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to coralline page Sectioning
Sectioning may be done on material embeded in paraffin, or epoxy. Woelkerling (1988) provides a good account of preparing specimens for epoxy sectioning. At the University of the Western Cape and the University of the South Pacific, all sectioning is done using a carbon dioxide freezing microtome. Similar microtomes are available based on electronic cooling from a Peltier device, and one such device is in use by Dr. Y.M. Chamberlain at the University of Portsmouth, England. Freeze-sectioning gives quick results, but the sections are invariably too thick to be useful for microphotography. For this reason, camera lucida drawings of critical structures are generally made for publication.
Decalcified pieces of material for sectioning are placed in 70-90% ethanol for at least an hour to harden the material and make it manageable. Each piece is removed from alcohol and blotted for 2-3 minutes on paper towel. Thicker material may be soaked in Hamilton’s freezing solution (see below) in a watch glass for a few minutes prior to sectioning. Thin, flexible specimens can be made into artificial epiphytes to facillitate sectioning by gluing them to a piece of Sargassum, seagrass or even a bit of terrestrial plant leaf using a tiny amount of Hamilton’s freezing solution. Sections are frozen to the microtome stage using Hamilton’s freezing solution. Care must be taken not to freeze the specimen so much that it becomes hard and brittle when using carbon dioxide to freeze the specimen. This will require a bit of practice.
Freezing medium
Freeze sectioning requires a mounting medium to prevent the material from freezing in a form that is too brittle to section. Hamilton’s freezing solution is based on sucrose and gum Arabec, which allows some elasticity if it is not frozen to too low a temperature. Hamilton’s freezing solution is made as follows: 100 ml distilled water 1 g gum Arabec 30 g sucrose 1-2 crystals of thymol dissolved in minimal 100% ethanol (use only enough to dissolve the thymol, use a glass beaker as thymol dissolves many plastics as well as the paint on lab benches as I discovered the hard way!) The stock solution should be kept in a refrigerator.
Staining and mounting medium
I use almost exclusively a 50% solution of Karo or other corn syrup for preparing slides. Prepare a dropper bottle of it by making a 50% solution, and mixing with a thymol crystal dissolved in c. 1 ml ethanol (use the minimal amount of ethanol needed to dissolve the thymol, and use a glass beaker as thymol dissolves many plastics). Once slides are prepared they can be left to air dry (be careful of sugar ants, they can dry up a slide in minutes!), or dried on a slide tray or in a warming oven. To obtain well hardened mounts fairly quickly, have the slide tray operating on a 12 h on, 12 h off cycle. This allows some of the moisture under the coverslip to moisten the harder material at the edge of the coverslip when the warmer is switched off, and this evaporates away when the warmer is switched on. This gradually draws away excess moisture from under the cover slip, and hardens the medium over a period of 1-2 weeks.
Aniline blue is the stain which gives the quickest results. Make up a densely coloured, but not saturated solution of water soluble aniline blue, and keep it in a dropper bottle. Add one drop to a microscope slide, then add 5-7 drops of 50% Karo syrup and mix thoroughly using a small paint brush before sections are transferred to the slide. Transfer sections from the microtome blade using a fine sable hair brush, and orient them gently under a dissection microscope before adding a coverslip.
Another stain that is very useful, and that shows up cell connections very well, is 4% potassium permanganate. It works best if the specimen can be soaked for 1-5 hours in a watch glass after it has been decalcified but before sectioning. Then add one drop to the slide an prepare as noted for aniline blue above. This stain can also be used for paraffin or epoxy sectioning.
Derek Keats,
updated 03/09/01