Has anyone successfully removed insects from amber? If so - how?
I've had some success grinding and polishing to get insects close to the surface for photography (first pic), but when the piece is irregular or the insect is not positioned well to allow this - I struggle to get decent images (second pic).
I've read that chloroform will dissolve amber and leave insects intact almost as if "freshly killed", so I have a litre on order to try it. Assuming it works, how should the extracted insect be treated? Should I just let it dry or should it go through other stages (e.g. alcohols) to remove chloroform?
Any advice and/or other's experiences gratefully received. Cheers
That's a very interesting project. Maybe you should keep the insect always under a liquid. If you remove a very fragile object from a liquid, the surface tension of the liquid can tear off bits as you lift it through the surface layer.
It may vary from case to case and be different for amber from different ages and localities. At least in most of the cases I have seen, the exoskeleton of the insect is gone and what is left is an external mold of its surface (in practice, a hole in the amber replicating the surface of the insect). Even when the exoskeleton seems preserved (by looking through the amber), it may be just a thin film of opaque organic material deposited on the walls of the mold, rather than a truly preserved exoskeleton. Often, the amber is sectioned to expose the holes and metal-sputtered to observe the details of the mold surface by SEM. I am not sure of the details, but I think in some cases one vacuum-fills the holes with a material that hardens, and then the amber is etched away and the artificial filling observed by SEM.
My guess would be that Quaternary or subfossil amber (and copal, which is not fossilized), i.e. very recent materials on a geological scale, are the best bet to extract parts of the insect exoskeleton.
I have seen for instance a collection of fossil arthropods in Baltic amber in the museum of paleontology of Uppsala University, Sweden. The amber pieces have been cut and polished to provide approximately flat surfaces for optical microscopy of the specimens.
In Baltic amber, practically no organic material of the insects is left. There is just a very thin lining of the otherwise empty cavity where the insect once was.
I am with the others in that in many cases the insect is just a carbon ghost, and a finely detailed mold in the tree resin.
But since they have been extracted, then not all of them are 'gone'. I would favor trying this on recent amber (Baltic), and on a tougher object first (beetles, bits of bark).
My guess would be to use chloroform to dissolve the amber, and a series of chloroform washes to gradually remove the dissolved amber. From there, to get the specimen into the air you would need to negotiate things so the chloroform can be removed without surface tension crushing any of the softer stuff. This seems a similar problem to how soft tissues are totally dehydrated for scanning electron microscopy. The procedure there is to use what is called a critical point dryer, which gradually exchanges the solvent with liquid CO2, and then letting it sit in the CO2 bath in a pressure chamber. There, the CO2 is gradually sublimated to gas, without surface tension. Even really delicate tissues survive well after that.
So where to get a critical point dryer? The easiest answer is to inquire at your local universities. Biology departments will generally have a scanning em facility and they need this machine to do that sort of work. I expect one could find a biologist (preferably an entomologist) who would be interested in helping you, as this is a pretty cool looking project, possibly leading to a scientific publication.
Thanks for the responses everyone. By adding "-jurassic -park" to my google query, I was able to find much more published material on the subject. It does irk me that "interesting stuff" like this gets completely swamped once it becomes fictionalised (aka trivialised). Grrr! Anyway, your comments confirm my findings. Survival of a specimen is sample/age dependent with amber from Lebanon have the greatest chance of success. Others (most) just leave "ghost" cast holes in the amber. The type of resin involved is possibly the most important factor...
Unfortunately, my desire to experiment anyway is stymied. Chloroform is on the "restricted list" so I've been informed that I can't get it from my usual supplier (who has gone super-stringent on applying the rules of late). My order was refunded. Sigh. I'll just have to get better at grinding and polishing and resign myself to sacrificing parts of specimens to get their more interesting parts closer to the surface.
Thanks again for the detailed responses. This really is an extremely well-informed forum. Cheers.
As to chloroform restrictions, once again a biologist collaborator can help you on that. They can get it pretty easy. Those who work on fruit flies can have chloroform as it is a common back-up method for anaesthesia of the flies.
I wonder if there are alternatives. One could experiment with cheap amber, doesn't need to have anything in it. Like fingernail polish remover, or mineral spirits, etc. -- stuff you can get at a hardware store.
The best way to get a really good surface on amber is to use aluminium oxide metal polish. After I have the surface looking good with 1500 grit paper, I use 1 micron and then 0.05 micron oxide. Both should be used on very soft, short nape felt. You can get this from the same sources as the polish. I bought a small supply of 2 inch pads. They are sticky backed. The easiest way that I could find to keep them flat in use was to stick them to an old CD.
Any remaining imperfections can be overcome by immersing the amber in oil. Amber has a refractive index of about 1.53. This is only slightly less than immersion oil. However, immersion oil is quite expensive in the sort of amounts which you need to submerge pieces of amber. Johnson's Baby Oil has a refractive index of about 1.53, so is a perfect match for the amber. Citrus scented is pleasant to have open on the bench for prolonged periods.
I use an old Fujifilm 35 mm Provia cassette which I have cut down to a little over 10 mm high. You can cut disks of various different colours of paper or card and drop these in below the oil and specimen to see which gives the best contrast. Blue is often good, as are white and occasionally black. The film cassette acts as a very nice diffuser for your lights.
If You can get xylene, you can give it a try. It is sold under different brands, Xylol being one. Each brand is a bit different combination of the xylene isomers.
The oldtimers used it when working with canada balsam, the original one from conifers.
Like almost every chemical substance, it is nowadays found toxic and also slightly mutagenic. Also very flammable, caution is needed when used.
It's really dangerous but you already know it.
