RSS

Category Archives: Uncategorized

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.

Follow me on twitter (@skinpathology)

Keep an eye out for my website coming soon (www.skinpathonline.com)

Feel free to email me with any questions or queries on feedback@skinpathonline.com

Advertisements
 

Tags: , , , , , , , , , , , , , , , , , , , , , , , , , , ,

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.

SOLUTIONS

2% hydrochloric acid (HCL)

2% aqueous potassium ferrocyanide

1% aqueous neutral red

METHOD

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.

TIPS

– 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.

Thanks for reading and I welcome any comments.

Follow me on twitter (@skinpathology)

Keep an eye out for my website coming soon (www.skinpathonline.com)

Feel free to email me with any questions or queries on feedback@skinpathonline.com

 

Tags: , , , , , , , , , , , , , , , , , , , , , , , , , , ,

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.

Follow me on twitter (@skinpathology)

Keep an eye out for my website coming soon (www.skinpathonline.com)

Feel free to email me with any questions or comments on feedback@skinpathonline.com

 

Tags: , , , , , , , , , , , , , , , , , , , , , , , , , , , , , , ,

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.

SOLUTIONS

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.

METHOD

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.

TIPS

– 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.

Thanks for reading and I welcome any comments.

Any other questions or queries email me on feedback@skinpathonline.com

Follow me on twitter (@skinpathology)

Keep an eye out for my website coming soon (www.skinpathonline.com)

 

Tags: , , , , , , , , , , , , , , , , , , , , , , , ,

Persistent Melanocytic Nevi: A Review and Analysis of 205 cases


Great article in the June 2011 issue of the ‘Journal of Cutaneous Pathology’ regarding persistent melanocytic naevi. Good reading for those interested in the subject.

Things of note are

– female predominance (reason unclear)

– back is the most common site followed by abdomen then chest

– mean time between original biopsy then biopsy of persistent naevus was 9.7 months

– dysplastic naevi were most likely to recur

– persistent melanocytic naevi were more likely to be initially removed via shave biopsy

 

Link to the article below

http://onlinelibrary.wiley.com/doi/10.1111/j.1600-0560.2011.01692.x/abstract

Thanks for reading and I welcome any comments

Keep an eye out for my website COMING SOON (www.skinpathonline.com)

Follow me on twitter (@skinpathology)

 

 

 

 

Tags: , , , , , , , , , , , , , , , , , , , , , , , , , , ,

Melanoma Research – Sex differences in survival of cutaneous melanoma are age dependent


Came across an interesting article in the latest issue of the Melanoma Research journal regarding differences in survival rates based on sex. It has been previously observed that women have a better survival rate for melanoma than men. This has also been observed in other cancers such as lung adenocarcinoma and colon cancer.

The study reveals that the slight survival benefit women with melanoma experience, disappears after the age of 60. This is mirrored, but also conflicts with other studies referenced within the article.

Proposed reasons for this female survival benefit include women being more prudent in the personal examination of the skin, women having a greater percentage of lower limbs melanomas which are associated with a better prognosis and immune gender differences.

Below is a link to the article abstract

http://journals.lww.com/melanomaresearch/Abstract/2011/06000/Sex_differences_in_survival_of_cutaneous_melanoma.11.aspx

 I recommend getting the whole article if it is possible.

 Thanks for reading and I welcome any comments.

 Follow me on twitter (@skinpathology)

 Keep an eye out for my website coming soon (www.skinpathonline.com)

 

Tags: , , , , , , , , , , , , , , , , , , , , , ,

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.

Follow me on twitter (@skinpathology)

Keep an eye out for my website (www.skinpathonline.com) COMING SOON.

 

Tags: , , , , , , , , , , , , , , , , , , , , , , , , , ,