Mycorrhizal Roots Staining and study

In This Article we discuss the:

Clearing and staining roots for examination of arbuscular mycorrhizas, ericoid mycorrhizas, orchid mycorrhizas, and dark septate endophytes:

  1. Mycorrhizal Roots should be cleaned of soil and are usually fixed in 50% ethanol for at least 24 h. Roots can be cleared without prior fixation.
  2. Rinse 3× in water (deionized if available).
  3. Clear in either 5% or 10% KOH (depending on the fragility of roots) by either autoclaving for 20–40 min or placing containers in a water bath at 90°C for 2–3 h. Some very fine roots can be cleared at room temperature if left overnight or longer.
  4. Rinse 3× in water (deionized if available).
  5. If Mycorrhizal Roots roots are very pigmented they will need to be bleached (30–35% hydrogen peroxide: distilled water (1:1) and then add ammonium hydroxide for a final concentration of 0.05%). The length of time will depend on roots but usually, about 5 min is sufficient.
  6. Rinse 3× in water.
  7. Acidify in 2% HCl or 2% lactic acid for 1–2 min.
  8. Stain in either 0.03% Chlorazol Black E (made up of a solution of 80% lactic acid: glycerin in distilled water 1:1:1) for 2–3 h at 90°C or 0.05% Trypan blue made up as for Chlorazol Black E or 0.1% acid fuchsin made up in the same way. Staining with the latter two stains may require more time.
  9. Staining with acid fuchsin has been particularly useful when the material is to be examined by confocal laser scanning microscopy.
  1. De-stain in 50% glycerin for approx. 24 h.
  2. Mount roots on slides in 50% glycerin.

Embedding material for light microscopy:

To obtain uniform thin sections of a specimen it is necessary to embed the material in resin so that it can be sectioned with glass or diamond knives using a micro-tome.

After infiltration and polymerization in a resin, the tissue and the surrounding medium is of equal density. There are many protocols for this
type of work, depending on the type of investigation (see Ruzin 1999).

For most light microscopic work, we have routinely used the following method:

  1. Prepare 2.5% glutaraldehyde in either 0.1M HEPES or 0.1M Sorensen’s phosphate buffer and adjust the pH to 6.8. This must be done in a FUME HOOD.
  2. Cut specimens into small pieces (usually a few mm in length) and place them in vials containing buffered glutaraldehyde, again in a FUME HOOD.

Fix tissues for a minimum of 3 h at room temperature (can be left at 4°C overnight or longer).

  1. Remove the glutaraldehyde with a Pasteur pipette in a FUME HOOD, placing this solution in a waste- container.
  2. Dehydrate the tissue by means of a graded ethanol series:
  • 50% for 20 min
  • 70% for 20 min
  • 90% for 30 min
  • 95% for 30 min
  • 100% for 30 min
  • 100% for 45 min
  • 100% for 60 min (store overnight in 100% ethanol if necessary)
  1. Gradually infiltrate with LR White resin (Obtained from London Resin Company, P.O. Box 34, Basingstoke, Hampshire RG21 2NW, United Kingdom, or from MARIVAC LTD., Halifax, N.S., Canada). We use the Medium grade.
  2. The ratio of 100% ethanol to LR White resin:
  • 2: 1 for 30 min
  • 1: 1 for 30 min
  • 1: 2 for 30 min
  • 100% LR White for 60 min
  • 100% LR White for 60 min
  • 100% LR White overnight.
  1. Embedding in gelatin capsules:

This is a common method for LR White resin, as LR White polymerizes with heat in the absence of air. Capsules of 6.5 mm diameter (#1) are convenient.

We usually make a holder for the capsules out of small, light cardboard boxes with holes the diameter of the capsule poked in the top side with a pencil or similar instrument.

Remove the tops from the capsules and place the bottom half in the holes.Place fresh LW White resin in the capsules. Transfer one specimen into each capsule with a Pasteur pipette and then top off the capsule with fresh resin.

It may be easier to pour all the contents (e.g. root tips) of the vial into a small disposable dish and carefully transfer each specimen with a toothpick.

Replace the capsule lid tightly, remember that LR White polymerizes in the absence of oxygen, so that as little air as possible remains in the top of the capsule.

When the samples are deposited in the capsules, indicate on the box which specimens you have, and then place them in the oven at 60°C.

Samples usually polymerize overnight. This method works consistently, but the samples may end up in a twisted configuration at the bottom of the capsule. These need to be cut out and re-mounted on a new resin block in the desired orientation.

  1. Flat embedding with LR White:

When it is necessary to embed specimens on a flat surface (as in the case of long thin roots found in most AMs) then the gelatin capsule method is unreliable. In this case transfer the resin infiltrated samples into an aluminum weighing dish and cover them with at least 2 mm of fresh resin.

Place another aluminum weighing dish on top of the resin so that a seal is formed.(Mycorrhizal Roots)

Mark the specimens with a small strip of paper with a code written in pencil placed in the resin. Separate the specimens in the weighing dish with a toothpick so that they don’t clump together during polymerization. Place dishes in a 60°C oven and polymerize for 3–12 h.

Make sure the weighing dish or embedding molds are lying flat in the oven, or the samples will clump together at the lowest point.

  1. Flat embedding with LR White polymerized with UV light: LR White can be polymerized with UV light; the protocol for this is the same as in 8 but the weighing dishes are placed under a UV light source.
  2. When using the flat embedding methods, specimens are cut out of the resin using a small saw and mounted on resin stubs to fit the chuck of the microtome. (Mycorrhizal Roots)

Staining resin-embedded tissues for light microscopy

Samples embedded in either LR White or Spurr’s resin can be stained for light microscopy with a number of methods (see Ruzin 1999). We routinely use the following:

  1. Prepare a solution of Toluidine Blue O (TBO) by adding 0.05 g of TBO and 1.0 g sodium borate to 100 mL d H2O. Filter and store in a reagent bottle.
  2. After sections have been heat-fixed to a slide, add TBO to cover sections and heat gently.
  3. Rinse the stain from the slide and let dry.
  4. Permanent mounts can be prepared by adding a drop of immersion oil and a cover glass and then sealing the edges of the cover glass with nail polish(Mycorrhizal Roots).

Staining hand sections for light microscopy

A considerable amount of information can be obtained concerning root and mycorrhiza structure using fresh material. A detailed description of how to prepare hand sections and some of the stains that are useful can be found in Brundrett et al. (1994, 1996).

Share on:

Leave a Comment