---

I am a long time amateur microscopist, having recently begun mounting various specimens of diptera, I am not able to find any standards concerning positions or orientations which are desirable. Are there accepted standards for slide mounting (as their are for pinning) or is it more or less reliant on the intended use to which the slide will be put? Where may I locate information on accepted practice which is not taxa specific?

---

Have a go at the method described in this publication: http://people.ds....01983a.pdf.

---

I'm not familiar with the use of that mounting medium as described. In the past I have used the variation described by Doestschman 1944 (Trans. Amer. Micr. Soc., 63:175) for use in deep cell mounts prepared without pressure. The individual mounting of dissected portions described is just what I was looking for. I am surprised that more emphasis is not placed on individual dissection of mouth parts to facilitate identification, but such is the plight of the amateur.

Thank you for pointing me to that excellent publication.

---

I sincerely hope that Berlese fluid is not widely used anymore. I use euparal in my slides, most of my collaborators tend to use canada balsam.
Which parts are dissected and which aren't seems to be taxon-specific. When I make Psychodid preparations I first mount the wings and set the rest of the body to maceration -- ca 24 hours in KOH at room temperature. Then I clean the body in three subsequent baths of acetic acid, 70% and 100% ethanol (ca 10 minutes each) before dissecting off the terminalia and head. The head, wings, thorax and terminalia are mounted under separate coverslips. The slides take several months to dry, but can of course be examined to some degree before they are finished.
People who do Chironomids and Ceratopogonids also usually dissect the legs and antennae from the rest of the animal.

---

Surely one doesn't always require a permanent mount (the primary argument against Berlese as I see it)? I'm rather surprised to hear that Balsam is used by contemporary professionals. I would have expected various of the proprietary synthetic resins to prove more popular; the expense being less of an obstacle for non-amateurs.

If I may, is there a particular reason you elect to macerate at room temperature? I've found a certain degree of difficulty in obtaining satisfactory consistency when using a hydroxide macerating solution over a period of more than a few hours, too often finding species too weakly or heavily macerated. I nearly always employ an incubator to maintain a temperature of 50-70 degrees to speed the process so that I may actively monitor the results.

I take it the portions likely to be dissected vary depending on the features unique to species?

---

As a taxonomist, I do not make temporary slides. Any identification I publish or deposit in a public collection must be available for later corroboration or revision by other specialists. Otherwise, what I do would not be repeatable, and it would be less scientific.
I also do this for the species that can be identified in alcohol, to improve my understanding of their morphology and to ensure uniformity in the collections I work with. It is much harder to manage a collection of some alcohol specimens and some slides than it is to manage a collection where all specimens are stored in the same way.

Heated maceration regularly destroys important taxonomic characters of the antennae (in Psychodids, that is). Unheated maceration takes longer, but if you regularly check on the state of the specimens you will be able to avoid both overclearing and underclearing.

---

Although the mounts I produce are for my own enjoyment, I recognize that permanence and uniformity is certainly desirable from a scientific and curatorial position. The more I think of it the more I feel inclined to break from my habit of producing semi-permanent mounts in aqueous media, though I will sorely lament the added time slides must be left unexamined as they cure.

It honestly hadn't occurred to me that I had been destroying important structures by speeding the process along, no doubt because I am sufficiently ignorant to fail to notice the absence of the destroyed features!

If I might impose on your expertise one last time; I general employ a 10% solution of NaOH. Is potassium preferable, and is 10% simply too strong do you think?

In any case, thank you, sincerely, for taking the time.

---

I have always used 10% KOH, and have generally been happy with the results. Probably other alkalic solutions of about the same pH would work just as well; I can't say as I haven't tried clearing with anything but KOH.

---

Just to see, I made up a quick whole mount (with pressure) of an Anthomyiid using KOH at room temperature for 24 hours. I kept the solution in a dark cupboard as I had heard KOH is somewhat sensitive to light exposure. I suppose I'm pleased with the results even if I didn't bother cleaning it properly of debris prior to mounting, and it hasn't cured yet.

I was most struck by the fact that none of the flies I had processed under heat retained the level of detail in the ovipositor (I can only guess that is the identity of the long structure projecting from the abdomen?). Apologies for the image quality, I pressed into service a 48mm objective that is sorely in need of cleaning so that I could provide as large a field of view as possible.

Slow connection warning; the images are quite large.

Overview: http://imageshack...qJUTNt.jpg

Ovipositor: http://imageshack...EtxHuy.jpg