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Tag Archives: microbiology

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.

Solutions

– 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.

Method

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.

Tips

– 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.

Thanks for reading and I welcome any comment.

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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.

Solutions

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

Method

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.

 

Tips

– 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.

 

Thanks for reading and I welcome any comments

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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.

SOLUTIONS

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

METHOD

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.

TIPS

– 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.

I welcome any other tips and comments.

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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

Solutions

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

1% aqueous periodic acid – from PAS method.

Schiffs Reagent – from PAS method.

 

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

Tips

– 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.

Thanks for reading and I welcome any comments.

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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.

SOLUTIONS

Schiffs reagent

1% aqueous periodic acid

METHOD

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.

TIPS

– 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.

Thanks for reading and I welcome any comments.

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Ziehl–Neelsen Stain For Acid-Fast Organisms – Method and Tips


The Ziehl–Neelsen (ZN) stain is a common standard stain which is readily performed in a majority of histopathology laboratories around the world. It was first described by Dr. Franz Ziehl and Dr Friedrich Neelsen, a German bacteriologist and a German pathologist respectively. The ZN stain is mostly used to identify acid-fast mycobacteria, the most important of which is Mycobacterium Tuberculosis, the organism responsible for tuberculosis (TB). The ZN stain also stains other organisms such as Nocardia.

As Mycobacterium are unable to be visualised on standard haematoxylin and eosin (H+E) and gram stains, the ZN stain was developed. It is based on the tubercle bacilli having a lipid-rich cell wall that takes up phenol-dye solutions (eg. carbol fuchsin, the main dye used in the ZN stain) and after subsequent differentiation, retains the phenol-dye.

Below is the method used by this author.

Solutions

Carbol fuchsin – (1g basic fuchsin in 10ml ethanol) + (5g phenol in 100ml distilled water), then filter.

Methylene Blue – 0.2% methylene blue

Method

1. Take sections to water.

2. Cover section with filtered carbol fuchsin for 20 minutes.

3. Wash well in tap water.

4. Differentiate in 1% acid alcohol until section is a very pale pink.

5. Wash well in tap water.

6. Stain with methylene blue for 1 minute.

7. Dehydrate, clear and mount.

Tips

– Before covering section with carbol fuchsin try covering the section with a little filter paper to reduce precipitate on the slide.

– Some methods still say to the slide to steaming temperature after covering it with carbol fuchsin. This author has found this of no use and is an unnecessary extra step, plus removes the hazard of using a naked flame.

– Before differentiation with acid alcohol wash slide with 70% alcohol for about 1 minute to remove a majority of the stain. This will reduce your differentiation time.

– Blot dry your slide after washing in water after the methylene blue counterstain. This will reduce your dehydration time and therefore result in less leaching of the methylene blue counterstain from the section.

– Some tap water contaminants have been described that stain with carbol fuchsin and are resistant to differentiation. These appear on a different focal plane to true acid-fast organisms within the section.

I welcome any other tips and comments.

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