Periodic Acid-Schiff (PAS) Staining Kit 

Glycogen staining is one of the classical staining methods widely used in histological and pathological studies. In 1946, McManus first introduced and applied the Periodic Acid–Schiff (PAS) staining technique for the detection of mucopolysaccharides in tissues. Owing to its strong chromogenic capability toward a wide range of polysaccharides and glycoconjugates, this method has been extensively used for the detection of glycogen and other polysaccharide-containing substances.

The basic principle of PAS staining is based on the use of periodic acid as a strong oxidizing agent. Periodic acid specifically oxidizes the vicinal diol (1,2-glycol) groups present in carbohydrate molecules to generate aldehyde groups. The newly formed aldehydes subsequently react with the Schiff reagent, producing a stable magenta (reddish-purple) complex that can be clearly observed under a microscope. Through this reaction, tissue structures rich in polysaccharides or glycoconjugates can be distinctly visualized.

During the PAS staining process, the concentration of periodic acid and the oxidation time are critical factors affecting staining performance. In addition to oxidizing polysaccharide structures, periodic acid may also oxidize other cellular components. Therefore, oxidation conditions must be carefully controlled so that vicinal diols are efficiently converted into aldehydes while avoiding over-oxidation, which could further convert aldehydes into carboxyl groups. Such over-oxidation would prevent subsequent binding with the Schiff reagent. Proper control of these conditions is therefore essential to ensure the specificity and stability of the staining signal.

MCE Periodic Acid-Schiff (PAS) Staining Kit is developed based on the classical Periodic Acid–Schiff (PAS) reaction principle and is suitable for the detection of glycogen and polysaccharide-containing substances in paraffin-embedded or frozen tissue sections.

Through a standardized combination of staining reagents and an optimized staining system, this kit enables clear visualization of glycogen and related polysaccharide structures in tissue sections. It is commonly used for observing glycogen distribution in tissues such as liver and muscle, and is also applicable to basement membrane visualization, fungal detection, and pathological studies involving polysaccharide deposition. The method features simple operation and good reproducibility, providing reliable morphological evidence for histological research and pathological analysis.

Periodic Acid Solution, Schiff Reagent: 4°C, 1 year. Protected from light.

Hematoxylin Staining Solution, Acidic Ethanol Differentiation Solution: RT, 1 year.

Reagents Preparation

1. Fixative: 10% formalin

2. Other reagents: xylene, neutral mounting medium or mounting reagent, graded ethanol solutions, distilled water, etc.

Procedure

1. Tissue Fixation and Embedding: Fix tissue samples using standard protocols. 10% neutral formalin is generally recommended. After fixation, dehydrate and embed the tissues in paraffin following routine histological procedures.

2. Deparaffinization and Rehydration: For paraffin sections, deparaffinize sequentially with xylene, then rehydrate through a graded ethanol series, and finally immerse in distilled water. Frozen sections can be directly placed in distilled water.

3. Rinsing: Wash the sections in tap water for 2-3 min, followed by two rinses in distilled water.

4. Periodic Acid Oxidation: Immerse sections in periodic acid solution and incubate at room temperature for 5-8 min (generally not exceeding 10 min).

5. Rinsing: Rinse once in tap water, then twice in distilled water.

6. Schiff Reagent Staining: Incubate sections in Schiff reagent at room temperature in the dark for 10-20 min, then rinse in tap water for approximately 10 min.

7. Nuclear Counterstaining: Stain nuclei with hematoxylin for 1-2 min, followed by differentiation in Acidic Ethanol Differentiation Solution for 2-5 s.

8. Bluing: Rinse in tap water for 10-15 min, then replace with distilled water to ensure full nuclear bluing.

9. Dehydration, Clearing, and Mounting: Dehydrate sections through a graded ethanol series, clear with xylene, and mount with neutral resin or mounting medium. Observe results under a microscope.

Interpretation of Results

1. PAS-positive substances (e.g., glycogen or other polysaccharides): red to magenta.

2. Cytoplasm: blue.

3. Nuclei: varying shades of red.

Note: The intensity of staining largely depends on the duration of oxidation in periodic acid and staining with Schiff reagent. Proper control of oxidation and staining times is critical to achieve clear, specific, and stable PAS staining.

(Optional) Negative Controls

1. Amylase Digestion Control: Dissolve 1 g of amylase in 100 mL PBS (pH 5.3) to prepare the amylase solution. Incubate tissue sections in this solution for 30-60 min, and then process them together with the experimental sections in the periodic acid solution for subsequent staining. As glycogen is digested by amylase, the PAS reaction should yield a negative result.

2. (Alternative Method) Saliva Digestion Control: Treat sections with filtered saliva for 30–60 min, and then subject them to the periodic acid solution together with the other sections. Since saliva contains amylase capable of digesting glycogen, the PAS staining result should be negative.

3. (Alternative Method) Oxidation Step Omission Control: When using sections from the same tissue sample as controls, the periodic acid oxidation step can be omitted. In this case, the control sections are directly incubated with Schiff reagent without periodic acid treatment. Because aldehyde groups are not generated, no PAS-positive staining should occur (negative result).

1. This product is suitable for staining routine tissue sections and is not recommended for fungal samples, single-cell preparations, or extremely thin sections.

2. It is recommended to fix samples with 10% neutral buffered formalin to ensure optimal preservation of tissue morphology and structure.

3. Since ethanol is highly volatile, graded ethanol solutions should be freshly prepared before use to ensure consistent dehydration performance.

4. Deparaffinization of sections should be thorough; incomplete deparaffinization may compromise staining quality.

5. The oxidation time with periodic acid should not be excessive; the optimal oxidation temperature is 18-22°C.

6. Periodic acid solution and Schiff reagent should be stored at 4°C in a tightly sealed container. Avoid prolonged exposure to light and air during use. Prior to use, it is recommended to equilibrate the reagents to room temperature for 30 minutes while keeping them protected from light.

7. Acidic ethanol differentiation solution should be frequently refreshed. Differentiation time should be adjusted based on section thickness, tissue type, and the age of the differentiation solution. Sufficient rinsing with tap water is required after differentiation.

8. The incubation times in periodic acid and Schiff reagent are critical and should be optimized according to section thickness and tissue type.

9. Staining time for frozen sections should be minimized to avoid overstaining.

10. Once opened, reagents should be used promptly to ensure consistent staining performance in subsequent experiments.

11. After staining with this kit, subsequent immunofluorescence or other fluorescence-based staining is not recommended, as it may interfere with fluorescence signals or the interpretation of staining results.

12. This product is for R&D use only, not for drug, household, or other uses.

13. For your safety and health, plea