Xylanase Protocols

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These protocols are used to screen for and measure xylanase activity.

Congo Red Staining

Congo Red is a dye that will bind to carbohydrate polymers (e.g. cellulose and xylan).


  • Agar medium of your choice
  • Congo-Red solution (1mg/ml in H2O) vortex well to mix
  • Destaining solution (1M NaCl)
  • Xylan (or carboxy-methyl-cellulose (CMC))


  1. Make agar plates containing 0.01% xylan
  2. Apply a droplet of overnight culture (or you can streak for colonies)
  3. Incubate for 16 hours.
  4. Add 10 ml of Congo-Red solution to the top of the plate
  5. Incubate for 15 minutes to stain xylan.
  6. Pour off congo-red.
  7. Add 10ml of 1M NaCl.
  8. Incubate for 15 minutes to destain the unbound regions.
  9. Observe clearing zones around xylanase positive colonies.



  • Teather and Wood. 1982. Use of Congo Red-Polysaccharide Interactions in Enumeration and Characterization of Cellulolytic Bacteria from the Bovine Rumen. Applied and Environmental Microbiology 4 p. 777-780
  • Scheirlinck et al. 1990. Cloning and expression of cellulase and xylanase genes in Lactobacillus plantarum. Applied Microbiology and Biotechnology (1990)33:534-541

RBB Plate Screen

This is a quick procedure to screen for xylanase activity.


  • 2.5g Xylan
  • 2.5g RBB (Remazol Brilliant Blue)
  • 60ml Water
  • 20ml Sodium Hydroxide Solution (1.5g in 20ml)
  • 20ml Sodium Acetate Solution (0.675g in 20ml)
  • 200ml 96% Ethanol
  • 1L Wash solution (660ml Ethanol, 330ml Water, 1.35g Sodium Acetate)
  • Acetone

Preparing the Dyed Xylan

  1. Add 2.5g RBB dye and 2.5g Xylan to 60 ml Water.
  2. Add 20ml Sodium Acetate solution to RBB-Xylan solution.
    1. Add dropwise over 5 minutes while stirring at room temp.
  3. After mixing add 20ml Sodium Hydroxide solution.
  4. Stir for 90 minutes at room temperature.
  5. Add two volumes of 96% Ethanol to precipitate the xylan-RBB.
  6. Filter using a vacuum flask and watman filter paper.
  7. Wash the precipitate sequentially with 1L of wash solution.
    1. The filtrate should now be colorless.
  8. Wash the precipitate with 100ml 75% Ethanol.
  9. Wash the precipitate with 50ml Acetone.
  10. Dry at room temp overnight.


  1. Add 0.2% RBB-Xylan (weight per volume) to your agar media of choice and autoclave.
  2. Pour the plates immediately being sure to swirl frequently.
  3. Allow to solidify.
  4. Drop 5μL of overnight culture onto the plates (This will look like a large round colony).
  5. Incubate for two days.
  6. Observe clearing zones around xylanase producing cultures.

RBB Assay

This assay functions on the release of ethanol soluble RBB when the RBB-Xylan is cleaved by xylanase. To be sure that there is activity it can be useful to do duplicate samples and measure after 1 hour and then again after 2 hours.


  • RBB-Xylan (made according to the procedure outlines above)
  • Cell culture (containing the xylanase)
  • water


  1. Prepare a 0.1M Acetate buffer containing 11.5mg/ml RBB-xylan
  2. Add 250ml of RBB-xylan solution to 250ml cell culture
  3. Incubate for 2 hours at 30°C
  4. Add 1ml Ethanol to stop the reaction and precipitate RBB-Xylan
  5. Centrifuge for one minute at maximum speed
  6. Collect supernatant
  7. Measure absorbance at 595nM
  • Compare to medium with a no xylanase control
  • Higher absorbance means more xylanase activity

Somogyi Nelson Assay

This assay measures the release of reducing sugars (including xylose and arabinose) during the enzymatic treatment of xylan. If you have reducing sugars (e.g. glucose) in your media, it will also be detected, so you will always have to compare to a control culture with no xylanase.


  • Copper Sulfate (pentahydrate)
  • Ammonium Molybdate (tetrahydrate)
  • Sodium Acetate (anhydrous)
  • Sodium Bicarbonate (anhydrous)
  • Sodium Carbonate (anhydrous)
  • Potassium Sodium Tartrate (heptahydrate)
  • Sodium Sulfate (anhydrous)
  • Sodium Arsenate (dibasic heptahydrate)
  • Sulfuric Acid (72%)
  • Xylan (purified)
  • Xylose (purified anhydrous)

Preparing the Reagents

If you prepare the amounts listed you will be able to run 10 samples (not including the standard curve)

Standard solution

  • This solution is 1mg/ml xylose and you will need 250μL of it.

Substrate Solution

1. Dissolve the following in 10ml of water:
  • 82mg Sodium Acetate
  • 0.2g Xylan
2. Bring the volume up to 20ml by adding water.
3. Adjust the pH to 4.5 using HCl.

Copper Solution

1. Dissolve the following in 10ml of water:
  • 80mg Copper Sulfate
  • 0.32g Sodium Bicarbonate
  • 0.48g Sodium Carbonate
  • 0.25g Sodium Potassium Tartrate
  • 3.7g Sodium Sulfate
2. Bring the volume up to 20ml by adding water.

Acid Solution

1. Dissolve the following in 10ml of water:
  • 54mg Arsenic Acid
  • 0.13g Molybdic Acid
  • 1.26ml 72% Sulfuric Acid (12M Sulfuric Acid)
2. Bring the volume up to 20ml by adding water.


Standard Curve

Prepare the following samples:
Blank sample : 1.90 ml of xylan substrate
Standard 1 : 20ul of xylose standard and 1.88ml of water and 100ul of MRS
Standard 2 : 50ul of xylose standard and 1.85ml of water and 100 ul of MRS
Standard 3 : 70ul of xylose standard and 1.83ml of water and 100 ul of MRS
Standard 4 : 100ul of xylose standard and 1.80ml of water and 100 ul of MRS


Add 100ul of cell culture (in MRS) to 1.9ml of xylan substrate.
1. Swirl sample, standards, and blank, and bring to 30°C.
2. Add 100ul of water to the blank sample, swirl, and incubate for 10 minutes at 30°C.
3. Add 2ml of copper solution to each standard and blank sample and swirl each solution.
4. Place a marble over each tube and place each tube in a boiling water bath for 10 minutes.
5. Allow tubes to cool to room temperature.
6. Add 2ml of acid solution to each tube and vortex until all precipitate is dissolved and foaming has stopped. Centrifuge tubes.
7. Transfer solutions to cuvettes and measure the absorbance at 540nm.