Detecting Glycogen in Peripheral Blood Mononuclear Cells with Periodic Acid Schiff Staining

Summary

Periodic acid Schiff staining is a technique that visualizes the polysaccharide content of tissues. This article demonstrates periodic acid Schiff staining protocol adapted for use on peripheral blood mononuclear cells purified from human venous blood. Such samples are enriched for lymphocytes and other white blood cells of the immune system.

Abstract

Periodic acid Schiff (PAS) staining is an immunohistochemical technique used on muscle biopsies and as a diagnostic tool for blood samples. Polysaccharides such as glycogen, glycoproteins, and glycolipids stain bright magenta making it easy to enumerate positive and negative cells within the tissue. In muscle cells PAS staining is used to determine the glycogen content in different types of muscle cells, while in blood cell samples PAS staining has been explored as a diagnostic tool for a variety of conditions. Blood contains a proportion of white blood cells that belong to the immune system. The notion that cells of the immune system possess glycogen and use it as an energy source has not been widely explored. Here, we describe an adapted version of the PAS staining protocol that can be applied on peripheral blood mononuclear immune cells from human venous blood. Small cells with PAS-positive granules and larger cells with diffuse PAS staining were observed. Treatment of samples with amylase abrogates these patterns confirming the specificity of the stain. An alternate technique based on enzymatic digestion confirmed the presence and amount of glycogen in the samples. This protocol is useful for hematologists or immunologists studying polysaccharide content in blood-derived lymphocytes.

Introduction

Periodic acid Schiff (PAS) staining is an immunohistochemical technique that is widely used in muscle research and diagnostics. It is also utilized as a diagnostic tool on blood samples. The technique works by applying periodic acid solution to the sample, which oxidizes units within the polysaccharide creating aldehyde groups that react with the colourless Schiff's reagent thereby producing a deep magenta product. The steps of this procedure are shown in Figure 1. The stain turns anything with polysaccharides magenta, including glycogen, glycoproteins, glycolipids, mucins, or other molecules with polysaccharide moieties.

PAS staining is often used to measure glycogen levels in muscle fibers. Muscles tissue sections are ideal for the technique as they firmly attach to the slide and withstand multiple washing and staining steps. Glycogen is most present in fast twitch Type II muscle fibers, which have a high demand for rapid ATP production requiring glycogen for maximum performance^1,2. Glycogen is a branched polymer of glucose that can be broken into free glucose through the action of glycogen phosphorylase enzymes. In times of rest and nutritional-sufficiency, glycogen is replenished through the process of glycogenesis, while in times of nutritional insufficiency or high-energy demand; glycogen is broken down into glucose by glycogenolysis. From as early as the 1950's clinician scientists have explored PAS staining on blood samples to analyze glycogen content in various diseases^3-7. For example, in Pompe disease-a bonafide glycogen storage disease- white blood cells accumulate large amounts of glycogen that differs significantly from healthy controls⁸.

This video-article demonstrates an adapted version of PAS staining for use on peripheral blood mononuclear cells (PBMC) samples from venous blood of healthy human subjects. PBMCs contain mostly lymphocytes of the T lymphocyte and B lymphocyte families, as well as other immune cells such as natural killer cells and monocytes. The first purification step removes erythrocytes, neutrophils, and other granulocytes. This technique provides data on a concentrated proportion of lymphocytes allowing for more robust enumeration of PAS-positive cells as compared to using whole blood smears.

Figure 1: Step by step methodology of PAS staining on PBMC. (A) First, isolation of PBMC is achieved through ficoll gradient, the left panel shows the preparation before centrifugation, the right panel shows it after centrifugation where the buffy coat containing the PBMC is observed in the center of the tube. (B) Isolated PBMCs are fixed onto the slide using formalin-ethanol fixative solution. The slide is gently rinsed with distilled water from a plastic wash bottle. (C) The slide is then placed in a 100 ml beaker half way filled with amylase solution, which will dissolve glycogen. The slide is gently rinsed. (D) The slide is treated with periodic acid solution, where oxidation of saccharides takes place. Slides are gently rinsed; this will remove the excess periodic acid and stop the oxidation step. (E) When the Schiff reagent is added to the slides, it will react with aldehydes created during the oxidation step. This colorless reagent will then result in a deep red magenta product. Slides are gently rinsed to remove the excess Schiff reagent.

Protocol

Research with human blood samples was approved by Concordia University Ethics Review Board, certificate number 10000618. The work on mouse muscle was approved by Concordia University Ethics Review Board, certificate number 2010BERG.

1. PBMC Isolation from Whole Blood

NOTE: Carry out this procedure in a biosafety cabinet using sterile technique and manufacturer-sterilized equipment.

1. Carefully pour 10-15 ml of whole blood from the heparinized (anti-coagulant) blood collection tubes into a 50 ml sterile conical tube. For minor blood spills, wipe with ddH₂O and 70% EtOH using cleaning tissue.
2. Dilute the blood in the conical tube to a 1:1 ratio with Phosphate Buffer Saline (PBS 1x) pH 7.4. Ensure that the maximum total volume does not exceed 30 ml.
3. Mix gently using a serological pipette gun avoiding bubbles.
   NOTE: Do NOT mix by inverting the tube to avoid blood accumulation in the cap and subsequent seepage during centrifugation.
4. Add 13 ml of Ficoll-paque (see Materials and Equipment table), to a new 50 ml capacity conical tube. Keep the tube upright in the rack.
5. Using a transfer pipette, take diluted blood, touch the transfer pipette tip to the inside wall of the tube near the top. With slow and steady pressure, transfer the blood along the inside wall of the tube forming a blood layer on top of the sucrose layer. Do this step several times until all blood has been transferred.
6. Cap the tube tightly and centrifuge the tube at room temperature for 30 min at 700 x g in a swing rotor with medium acceleration set to 5, and deceleration set to ZERO.
7. Slowly take out the tube and without disturbing the layers, take it back to the biosafety cabinet.
8. Carefully collect the buffy coat (thin cloudy white layer), where PBMCs are located, placed between the PBS/plasma and ficoll layers using a transfer pipette. Avoid collecting sucrose layer and do not disturb the red blood cell layer.
9. Transfer the buffy coat into a new 50 ml sterile conical tube. It may take several times of repeating step 1.8 to collect all the PBMC.
10. Add PBS pH 7.4 to the tube with the PBMC and fill up to the 45 ml mark. Shake tube thoroughly. Do not vortex.
11. Wash 1: Centrifuge for 15 min at 480 x g with both maximum acceleration and deceleration set to 9. Observe formation of a pellet of PBMCs at the bottom of the conical tube.
12. Discard the PBS (supernatant) into a plastic beaker with 10 ml of bleach.
13. Loosen cell pellet by gently “racking” against an undulated surface (i.e., empty rack). Do not vortex.
14. Add 25 ml of fresh PBS pH 7.4 to the tube. If more than one tube is being processed, pool the pellets together in this step.
15. Wash 2: Centrifuge the tube for 12 min at 480 x g with both maximum acceleration and deceleration set to 9.
16. Discard the PBS (supernatant) into the plastic beaker with a splash of bleach.
17. Gently “rack” against an undulated surface (i.e., empty rack). Do not vortex.
18. Add 25 ml of fresh PBS to the tube.
    1. Take out 50 µl of the cells and transfer into a microcentrifuge tube for viability count.
    2. Add an equal amount (50 µl) of trypan blue (a viability stain) and pipette up and down to mix gently. 
19. Take out 10 µl and transfer it to a hemocytometer to check the viability of cells per ml. Record the number of cells/ml.
20. Take out the desired amount of cells and centrifuge the tube for 12 min at 480 x g with both maximum acceleration and deceleration set to 9.
21. Discard the PBS (supernatant) into the beaker with bleach.
22. Gently “rack” against an undulated surface (i.e., empty rack). Add 80 µl of PBS into the tube.

2. Making the PBMC Slide

1. Place 80 µl of the cells onto a microscope slide. Smear the drop with the help of another slide or place 2 drops of 40 µl each on both ends of the slide.
2. Leave slide in the biological safety cabinet to dry. Label the slide with a pencil on the frosted side.

3. Fixing the Samples on the Slides

1. Prepare the fixative solution by mixing 0.5 ml of 37% formaldehyde to 4.5 ml of 99% ethanol.
2. After the slides are dried, take out 2 ml of the freshly made fixative solution and pour it on the slide so that the entire surface of the slide is covered.
3. Leave the solution on the slide for 1 min. Rinse the slide for 1 min with tap water and leave it to air dry.

4. Making the Amylase Solution for Negative Control

1. Take 0.25 g of amylase powder and dissolve in 50 ml of distilled water. Pour the solution in a clean 100 ml beaker.
2. Immerse the slide in the beaker so that half of the slide receives the treatment and the other half remains untreated and then incubate for 15 min at room temperature (Figure 1C).
3. Make a note on which side of the slide is receiving the amylase treatment. Draw a line on the back of the slide indicating the border between the treatment and control.
4. Wash the slides with ddH₂O to remove the amylase solution and leave the slide to air dry.

5. Perform Periodic Acid Schiff (PAS) Staining and Imaging

NOTE: PAS reagents are toxic by inhalation and are corrosive, so the steps need to be done in a chemical fume hood, and the waste products must be properly disposed of according to institutional guidelines.

1. Place the slide on a flat surface and pour 1.50-2.00 ml of Periodic Acid Solution on the sample. Incubate for 5 min at room temperature.
2. Rinse the slides in several charges of distilled water.
3. Pour 1.50-2.00 ml of Schiff’s reagent on the slide and incubate it at room temperature for 15 min.
4. Wash the slide with distilled water for 5 min and leave it to air dry.
5. Apply 10 µl mounting media on the slide and cover with two small coverslips, or apply 50 µl and use one large coverslip.
6. Apply clear nail polish on the edges of the coverslip, let dry overnight.

6. Obtain Images with the Binocular Light Microscope Using the 100X Objective

Results

To validate the reagents and basic technique, PAS staining was performed according to manufacturer’s instructions on mouse soleus muscle sections. The staining was done same day as the sacrifice and in the last step of the staining, the sections were fixed by xylene. The soleus muscle is known to contain ~35% glycogen-positive cells⁹. The stained muscle cells displayed two distinct PAS-positive features- punctate granules within the cell, and a continuous line demarking the cell membrane (Figure 2A

Discussion

The critical steps of this video article were during washing and amylase treatment of the cells. While washing the slides, the key step was using a plastic squeezable washing bottle and letting the water gently run through the sample on the slide and not aiming directly onto the samples. Even the slightest direct water pressure would cause the cells to come off the slide. Another key step was to use the same slide for ± amylase conditions. After the PBMCs were adhered to the slide, the slide was carefully placed int...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Periodic Acid Shiff Kit	Sigma-Aldrich	395B	Bring to room temperature prior to use. Materials in this kit are toxic and harmful. Use caution.
α-Amylase from porcine pancreas	Sigma-Aldrich	A3176
Binocular Microscope	Carl Zeiss Microscopy	Axio Lab A0
Glycogen Assay Kit	Sigma-Aldrich	MAK016
Ficoll-Paque PLUS	VWR, GE Healthcare	17-1440-02	Nonionic synthetic polymer of sucrose.
Centrifuge			For PBMC isolation, swing buckets were used.