Following on Terri Taylor's (1993) report on drying methods, a further method, utilizing acetone as the effective agent, needs to be publicized. The acetone drying procedure is a simple and cost-effective method for drying weakly sclerotised or small insects preserved in alcohol in preparation for dry mounting or scanning electron microscopy. The principle of the technique relies on the replacement of the alcohol contained in the specimen with highly volatile acetone, which is then speed evaporated under heat, leaving the specimen in a perfectly preserved state.

The technique was first described by Truman (1968) who used the method to preserve larval and adult mosquitoes. Walpole et al. (1988) applied the technique to representatives of Diptera, Hemiptera and Anoplura for SEM preparation. Tony Ware of the now disbanded Rhodes University fig team, headed by Steve Compton, was the first to apply this technique to Hymenoptera, specifically fig wasps, as an alternative to critical point drying for SEM preparation. Together with Rob Cross of the Electron Microscopy Unit at Rhodes University, they reported on the success of this treatment for the preparation of Agaonidae (Ware & Cross, 1989).

I have been using the technique for the mounting of fig wasps, and chalcids in general, for several years now. This includes both specimens preserved in alcohol (under sub-optimal conditions) for twenty plus years and freshly collected material from Malaise traps, yellow pan traps etc. The method works equally well for old and freshly collected material and is particularly useful for families with weakly sclerotised or small representatives. More recently I have subjected proctos (s.l.), cynipoids, ichneumonoids and smaller sized aculeates, such as dryinids to this treatment, with equal success. A colleague, Hamish Robertson, applies the technique successfully to ants. Many chalcids and proctos, being strongly sclerotised, do not really require this special treatment. I apply it as a matter of course, in an attempt to preclude any possibility of antennal or compound eye collapse. In some taxa, particularly ichneumonids, the gaster (metasoma) often distends when stored in alcohol. Acetone drying preserves the distention, creating an artificial appearance to the gaster, but this does allow for clear all round observation of the tergal and sternal plates. It is, nevertheless, possible to control the final extent of distention by allowing the specimen to dry out until the gaster returns to normal, before placement in the acetone environment.

The acetone environment can be created by saturating a layer of cotton wool in the bottom of an airtight glass (or acetone-proof plastic) container with acetone and placing or pinning the specimens on some sort of platform (I use a piece of SPX foam) above the cotton wool. Specimens may either be placed directly into the acetone environment from alcohol or first mounted. For direct placement it is best to float the specimens out of the alcohol onto thin card, as it is essential to achieve the desired position for later mounting at this stage. A certain amount of manipulation can be carried out after acetone saturation and before drying, but the final position has more or less been determined at this point. Alternatively the specimen can be micro-pinned and double mounted (the synthetic "polyporous" silicon strip is not affected by acetone) or card mounted prior to placement in acetone vapour. I have used both water soluble glue and shellac gel (with little success) for card mounting. The adhesive must be allowed to dry sufficiently before placement in the acetone environment, so that the properties of the glue are not affected. Water soluble glue turns opaque and shellac gel looses its adherence properties. The latter is presumably as a result of the alcohol in the gel being replaced by the acetone and affecting the resultant bond. It is absolutely critical to achieve a balance between the glue drying sufficiently and the specimen remaining sufficiently wet to prevent any collapsing prior to acetone treatment. This sounds as if it involves much fussing, but with a bit of experimentation it is not difficult to get the timing right. I find that the ease of handling mounted specimens out weighs the disadvantages involved with gluing the specimen first.

The specimens are left for a minimum of three hours in the acetone jar, although no ill effects arise if they are left overnight, before removal and placement under a desk lamp (close to the bulb) for at least half an hour. A 60-watt bulb provides sufficient heat to speed dry the volatile acetone, leaving the specimens in a well preserved and uncollapsed state. With this method, good results can be achieved quickly and at low cost without requiring access to freeze drying or critical point drying equipment.

Taylor, T.l. 1993. Chalcid Forum, 16: 10

Truman, J.W. 1968. Annals of the Entomological Society of America 61: 779-780

Walpole, D.e., Coetzee, M. & Lalkhan, C.M. 1988. Journal of the Entomological Society of

Southern Africa 51: 293-296

Ware, A.b. & Cross, R.H.M. 1989. Proceedings of the Electron Microscopy Society of

Southern Africa. 19: 39-40

I am afraid that I am not well plugged into the network of chalcid workers and the methods I write about here may be old hat to most. The distortion of dried chalcid specimens is of course a well known problem, especially for specimens collected into alcohol. From what I have learned in the literature, the solution to this is critical point drying, which requires fairly expensive equipment and, I am informed by some, can be dangerous. For years, our electron microscope technician has been preparing small arthropods by aldehyde fixation, dehydration to 100% alcohol, and then through hexamethyldisalizane (HMDS). The HMDS is allowed to evaporate leaving specimens that show no collapse.

I have adapted this method to specimens that I collected into 95% ethyl alcohol with a Malaise trap. The results, I think, have been good and work well not only for chalcids but any soft bodied insects including braconids, Diptera, aphids etc. I remove the desired specimens from the trap jar, rinse in clean 95% alcohol, transfer them to 100% alcohol, change this once, and then transfer them through two baths of HMDS. I then allow the HMDS to evaporate and this leaves the specimens ready for mounting on cards or points. The specimens are of course brittle and must be handled with care.

HMDS is nasty stuff and must be handled with great caution: no contact with the skin, and no breathing fumes. Always work under a hood. This chemical is not exorbitantly priced and only small amounts are needed for a sample of several dozen insects. HMDS is available from Polysciences, Inc. Warrington, PA 18976 (1-800-523-2575), Catalog # 00692.

Previously, I wrote about the use of Peldri II as a chemical alternative to critical-point-drying (CPD) (Brown, 1990). Using Peldri II for specimen preparation has the advantage of not requiring expensive equipment and huge canisters of CO2, but the chemical is somewhat expensive and requires some equipment, i.e. a hot plate.

Recently I came across a material that seems to overcome all the drawbacks of Peldri II. This chemical, called hexamethyldisilazane (HMDS) is readily available and cheap, costing $18.00/250g, versus $58.00/250g for Peldri II. No heating or cooling is needed for using HMDS: one merely dehydrates specimens to 100% alcohol, then do two soaks of 1/2 hour in pure HMDS (i.e. change the HMDS once). I use small vials for the soaks. After the second soaking, I pour the HMDS and flies into shallow depressions or small dishes, and allow the liquid to evaporate under a fume hood. Specimens come out exactly like CPD prepared specimens, ready for SEM or for general mounting for the collection. Other users agree, finding that HMDS is just as or more effective than CPD for producing perfect specimens of various tissues (Adams et al., 1987; Nation, 1983). Note for larger samples (for instance, 50 small flies at once), I recommend an extra change of HMDS.

Adams, J.L., Battjes, C.J., & Buthala, D.A. 1987. Biological specimen preparation for SEM by a

method other than critical point drying. Proceedings of the 45th Annual Meeting of the Electron Microscopy Society of America (ed. by G.W. Bailey). San Francisco Press, San Francisco.

Brown, B.V. 1990. Using Peldri II as an alternative to critical point drying for small flies.

Fly Times, 4, 6.

Nation, J.L. l983. A new method using hexamethyldisilazane for preparation of soft insect tissues

for scanning electron microscopy. Stain Technology. 58: 347-351.