic-Acid Schiff Diastase (PASD) Special Stain – Method and Tips

In my previous post I covered the Periodic Acid-Schiff reaction (PAS) special stain which is by far the most common stain performed in a routine histology laboratory. A variation on this technique call the Periodic Acid-Schiff Reaction with diastase digestion (PASD) is another commonly performed special stain which I will be covering in this post.

The variation on the PAS technique involves simply exposing the section to the diastase enzyme amylase prior to continuing with the standard PAS method. The term ‘diastase’ refers to any enzyme that catalyses the breakdown of starch into maltose the dextrose. The diastase enzyme acts by cleaving the a-glucosidic 1-4 linkages of starch or glycogen (aka animal starch) leading to the formation of maltose and dextrose (maltose and dextrose are water-soluble sugars). So when sections are pre-exposed to diastase before commencing the PAS technique the glycogen within the tissue is broken down into maltose and dextrose which are dissolved and washed away when the section is rinsed sufficiently in tap water.

The diagnostic purpose of performing the PASD technique include

–         the removal of glycogen to make it easier to identify mucins stained by the PAS technique

–         analysis of glycogen deposits within the liver

–         highlighting a thickened basement membrane for example in lupus


Diastase solution – 1 part human saliva to 9 parts distilled water.

1% aqueous periodic acid – from PAS method.

Schiffs Reagent – from PAS method.




1. Take sections to water.

2. Expose sections to diastase solution for 30 minutes at room temperature.

3. Wash sections thoroughly in tap water

3. Continue with PAS method from step 2


– Always run a PAS and a PASD control with every batch of PASD stains to ensure your diastase solution is working. This author has found a glycogen rich liver control to be most sufficient. This author also runs a PAS and PASD for all PASD requests.

– Commercial amylase is available instead of using a saliva solution. Commerical amylase sometimes requires different incubation temperatures and conditions so check this before using. This author has found that a saliva solution is easiest due to its ease of preparation, availability and plus it is free.

– Put all sections onto to ‘sticky’ slides ie. Superfrost plus slides or their equivalent as the saliva solution causing some lifting of the section from the slide. This is reportedly more prevalent in sections exposed to the commercial amylase solution.

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Periodic-Acid Schiff (PAS) Special Stain – Method and Tips

The Periodic-Acid Schiff (PAS) technique (and its numerous variations) is by far the most commonly performed special stain within the histopathology laboratory, therefore knowledge of its method is a vital arrow in any medical scientist’s quiver of knowledge.

The PAS technique is most commonly used to highlight molecules with a high percentage carbohydrate content such as mucins, glycogen, fungi and the basement membrane in skin.

The PAS method works by exposing the tissue to periodic acid. This acts an oxidizing agent which oxidizes vicinal (neighbouring) glycol groups or amino/alkylamino derivatives. This oxidation creates dialdehydes.These dialdehydes when exposed to Schiff’s reagent create an insoluble magenta compound which is similar to the basic fuchsin dye within the Schiff’s reagent.


Schiffs reagent

1% aqueous periodic acid


1. Take sections to water.

2. Expose sections to periodic acid solution for 10-15 mins.

3. Rinse well in tap water.

4. Expose sections to Schiff’s reagent for 10-15 mins.

5. Wash in running tap water for 5-10 mins

6. Counterstain with a haemtoxylin for approx. 15 secs.

7. Differentiate (if necessary) and blue.

8. Dehydrate, clear and mount.


– Periodic acid and Schiff’s reagent are easily available commercially prepared, the technique for self-made Schiff’s reagent is arduous by comparison but can be found.

– Keep your Schiff’s reagent out of UV light and refrigerated when not in use. Failure to do so will result in the loss of sulphur dioxide in your Schiff’s reagent leading to the solution turning from colourless to a magenta colour resembling the original basic fuchsin colour. When this happens replace your solution. Also keep your periodic acid solution refrigerated when not in use.

– The purpose of washing in running tap water after exposing the sections to Schiff’s is to intensify the magenta colour. This author has found that when the water has runs from a magenta colour to a clear colour the colour isn’t going to intensify any further therefore the washing in running water can be ceased. This may vary from lab to lab.

– There are numerous variations of the PAS technique (eg. PAS + diastase, PAS + Alcian Blue). This will be discussed in a further blog post.

 Below is a PAS stain of a section of skin from the scalp.

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