Periodic acid Schiff- diastase (PAS – Diastase technique) is a test method of using diastase enzyme in combination with PAS stain.
Diastase is an enzyme. It breaks down glycogen.
PAS – Diastase technique stain is a stain that useful for the staining of glycogen, glycoproteins, glycolipids, and mucins in tissue sections.
When using PAS technique, it is difficult to distinguish mucins from glycogen.
Therefore, a glycogen digestion step is added to the PAS staining procedure.
It is called as PAS – Diastase technique.
Alpha-amylase (α-amylase) may be used for this.
Hydrolysis of glycosidic bonds of glycogen is catalyzed by alpha-amylase.
By the breakdown of large glycogen molecules, water-soluble disaccharide known as maltose is produced.
By this method removal of glycogen from the tissue section is done prior to PAS staining.
PAS used as a stain for glycogen, glycoproteins, glycolipids, and mucins in tissue sections.
When diastase is added to the tissue sections, diastase act on glycogen and glycogen is breakdown into water soluble sugar namely maltose.
This water-soluble sugar washed out from the section.
Periodic acid solution oxidizes 1,2-glycol groups producing dialdehyde.
By treating with Schiff’s reagent, the resultant dialdehyde is precipitated as insoluble magenta colour complex.
When the glycogen has breakdown by diastase, magenta colour precipitate cannot be seen.
Instead of that pale pink colour background can be seen.
Preparation of periodic acid solution
Mix 1 g of periodic acid with 100 ml of distilled water and dissolve well
Preparation of Schiff reagent
- Liquidate 1 g of basic fuchsin and 1.9 g of sodium metabisulfite (Na2S2O5) in 100 ml of 0.15 M HCl (hydrochloric acid).
- Then shake the solution well.
- Shaking should be done at intervals on a mechanical shaker for 2 hours.
- The solution should be clear and colour should be yellow to light brown.
- After that add 500 mg of activated charcoal.
- Then shake 1-2 minutes.
- Next filter the solution using a no 1 Whatman filter paper into a bottle.
- This solution should be clear and colourless.
- If the solution is yellow in colour, repeat the charcoal decolourization using a fresh lot of activated charcoal.
- This solution should be stored at 4°C and it is stable for several months.
Dissolve 1.97 g Sodium phosphate (monobasic) and 0.28 g of Sodium phosphate (dibasic) in 1000 ml of distilled water.
Dissolve 0.1 g Malt diastase in 100 ml of phosphate buffer
- First dewax the slides and bring sections down to the water through graded alcohol solutions.
- Then treat one slide with diastase solution for 1 hour at 37°C.
- The other slide is an untreated control and may remain in water for 1 hour (The sections are also can be treated with 1% diastase at 56°C for 10 minutes).
- Then wash both slides in tap water for 5-10 minutes.
- Then treat with 1% periodic acid for 10 minutes.
- Then wash with tap water.
- Next cover the sections with Schiff reagent for 15 minutes.
- Rinse in running tap water for 5-6 minutes.
- Then stain with Carazzi’s haematoxylin for 3 minutes.
- Blue the nucleus in running tap water for 5-10 minutes.
- Finally dehydrate the sections, clear and mount with DPX.
Glycogen should be stained in magenta colour in untreated slide.
Glycogen stained magenta colour cannot be seen in the diastase-treated slide.
Malt diastase, which contains both α- and β-amylases is widely used in this method.
Moreover, human saliva is also effective for the digestion of glycogen in tissue sections.
However, due to safety problems and lack of standardization human saliva is not used widely for this method.
For this method two slides should be used.
A known positive control slide should be used to verify the potency of the enzyme.
Commercially available diastases are available.
However, activity and purity may be varied from batch to batch.
Contamination of the enzymes should be avoided.
Because they can be digested materials other than glycogen.
No responses yet