2.1 - Liquid Agents for Killing and Preserving

Insects and mites of all kinds may be killed and preserved in liquid agents, but it is first necessary to determine the advisability of using a liquid killing agent rather than a dry gaseous agent. Some kinds of insects are best kept dry; it may not be advisable to allow others to become dry. Directions for the treatment of various insects are given in the last part of this publication under the various orders.

Ethanol (grain or ethyl alcohol) mixed with water (70 to 80 percent alcohol) is usually the best general killing and preserving agent. For some kinds of insects and mites, other preservatives or higher or lower concentrations of alcohol may be better. Because pure ethanol is often difficult to obtain, some collectors use isopropanol (isopropyl alcohol) with generally satisfactory results. Isopropanol does not seem to harden specimens as much as ethanol, and at least it is satisfactory in an emergency. Although there is controversy over the relative merits of ethanol and isopropanol, the choice of which to use is not so important as what concentration to use. This choice depends on the kind of insect or mite to be preserved.

Parasitic Hymenoptera are best killed and preserved in 95 percent alcohol. This high concentration prevents the membranous wings from becoming twisted and folded, hairs from matting, and soft body parts from shriveling. This concentration may also be desirable if large numbers of insects are to be killed in a single container, such as in the killing jar of a Malaise trap, because the insect body fluids will dilute the alcohol. On the other hand, soft- bodied insects, such as aphids and thrips, small flies, and mites, become stiff and distorted if preserved in 95 percent alcohol and should be preserved in alcohol of a lower concentration. Adult bees should not be collected in alcohol because their usually abundant body hairs become badly matted. Adult moths, butterflies, mosquitoes, moth flies, and other groups with scales and long, fine hairs on the wings or body may be worthless if collected in alcohol regardless of the concentration.

Formalin (formaldehyde) solutions should not be used because the tissues become excessively hardened and the specimens then become difficult to handle.

Larvae of most insects should be collected in alcohol and subsequently killed in boiling water to "fix" their proteins and prevent them from turning black. Larvae should be left in hot water for 1-5 minutes, depending on the size of the specimens, then transferred to 70-80 percent alcohol. Large specimens or small specimens that have been crowded into one vial should be transferred to fresh alcohol within a day or two to reduce the danger of diluting the alcohol with body fluids. If the alcohol becomes too diluted, the specimens will begin to decompose. Water is not a preservative.

For some groups, preservation is better if certain substances are added to the alcohol solution. Thrips and most mites, for example, are best collected in an alcohol- glycerin-acetic acid (AGA) solution, and for many larvae a kerosene-acetic acid-dioxane (KAAD) solution is preferred. If KAAD is used, larvae need not be killed in boiling water. Formulas for these and other solutions are given in the Appendix.

For some histological, cytological, or physiological studies, specimens must be in a certain critical condition and must be preserved in special media (see Walker & Boreham 1976).

Larvae and most soft-bodied adult insects and mites can be kept almost indefinitely in liquid preservatives; however, for a permanent collection, mites, aphids, thrips, whiteflies, fleas, and lice usually are mounted on microscope slides (see p. 36). Larvae are usually kept permanently in alcohol, but some may be mounted by the freeze-drying technique (see p. 34) or by inflation (see p. 34). Many insects collected in alcohol are later pinned for placement in a permanent collection. Hardbodied insects such as beetles can be pinned directly after removal from alcohol, but for them and all softer insects such as flies and wasps special procedures must be followed.

2.2 - Temporary Storage of Specimens

After specimens have been collected, time is often not immediately available to prepare them for permanent storage. There are several ways to keep them in good condition until they can be prepared properly. The method used depends largely on the length of time that the specimens may have to be stored temporarily.

2.2.1 - Refrigeration and Freezing

Medium to large specimens may be left in tightly closed bottles for several days in a refrigerator and still remain in good condition for pinning as will smaller specimens if left overnight. Some moisture must be present in the containers so that the specimens do not become "freeze-dried," but if there is too much moisture, it will condense on the inside of the bottle as soon as it becomes chilled. Absorbent paper placed between the jar and the insects will keep them dry. When specimens are removed for further treatment, place them immediately on absorbent paper to prevent moisture from condensing on them.

Insects may be placed in alcohol, as described previously, and kept for several years before they are pinned or otherwise treated. However, it has been shown that many insects, especially small ones, can deteriorate in alcohol stored at room temperature. Long term storage of specimens that suffer from this kind of deterioration can be lessened by storing the containers in a freezer. Even though the alcohol will not freeze at the temperatures obtained by most ordinary freezers, the lower temperature seems to slow or stop deterioration of the specimens.

2.2.2 - Dry Preservation

It is standard practice to place many kinds of insects in small boxes, paper tubes, triangles, or envelopes for an indefinite period, allowing them to become dry. It is not advisable to store soft-bodied insects by such methods because they become badly shriveled and very subject to breakage. Diptera should never be dried in this manner because the head, legs, and most of all the antennae become detached very easily.

Almost any kind of container may be used for dry storage; however, tightly closed, impervious containers of metal, glass, or plastic should be avoided because mold may develop on specimens if even a small amount of moisture is entrapped. Nothing can be done to restore a moldy specimen.

Dry-stored specimens must be labeled with complete collection data in or on each container. Avoid placing specimens collected at different times or places in the same container. If specimens with different collection data must be layered in the same container, include a separate data slip with each layer.

To insure that specimens do not slip from one layer to another, cut pieces of absorbent tissue, glazed cotton, or cellucotton a little larger than the inside of the container. Place a few layers of this material in the bottom of the container, then a few insects (do not crowd them), then more layering material, and so on until the container finally is filled. If much space is left, use a little plain cotton, enoush to keep the insects from moving about but not enough to produce pressure that will damage them. To prevent parts of the insects from getting caught in the loose fibers, use plain cotton only for the final layer. Insect parts are very difficult to extract from plain cotton without damage.

One method of keeping layered specimens soft and pliable for several months includes the use of chlorocresol in the bottom of the layered container and a damp piece of blotting paper in the top. The container must be impermeable and sealed while stored; plastic sandwich boxes make useful containers to use with this method. Add about a teaspoonful of chlorocresol crystals to the bottom, cover with a layer of absorbent tissue, follow with the layers of specimens, then a few layers of tissue, and finally a piece of dampened blotting paper as the top layer. The cover is then put in place and sealed with masking tape. It is best to keep boxes of layered specimens in a refrigerator.

Reference: Tindale 1962.

Some insects, such as small beetles, should be glued to points (see p. 29) directly from the layers for permanent preservation, but if they are to be pinned or otherwise treated, they must be relaxed as described on page 25.

2.2.3 - Papering

Although pinning specimens when they are fresh is preferable, the storage method known as papering has long been used successfully for larger specimens of Lepidoptera, Trichoptera, Neuroptera, Odonata, and some other groups. It is a traditional way of storing unmounted butterflies and is satisfactory for some moths, although moths too often will have their relatively soft bodies flattened, legs or palpi broken, and the vestiture of the body partly rubbed off. To save space in most large collections, file Odonata permanently in clear plastic envelopes instead of pinning them.

 

Figure 16. Lepidoptera temporarily in paper and in glassine envelope

Papering consists of placing specimens with the wings folded together dorsally (upper sides together) in folded triangles (fig. 16) or in small rectangular envelopes of glassine paper, which are the translucent envelopes familiar to stamp collectors. Glassine envelopes have become almost universally used in recent years because of the obvious advantages of transparency and ready availability. In many collections, glassine has become partly superseded by plastic. However, many collectors still prefer folded triangles of a softer, more absorbent paper, such as ordinary newsprint, and believe they are superior for preserving specimens. Specimens can become greasy after a time, and the oil is absorbed by paper such as newsprint but not by glassine. Moreover, glassine and plastic are very smooth, and specimens may slide about inside the envelopes during shipping, losing antennae and other brittle parts. Although softer kinds of paper do not retain creases well when folded, this shortcoming may be circumvented by preparing the triangles of such material well before they are needed and pressing them with a weight for a week or so. Triangles are easy to prepare.

Some Lepidoptera are most easily papered if first placed in a relaxing box (see p. 24) for a day or two. The wings, often reversed in field-collected butterflies, may then be folded the proper way without difficulty. Do not pack specimens together tightly before they are dried or the bodies may be crushed. Do not store fresh specimens immediately in airtight containers or plastic envelopes or they will mold. Write collection data on the outside of the envelopes before inserting the insects.

2.2.4 - Liquid Preservation

Preservation of insects in alcohol is a complex subject and like many things, it varies somewhat from one group to another. For example, spiders preserve well in ethanol, but tend to become to flaccid in isopropyl. The opposite is true for many myriapods. If one specializes in an insect group suited to preservation in one or another kind or concentration of alcohol, consult specialists in that group or experiment to find what yields the best results.

In general, ethanol and isopropanol mixed with water is the most widely used preservation fluids. Most commonly, a mixture of 75% alcohol to 25% water is used. The water should be distilled to ensure a neutral PH and the solution should be thoroughly mixed since alchols and water do not mix easily by themselves. Additives should be avoided.

Special care should be taken with labels placed in alcohol. Paper should be high quality rag or linen and acid-free. The ink should contain vegetable gum (such as India inks) as these seem to withstand the constant exposure to the alcohol the best. Typewritten labels and computer generated (laser printed) labels are generally unacceptable. The best system is still professionally printed labels.

Shell vials plugged by cotton or with polyethylene stoppers are recommended. Avoid stoppers made from cork, rubber, or neoprene as they tend to degrade and/or leach chemicals into the alcohol. Shell vials are preferred over necked vials as it is easier to remove the specimen and the chance of damage is reduced. Each vial should be individually labelled with complete collection data.

The shell vials are kept in wide mouthed, gasketed bail-top jars with straight sides (fig. 17). Avoid metal screw caps, bakelite lids, greased glass, and ground glass as they may rust, warp, crack, leak, or allow the alcohol to evaporate. Generally it is recommended that each jar contain between 10 and 40 vials. Avoid glass-glass contact by plaing a folded paper towel in the bottom of each jar. Keep vials upright within jars. Each jar should be filled with alcohol to just below the level of the gasket. If material is going to be stored for long periods of time, the jars should be checked periodically and the alcohol topped off. Labels may also be placed on the outside of the jars to indicate the enclosed contents.

 

Figure 17. Alcohol storage jars

Light is the chief enemy of alcohol preserved material, and as a result, jars should be stored inside cabinets. Vibration can also damage specimens and cause lids and caps to come loose so it is a good idea to place cabinets in a location where vibration is at a minimum.

Fire safety is always an important consideration when storing or working with alcohol collections. Concentrations of vapors can be very hazardous so care should be taken that work areas are properly ventilated and that there is no source of open flame nearby. In larger collections, special cabinets, exits, and other precautions may be necessary to meet the fire code.

References (J. Coddington, personal communication; Roth, 1952; Levi, 1966; Jocqué, 1983).

2.3- Preservation for Molecular Studies

Systematists are increasingly using molecular methods to study insect relationships, make identifications, and determine species limits. Some of these techniques, such as the study of cuticular hydrocarbons, can be used on dried insects, even those stored in museum collections. However, many others require that specimens be treated so that DNA or other molecules are preserved. In general, specimens for molecular work should be collected in 95% or absolute (100%) ethanol (ethyl alcohol). It is best if specimens are thoroughly dehyrated by changing the alcohol at least a couple of time before the specimens are stored for any length of time. It is also advisable to keep specimens cold (frozen if possible). For more detailed information on specimen preservation for molecular work see Hillis, et al. (1996).