Many of us have had the unpleasant experience of encountering vials, tubes, and jars in our collection of specimens from which most or all of the fluid has evaporated. This may be for a variety of reasons.

1) It may have been sitting on a shelf for years while you were busy with other things. Recently, I found a jar of the algae Nostoc which after 37 years was still in good shape. Next to it were 5 jars of caddis fly larvae, 4 of which were fine and 1 of which had dried up. In this case, it didn’t really matter, because what I was interested in was the tubes they had built and not the larvae themselves. Why were 4 in good condition and 1 not? The clear explanation is the annoying intervention of the impish god Whimsillycus.

2) A frequent factor is the sort of container in which the specimen is stored. After many years, I have discovered a few basics.

a) Vials or tubes with bakelite or plastic screw caps are notoriously unreliable. Over time, it seems that these caps inevitably loosen. There is a marvelous product (and, no I’m not a lobbyist for them) called Parafilm; it is a thin, waxy strip about 1½" wide which stretches nicely and can be used to form a seal around caps of bottles and tubes. However, with some kinds of tubes, vials and bottles even the highly useful Parafilm won’t seal indefinitely. This clearly depends on the container, because I have successfully used it to seal bottles of absolute alcohol, acetone, xylene, etc., not for as long as 30 years, as is the case with some of my specimens, but for 10 years, which for volatile solvents, is excellent. Clearly, the choice of container is critical.

c) Another sort of jar which is frequently used is the clear plastic one with a metal screwcap lid with a coated paper liner. These are problematic, not only in relation to evaporation, but some preservatives, depending on the type of plastic, will attack the container and sometimes produce sufficient scarring that you can’t see what the specimens are. Furthermore, metal caps, lined or not, are, in the long run, almost always a problem. Formaldehyde, even when significantly diluted to be used as a preservative, begins to turn acidic and attacks the metal of the lid and over time, you will find bits of oxidized metal from the lid deposited on your specimens. If that weren’t bad enough, there is also the possibility of deposits of paraformaldehyde, an insoluble, white precipitate. The formaldehyde can be buffered to maintain a neutral pH or one can simply add a piece of marble stone to the solution.

Sometimes we tend to forget that certain alcohols, especially in concentrated form, are very aggressive solvents which is the major reason one should never use alcohol to clean lenses.

d) Some earlier workers used corks for vials, tubes, and small bottles. These, however, need to be sealed with wax and resealed each time after they are opened. Cork is a porous material and solvents, such as alcohol, will evaporate through it rather quickly.

e) Occasionally, I have used small rubber stoppers, but some preservatives cause the part of the stopper within the bottle to swell making it difficult to remove. I have found this especially to be a problem in preservatives containing acetic acid, such as, F.A.A. (Formol-Acetic Acid-Alcohol) which is both a fixative and a preservative used for a wide variety of plant material and some aquatic invertebrates.

As you can see the problem of finding the right sort of container with a lid that seals properly for long periods of time is not a casual matter.

3) A different sort of strategy is to protect your specimens from complete drying even if the basic preservative evaporates. This only works for certain types and sizes of specimens. The usual procedure is to add enough glycerine to the solution to cover the specimen and keep it in fluid even if the preservative evaporates. This has pros and cons but, in general, the advantages outweigh the disadvantages. Glycerine is a very peculiar substance. It is somewhat viscous; it is hydrophillic, that is, it extracts and absorbs water; and at room temperature, there is no significant evaporation. If the preservative starts to evaporate, the glycerine penetrates into the organism or tissue with increasing concentration and keeps the material moist and in generally good condition. Then, why you might ask, don’t we just store everything in pure glycerine? Microscopists in the 19^th and early 20^th Centuries frequently made mounts using pure glycerine or glycerine jelly. If you want to mount specimens in glycerine, you can’t just pop them into the “pure” stuff--if you do, since glycerine is hydrophillic, you will get specimens that are shriveled and distorted. You have to go through a series of stages parallel to those involved in the dehydration process used for mounting in balsam or a synthetic resin. In this case, however, the procedure is much simpler, because you place the specimen in a tube or dish of glycerine diluted with distilled water and then set it aside in a dust-free place and then days or weeks later, at your convenience, you can make a mount.

So, for preservation, and let’s say the specimens are in alcohol, add sufficient glycerine to cover them and then close the container tightly. Now, if the alcohol evaporates, you won’t lose your specimens, although it may take some work if you want to get them back into alcohol or transfer them into water for sectioning or staining. Again, this must be done gradually.

Another disadvantage of using glycerine is that it will gradually attack calcareous structures which are one of my primary interests; so, if you have specimens with delicate calcareous spicules, plates or spines which you wish to study, don’t add glycerine to your preservative.

4) If you have specimens that are rare or of special interest and importance to your investigations, then you may want to take special precautions, especially if they are to be stored for a considerable period of time. This method is only suitable for small or medium-sized specimens; it won’t work for sharks, whales, elephants, or rhinoceros unless you have some REALLY BIG containers. Take your vials, tubes or small jars and make certain that the lids are tight and that the containers are filled to the top with preservative. Seal them with some Parafilm or melted paraffin. Then place the containers in a one gallon wide-mouth jar or something similar and fill the large jar with preservative and make sure that it is tightly closed. Now, if one of the small containers develops a breach, the surrounding fluid will seep in and keep the specimen container full.

5) If you have specimens where all or nearly all of the fluid has evaporated, then you have several options.

a) You can decide that you were never really very interested in studying those specimens anyway and toss the whole mess into the garbage.

b) If the specimens have some calcareous or siliceous spicules, spines, plates or disks that are of interest then, in my view, the best choice is to remove the specimens from any remaining fluid and let them dry thoroughly.

c) If the specimens have dried out completely (and there was no glycerine in the container) and the structures you are interested in are not primarily calcareous or siliceous then, your options are limited and the results are sure to be highly variable and after treatment the material will not be as desirable as it would have been had it not gone through the trauma of dehydration.

I’ll give you an example from my own experience. I have a small jar of Amphioxus which I obtained from a biological supply house. It is a glass jar with a metal lid and a coated heavy cardboard liner. (Hello, out there, biological supply companies. Are you listening? These are not good containers for the long-term storage of specimens! You overcharge us for this stuff and then send it in crumby containers so the fluid will evaporate and we have to order more of your overpriced stuff.) I now have 12 dried Amphioxus stuffed into a very small jar and when they were in fluid, they were, I am sure, quite flexible. Now, however, they are rigid and certainly any attempt to extract them in this condition would result in their breaking apart.

Amphioxus is one of those fascinating borderline organisms, a transitional form between the invertebrates and the vertebrates; it has a dorsal nerve cord, so it is, strictly speaking, a chordate, but there is not a supporting skeletal structure. (In case you’re confused about nerve cord, chordate, and nerve chord, both nerve cord and nerve chord are accepted usage. Chordate refers to organisms with a nerve cord, but “cordate” signifies the heart-shaped form such as is characteristic of many kinds of leaves. No wonder English is such a nightmare to learn!) There may, however, be some tiny calcareous plates and I would like to be able to extract some intact specimens to check that out. Since the effect of glycerine on calcareous structures is relatively slow, I begin with a solution of 15 parts of distilled water, 2 parts of 70 % Isopropyl alcohol, and 1 part of pure glycerine. When these are mixed, there is some turbulence, so combine them and after a minute or so when they are well-mixed, add the solution to the jar of specimens. Leave it uncovered in a dust-free place and as the alcohol and water evaporate, the concentration of the glycerine will gradually increase. The alcohol and glycerine will cause some shrinkage which one hopes is in significant part counterbalanced by the water. After a day, add another part of glycerine and another part of alcohol. As soon as the specimens are flexible enough to be safely removed, transfer them to a Petri dish. In this case I put 6 of the Amphioxus in each of 2 dishes. I immediately removed the fluid in the first dish with a Pasteur pipet, flooded them with distilled water, and then added a few drops of bleach. One wants only enough bleach to make the specimen relatively transparent, but not enough that it begins to dissolve away the tissue. The usual recommendation is for the use of 2% KOH (Potassium hydroxide). This should be monitored every few hours to make sure that is not actually removing the tissue and disassociating it from the specimen. Just out of curiosity, I’m going to try household bleach separately on a couple of specimens as well.

So, this step should be implemented conservatively and with patience. When the specimens are suitably transparent, transfer them into distilled water with a few drops of Alizarine Red S which is a stain with an affinity for calcium. This sort of technique has been used with fingerling fish to demonstrate the skeleton in a spectacular way. If any of the Amphioxus experiments turn out to be interesting, you can be sure an essay on them will magically appear.

The final stages of attempting to restore these specimens involves moving them through higher and higher concentrations of glycerine–30%, 50%, 80%, pure glycerine. Alternatively, one can put them in the 30% solution and then wait weeks or months for the water to evaporate leaving the specimens in “pure” glycerine. Personally, I don’t have that kind of patience, especially at my age.

Finally, for larger specimens–sea cucumbers, echinoids, skates, rays, crabs, etc.–I leave them in 2 gallon or 5 gallon plastic buckets of the type that are often used to ship specimens. If they are in formaldehyde, then I try to anticipate when I might want to work with them and transfer them into one of the buckets containing alcohol. There is little point in keeping preserved specimens if they are not properly maintained. Clearly, only the most compulsive sorts of amateurs will never have any specimens dry up on them and few of us can afford to hire a laboratory assistant to attend to such matters, so if you have a fairly extensive collection, perhaps the best procedure is to set aside a day every two months and go through everything to check fluid levels and replenish them if necessary. In the long run, such a schedule will likely result in saving you a considerable amount of time and effort in not having to try to restore specimens.

All comments to the author Richard Howey are welcomed.

Editor's note: Visit Richard Howey's new website at http://rhowey.googlepages.com/home where he plans to share aspects of his wide interests.

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