Preparing coralline algae for
transmission electron microscopy

Specimen preparation for TEM seeks to preserve cell structure with minimal alteration. Some common pitfalls and how to avoid them are described here. Preservation of membrane lipids is critical; without lipids, membranes are poorly defined and images have little contrast. Loss of lipids increases with temperature, and organic solvents used in dehydration cause lipid extraction. Therefore, specimens should be kept cool and must be fixed with osmium tetroxide, a dangerous lipid preservative and stain. Another concern is mechanical damage. Collecting and subdividing corallines may require force. The shock of a chisel blow may be transmitted widely through the thallus, shattering walls and cells. Drying causes severe damage; specimens must be kept continuously moist before and during processing. Decalcification of corallines by EDTA is a slow process, chelating the calcium to slowly dissolve the carbonate and release CO2. Decalcified thalli are easily damaged. Thalli should be sliced, not chopped. Decalcification progresses deeper into the specimen with time. If your interest is only the upper surface, the decalcified portion can be gently sliced off. Chemicals used in electron microscopy pose immediate and long-term hazards to users. Osmium tetroxide vapors can cause temporary blindness; formaldehyde and some embedding resins are possible carcinogens. Some buffers and most stains contain dangerous heavy metals. Do not expose unsuspecting people to dangerous chemicals, or ask others to handle dangerous chemicals without proper instruction and protection. Transporting hazardous chemicals in luggage or shipping through the mail without proper packaging is dangerous and illegal.

Recommended primary fixative: 3-5% glutaraldehyde, 0.2 M sucrose in 0.1 M phosphate buffer (pH 7.0). Phosphate buffer is prepared from stock solutions of 0.1 M NaH2PO4 and 0.1 M Na2HPO4 . These are added together in differing proportions to obtain the pH desired. One part dihydrogen stock and two parts disodium stock will give a 0.1 M buffer of pH 7.0; equal volumes of stock solutions produce a buffer of pH 6.75. Sucrose in the fixative solution raises the osmolarity. In a pinch, seawater can be used as a buffer. Specimens should be fixed for 2 hours to overnight and should be protected from heat and sunlight. Use gloves when handling fixatives and avoid the fumes. Phosphate buffers are physiological buffers, therefore fungi and bacteria can attack specimens stored in pure buffer. Cacodylate buffer does not have this problem, but it contains arsenic and must be handled with gloves and in a fume hood, and it must be disposed of as hazardous waste. For these reasons, it is not appropriate to use in the field.

After primary fixation, the fixative solution is removed and replaced with pure 0.1 M phosphate buffer. After a brief (e.g. 15 minutes) rinse, transfer to buffer plus 5% disodium EDTA (or to saturation). When the rate of bubble generation slows, the solution should be replaced. Some specimens still may be partially calcified after a week of decalcification. Fortunately, the presence of EDTA prevents growth of organisms. When decalcification is complete, rinse in two changes of pure buffer.

Osmium tetroxide fixation must be done in a fume hood; bottles and vials should be tightly capped. Use 1% or 2% OsO4 in buffer or distilled water for 2 hours to overnight. Specimens and other organics, including skin, are blackened by contact with OsO4. Waste must be properly disposed of. Do not attempt to use OsO4 without advice from an experienced user. At the completion of postfixation, the OsO4 is pipetted away and replaced with cold distilled water. After two water rinses, the specimens are dehydrated by changes of increasing concentrations of acetone or alcohol (ethanol or methanol). Usual steps are 10 minutes each in 25%, 50%, 75%, 95%, and three changes in 100% dehydrant that has been dried over beads of hygroscopic molecular sieve (e.g. type 4A, 8-12 mesh). Infiltration with increasing concentrations of resin (in acetone or alcohol) follows dehydration. Use 25%, 50%, 75%, and 100% resin in steps for a minimum of 2 hours each. Transfer the specimens to pure resin in silicone molds, which allow for easy orientation. Slightly overfill the mold to allow for resin contraction during polymerization. Polymerize the resin in a stable oven at a temperature appropriate for the resin used, usually 60o or 70o C. I prefer Spurr's resin, because its low viscosity allows infiltration of specimens other resins may not. However, decalcified corallines are relatively easy to infiltrate, and Epon and Epon-Araldite are less hazardous. Waste resin and resin on consumable items, such as vials and pipettes, should be polymerized before disposal.

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Derek Keats,
updated 02/01/01