Laboratory of Ann Hirsch

Sweetclover Methods

Techniques for light microscopy

A - Plastic sections*

1. Fixation
   1. Tips: Work with a small sample so that the tissue can be adequately fixed and the embedding material can easily penetrate. The size of a particular sample has to be determined by trial and error, but generally we do not use material that exceeds 2 or 3 mm in length if the tissue is thick. Tissue is generally fixed with aldehyde-based fixatives which cross-link many cellular components, especially proteins. Lipids generally require additional fixation and in some cases, the tissue can be post-fixed with osmium tetroxide. The other important factor is pH. Most fixatives and buffers have a pH between 6.8 and 7.1.
      
   2. Fix tissue in 3.5% (w/v) glutaraldehyde, 1.5% (w/v) paraformaldehyde in 0.1 M cacodylate buffer, pH 7.0. Note: 4% glutaraldehyde in 0.1 M phosphate buffer (pH 6.8) works well also. Fix for a minimum of 4 hours, but do not overfix. An overnight fixation is usually fine. We usually fix at 0oC but it is okay to fix at room temperature.
   3. Rinse twice in 0.1 M cacodylate (or phosphate buffer). 15-30 minutes per rinse.
   4. Post-fix in aqueous 1% (w/v) OsO4 at 0oC for 4 to 6 hours. Do not do this step if you are planning to embed in LR White resin.
   5. After removing the osmium tetroxide, wash the tissue with distilled water twice, 15-30 minutes each rinse depending on the size of the tissue. Continue at 0oC.
      
2. Dehydration
   
3. Dehydrate in a graded ethanol or acetone series, e.g., 10%, 30%, 50%, 70%, 95%, and three 100% steps.
   
4. Tips. Whether you use ethanol or acetone usually depends on the type of resin that will be used for embedment. The important thing is to dehydrate slowly and gradually so that the water is replaced with the solvent in which the embedding medium is soluble. The timing can be as short as 15 minutes per change or as long as 1 hour depending on the size of the tissue. The temperature for dehydration is also variable; you can dehydrate at 0oC or at RT.
   
5. Embedment
   1. Before embedding, rinse the tissue three times with the most concentrated (100%) ethanol or acetone.
   2. From 100% acetone: embedding into Spurr standard low viscosity medium (Polysciences Inc. Warrington, PA).
      1. Make fresh Spurr resin. Dilute some to 25% and also to 50% with acetone.
      2. Transfer tissue from 100% acetone to 25% Spurr resin. Let infiltrate at RT for 4 hours.
      3. Transfer to 50% Spurr resin for another 4 hours.
      4. Make 3 changes into 100% Spurr resin; let one infiltrate overnight.
      5. Change to fresh 100% Spurr resin and then embed into BEEM capsules, molds or aluminum weighing dishes.
   3. From 100% ethanol (you can do the entire dehydration series at RT), embedding into LR white (Polysciences Inc. Warrington, PA).
      1. Infiltrate specimens with a graded series of resins. 25% for 30-60 min.
      2. 50% for 30-60 min.
      3. 75% for 30-60 min.
      4. 100% for 30-60 min
      5. 100% for 12 hours to overnight.
      6. 100% for 6 to 8 hours.
      7. Transfer to air-tight containers (e.g., BEEM capsules or flat embedment between two aluminum weighing dishes) and polymerize at 55 to 60oC until hard (usually overnight or the following day). LR White will not polymerize in the presence of oxygen so it is essential to exclude all oxygen.
   4. Tips. There are various resins on the market. We have had good luck with both Spurr and LR White resins. In any case, use these materials in the hood and dispose of used resin and ancillary materials properly. The various resins differ in viscosity which may be an important factor in infiltrating larger specimens. The procedures of infiltration and embedment are similar to the dehydration steps. The idea is to replace the solvent with resin slowly and gradually to avoid cell and tissue collapse.
      
6. Sectioning

1. Trim specimens and on an ultramicrotome, cut sections for light microscope of a thickness of 0.5 to 1 or 2 µm using a glass or diamond knife. Transfer the sections to a drop of filtered water on a microscope slide and allow to dry on a hot plate. You can coat the slides with a gelatin adhesive or with poly-lysine if you are having trouble getting the sections to stick on the slides.
2. Stain the specimens on the slide with 0.05% toluidine blue O in 1% Borax. Put a drop of the stain over the specimen and gently heat for 30 sec or less. Wash off the excess stain with a wash bottle. After the slides are dried, place some Eukitt, Permount or other adhesive on the slide and cover with a cover glass.

*These steps can also be used for electron microscopy, but a diamond knife needs to used for sectioning and the sections must be considerably thinner.