Fixatives - Materials and methods in Practical microtechnique

Fixatives

Alcohol and formaldehyde-based fixatives

Seventy per cent alcohol is typically used as a plant fixative in schools and laboratories, because it has little effect on the user should they accidentally spill it over themselves provided it is immediately rinsed off with water. Al-cohol by itself tends to harden the plant tissues and can cause changes in shape. It is to be avoided as a histological fixative, as no cell detail will be preserved at all.

Formaldehyde has to be treated with care and a great deal of respect. Do not breathe the vapour, and use a fume cupboard. Plant tissues require quite aggressive fixation techniques, and there is no reason why students should not use more potent liquids for fixing plants if suitable care is taken. For general histological purposes, the formalin–acetic acid–alcohol (FAA) mixture given in Table 10.1 works reasonably well. Always use a fume cupboard.

Note: FAA is a corrosive liquid, and if it comes into contact with the skin it should be washed off immediately. The best preventive measure is towear laboratory gloves. It is well worth the trouble and care to use material fixed in FAA because materials that are preserved in it section well and can be kept in the reagent indefinitely. However, be careful to store vials and bottles containing FAA in a well-ventilated space, as the fumes are harmful and should not be inhaled.

Irrespective of the fixation technique used, plant material to be fixed is normally cut into portions to enable rapid penetration of the fixative. Care should be taken to ensure that the portions of plant tissue are cut so that they can be readily identified and oriented. Bottles with wide mouths and polypropylene screw or push-on tops are ideal for storage, and can be ob-tained in a range of sizes. It is best to keep the plant in fixative for at least 72 hours before continuing on with the preparation process. Plant material may be kept in FAA and can be stored for as long as required, but the bottles should be inspected regularly for evaporation and topped up with 70% alcohol if necessary. This is the most volatile of the constituents.

Specimens to be sectioned are removed with forceps and washed in run-ning tap water for 30 minuets to 1 hour. They can then be handled safely.

Non-coagulating fixatives

There are other fixation options available for detailed anatomical and cyto-logical studies, which will preserve cytoplasmic details as well as prevent se-rious plasmolysis which is usually evident when using FAA and its associated dehydration procedures. Feder and O’Brien (1968) reviewed principles and methods used in Plant Microtechnique, which introduced the concept of using non-coagulant fixatives such as osmium tetroxide, acrolein glutaral-dehyde and formaldehyde, coupled with the use of plastics such as glycol methacrylate polymer instead of wax. The advantages of using polymers, is that thin sections (1–3 µm) may be made, and these sections show excellent cell structural detail. Osmium tetroxide is particularly dangerous, and should be used only by competent, trained people in a fume hood, following all the safety precautions.

We recommend acrolein as an alternative fixative, especially where the researcher is intent upon preserving (and needs to resolve) cellular details which would not be preserved at all with FAA. Ten per cent acrolein is rou-tinely used as a biological fixative. It may be made up in tap water or added to a suitable buffer solution, such as a phosphate buffer, not unlike that used in electron microscopy preparative techniques. Dehydration is a little more complicated than with FAA, but well within the scope of the average anato-my laboratory. Caution: Wear protective gloves and use a fume hoodwhen working with acrolein.