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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.

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.

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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

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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.

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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.

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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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Alcian Blue Stain For Acidic Mucins – Method and Tips


The alcian blue stain is this author’s preferred stain for the demonstration of acidic mucins. The dye was originally used for the dyeing of cotton before being discovered as a by Steedman in 1950.

The alcian blue itself is a cationic copper phthalocyanine dye which stains mucopolysaccharides and glycosaminoglycans a bluish colour. Within skin, acidic mucins can be found in many differing conditions such as a mucinoma, lupus and alopecia mucinosa.

Below is the preferred alcian blue method of this author

METHOD

Solutions

1g alcian blue in 3% acetic acid (check pH = 2.5)

1% safranin

1. Take sections to water

2. Cover slide with FILTERED alcian blue solution and leave for 20 minutes

3. Rinse in tap water

4. Counterstain with FILTERED 1% safranin for 10-15 seconds

5. Rinse in tap water

6. Dehydrate quickly, clear and mount.

Tips

– this author prefers an alcian blue staining time of about 20 minutes but can be done within the range of 10-30 minutes if desired.

– by reducing the pH to 0.2 the stainer can select for only strongly sulphated mucins. A pH of 1.0 stains both weak and strongly sulphated mucins. If using a lower pH method be sure no to rinse in tap water between the steps for too long as this can affect the alcian blue staining.

– this author prefers safranin as a counterstain due to its crisper staining, but safranin leeches out quickly in the dehydrating alcohols therefore blot dry after counterstaining and quickly dehydrate through the alcohols.

– neutral red can also be used as a counterstain.

– the alcian blue staining solution expires after approximately 6 months.

Thanks for reading and I welcome any comments and other tips.

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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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