Edited and expanded: November 2025.

I mount all my aphids in Canada balsam, which is made from the resin of the balsam fir tree.  Following is a brief description of the procedures, tools, and fluids I use.

Before getting to slide making, I should mention how I preserve my specimens.  I learned early on that aphids preserved in ethanol of 70-95% concentration quickly (i.e. within a month) become brittle and difficult to mount without serious dismemberment problems.  Consequently, I adopted a collecting solution that was suggested by my acarology professor, Gerald Krantz.  He presented a recipe in his book (A Manual of Acarology, 1978) and called it Oudeman’s fluid: Glycerine — 5 parts, 70% ethanol — 87 parts, glacial acetic acid — 8 parts.  This solution not only keeps fresh material pliable for years, it also can soak and render flexible previously dried or damaged specimens.  Best mounting results with aphids preserved in this fluid, following my methods below, are achieved when the specimens are soaked for a month or more.  Less than a month and NaOH clearing is less effective for some reason.

Equipment and Supplies

1. 9.1% NaOH (sodium hydroxide): I mix this using 100 grams of distilled water (tap water will work, but the minerals precipitate and cause the solution to become ineffective sooner) and 10 grams of NaOH crystals. I weigh this using a laboratory balance I found at a thrift store. The NaOH is sold as “lye” in the drain cleaner section of hardware stores. Be sure to get pure lye crystals, not any sort of prepared drain cleaner. Be careful with NaOH, as it is extremely caustic and dangerous. As it goes into solution it will generate quite a lot of heat, so I let it cool to room temperature before putting the lid on to avoid having the lid get sealed on with suction. Store the NaOH crystals and the 9.1% solution in the fridge.
2. 50, 70, and 95% ethanol: these solutions are easy to make assuming you remember a little bit of your high school chemistry and can get ahold of 95% ethanol. The latter can be a pain — for many years I bummed ethanol off my colleagues employed at university research labs. Then I bought my own from Bioquip, the now defunct entomology supply store. My last batch was secured from Carolina Biological Supply; as a non-institutional customer I was allowed to purchase it from them but only 2 liters at a time (plenty for several years of collecting and mounting). Note that most of the ethanols sold on Amazon seem to be impure or are not even ethanol. I imagine some of you wonder whether denatured ethanol will work. As far as I can tell, it is fine. Another likely question is whether isopropyl alcohol will work. My honest answer is, “I don’t know.” I’m very reluctant to change something that works, so I have never tried isopropanol. I store my ethanols in small plastic squirt bottles one can get at a craft store.
3. Clove oil: I use pure clove oil sold by the company called NOW Foods. I gather that one can find various other things sold as clove oil that are apparently mixtures with other things meant to smell a certain way for perfumes as such. So, buyer beware: only 100% clove oil for you. The same clove oil can be used for preparation of many specimens.  I replace the clove oil in my contact lens holders (see below) after about 150 slides.
4. Orange oil: I also get this oil from NOW Foods. It must 100% pure orange oil, no mixtures.
5. Canada balsam: I buy Canada balsam from Woodfinishing Enterprises in Wauwatosa, WI. It costs about 10% as much from him as it does from Fisher, VWR, and similar, and his works just as well. Thin your balsam using orange oil. I have been asked whether clove oil can be used to thin the balsam since it is used in the last clearing step. Why not simplify the supply list? My answer was, “I don’t know.” Given that I know balsam thinned with orange oil will be good for a few decades (and therefore probably indefinitely), and I don’t know anything about the fate of balsam thinned with clove oil, I am reluctant to change. I thin my balsam more than I gather others may — I want it quite fluid, perhaps a bit thinner than molasses (treacle). I store my thinned balsam in the glass container shown in the photo below. This is one item that might be hard to find outside a laboratory supply company — I’ve had mine since graduate school. A key feature is a lid that prevents evaporation and does not easily get glued shut by the sticky balsam.
6. A dish for processing samples from collection vial to clearing solution. See the photo for the 2-chambered dish I got at a thrift store many years ago. This is very handy because I clear 2 vials of aphids at a time.
7. Vials or test tubes for soaking specimens in NaOH. I use large vials because they are flat-bottomed and therefore stand on their own in my soup-warming cup.
8. A heating device, in my case a cheap soup-warmer and its associated ceramic cup. I think it is arbitrary what is used here. I have not experimented with different devices since the one I have works fine. In grad school I had an actual research device called a dry bath. I think I set it to 80 or 90°C. Of course the temperature one uses affects the lengths of time needed for clearing and the amount of time that results in excessive bleaching during the NaOH steps.
9. Eye droppers for moving fluids. These can be found at drug stores and similar.
10. Micro-tools for manipulating and dissecting specimens. One gotcha that others have found is that the available microtools that one could buy from Bioquip (before they went out of business; see the blue-handled ones in the photos) or similar are too large and clunky for most aphids. The needle tool and the microspatula I use are homemade, gifted to me by my postdoc advisor who had them custom made. They are made from some kind of very durable and flexible metal (I’ve sometimes wondered if bra underwire might work). The spatula is just over 1 mm wide and is pretty much ideal. The spatula from Bioquip is 2 mm wide, and is too big for all but the largest aphids. The needle from Bioquip is similarly too coarse.
11. Forceps. I use these for moving labels or debris in submitted samples.
12. Little dishes. You need little dishes of some kind for the ethanol and clove oil soaking stages. For the ethanol stages I use the mini watch glasses shown in the photo. Watch glasses like this were once ubiquitous in university research labs, but they seem less common in modern times. Getting little ones like I have is very difficult, so I might suggest using contact lens storage thingies — you know, the ones marked with color coding and little nubbins on the lid to indicate left and right. I use such a thing for clove oil because they seal very well with screw-on tops, making it convenient to use the same clove oil for several batches of slides.

Procedures

1. When ready to do some clearing, bring out the NaOH solution and pre-heat your vial(s) of solution indented to clear your specimens in your warm water bath. Puncture the abdomen of each aphid and transfer them to a vial of 9.1% NaOH (plumber’s lye) or KOH. Heat in a water bath using a soup warming hot plate (about 38°C) for about 20 minutes. Don’t soak in warm NaOH too long — it bleaches the pigment in the aphids’ integument.
2. Pour these into a dish and pump out most of the contents of each aphid using a small spatula. To get good clearing it is sometimes necessary to slice the abdomen open just a bit to squeeze out more of the body fluids. It is ideal to leave a couple embryos in the abdomen, fully cleared of course (easier said than done). Heat aphids in fresh NaOH (or KOH) for another 20 minutes.
3. In a dish, finish pumping colored body contents, then transfer clear aphids to 50% ethanol, let stand without heat, covered, for at least 10 minutes. This treatment can often be omitted, leaving only the subsequent 70% and 95% ethanol treatments, however, with some aphids, whiteflies, and other especially delicate arthropods, the appendages shrivel without the 50% ethanol rinse.
4. Transfer to 70% ethanol for 10 minutes at least.
5. Transfer to 95% ethanol for at least 20 minutes, up to several hours.
6. Transfer to unheated clove oil for at least 2 hours.  To do this, I use a microspatula to scoop one or more insects out of the ethanol, then I lightly touch an absorbent napkin to remove most of the excess ethanol from the specimens.
7. Mount in Canada balsam (thinned as needed with orange oil), labeled appropriately, etc. Press the cover slip enough so the legs lie flat, but the head and appendages are not deformed. Balsam can be added around cover slip if necessary to fill in empty spaces.
8. Slides are dried at room temperature (or on the shelf above the wood stove) for 2 or more weeks before being filed in 100-slide boxes.
   

I prepare 10-15 slides per day, representing 4-8 separate samples.  I mount up to 8 or10 specimens per slide, but usually 4 or fewer.

Certain taxa of aphids have biochemistry that interferes with some steps in the process (e.g. precipitation of crystals), but just forge ahead with it best as you can, and in the end the results are usually OK. Clove oil can cure a lot of problems in the last step.

 

Due to some unreliable lab technician work (by me), I’ve had the opportunity to try 9.1% NaOH, 10% NaOH, and something around 14% NaOH.  I have ended up preferring the 9.1% solution.  The ~14% solution was completely un-usable because the internal parts of the aphids were turned into a stiff gelatinous mass rather than dissolving.  With 10%, this same sort of phenomenon sometimes occurred, depending on the aphid taxa being prepared.  At 9.1% I don’t have such trouble; in some taxa the internal fluids tend to solidify in a different way, creating large rounded crystalline masses.  These, fortunately, dissolve in clove oil at the last step.

This description of my slide preparation technique is a tad brief – it is here primarily because a journal editor didn’t want to print it in one of my papers and wondered if I could refer to a website instead. So I did. I dry my slides at room temperature because I’m too cheap and space-limited to use a food dehydrator or similar.