CHAPTER 6

SPECIMEN PREPARATION

Insect specimen preparation methods are discussed in a wide variety of books and manuals. However, the salient aspects are treated here to correspond with preferences and protocols established for the INBio inventory.

Killing

All of the beetle specimens that you collect will be killed at some point in time. This is a necessary prerequisite for working with the specimens as data vouchers, integrating them into collections, and allowing them to be studied subsequently.

There are numerous methods for the lethal dispatch of beetles and other insects. The two most common chemicals are ethyl acetate and sodium cyanide. Both are effective and have their benefits and detractions. Ethyl acetate is inexpensive, usually easy to obtain, kills insects quickly, including large beetles, and has the added benefit of helping to maintain softness and flexibility of the specimens for long periods of time. Two negative effect of ethyl acetate are the moisture that readily develops in jars with this chemical can mat the hairs of a beetle, and that fatty tissues in the beetle can sometime liquify and spread and oily film over the body of the specimen. Both of these latter situations can be offset by specimen cleaning with chemical solvents (see below). Despite these negatives, many people prefer the availability, ease of use, and effectiveness of judiscious amounts of ethyl acetate. This chemical is the active ingredient in finger-nail polish remover, so despite the added perfume, this can be used in emergency situations.

The use of cyanide by collectors, usually sodium cyanide, is common. Potassium cyanide will also work, but has a shorter life-span. Both cyanide formulations are highly effective for the killing of insects and other arthropods. These chemicals usually are available in crystal form, but only from chemical suppliers, and are often heavily regulated as toxins in many countries. In the collecting jar, the crystals are activated by contact with water moisture and the reaction for hydrogen cyanide gas that actually does the killing of the beetles. Cyanide is fast and effective, and bottles with cyanide will usually stay dry, thus helping to prevent matting of hairs. Body fats of beetles are rarely liquified by cyanide treatment, so that the specimens will stay cleaner. Drawbacks to cyanide are a drying effect on the specimens requiring that material be prepared quickly or subsequently relaxed, color changes in pigments of some beetles if left too long exposed to the cyanide, and difficulty in obtaining the chemical.

Both ethyl acetate and cyanide collecting bottles require caution in their use. Both chemicals will harm the user, other persons, and other organisms, if not safely handled.

Killing jar
 

Pinning

Techniques of pinning were demonstrated by curators or specialists at workshops. Here, only general considerations are discussed. Refer to figures __-__ for placement of pins through the body of beetles. Generally speaking, the pin should be placed approximately 0.2-0.3X of the length of the right elytron from the base of the elytron, and at midwidth of the elytron. The specimen needs to be adjusted on the pin to permit finger-tip room at the top for handling and adequate room beneath for labels. For many beetles, approximately 10 mm of gap should be present from the top of the beetle to the pinhead.

Pins for preparing beetles for study or display are a special type designed for entomological use. They are specially made of spring steel or stainless steel. Spring steel pins are usually painted with a black protective coating that resists corrosion. Because of the corrosion problem, inventory programs are encouraged to use stainless steel pins, despite their higher initial cost, to enhance long-term protection of specimens and reduce re-pinning demands and increased curatorial costs in the future. Common, or sewing, pins are not adequate for working with insects because of their short length, thick diameter, and low quality leading to corrosion.

Entomological pins come in a wide variety of sizes to meet the many needs of entomologists. However, for beetles, the #2 and #3 sized pins are the most commonly used. Parataxonomists working with very large beetles, such as many scarabs or cerambycids, may need a supply of larger #5 pins.

The choice of pin size for a particular specimen is dependent upon evaluating the degree of damage done to a specimen by the insertion of the pin. In general, use the smallest pin available that will still provide adequate support for the specimen. Many specialists will not use a pin smaller than a #2 for direct pinning. If the width of the specimen is less than 0.2X the diameter of the pin, then the damage done to the specimen will often be considerable. Under this criterion, the #2 and #3 pins are generally the most used for beetles. Smaller specimens should be pointed.

 

Entomological pins

Pointing

Points are paper triangles to which small beetles are attached by glue. Points can vary in size for special uses, but standardized point sizes are available with special punches. Points can be made by using scissors, but this is not recommended unless necessary. A typical point is 7 mm long and 2 mm wide at the base. An insect pin is inserted through the base and the beetle is glued to the pointed tip.

Points

Specimens are glued to the tip of the point. Because of the rounded shape of many beetles, the tip of the point should be bent downward at an angle that is customized to fit the specimen at hand. Bending the point is easily done with the nails of the index finger and thumb, or a fine pair of forceps. Shaping the bend of the point with a curvature will help adhesion to many beetles, but must be assessed on a specimen by specimen basis by the preparer. Also, for many beetles, clipping a short length of the point to provide a broader transverse surface for glue may be beneficial.

Appropriate paper should be used for points. A thickened or multilayered paper of archival quality is best. However, common index cards are frequently used. Archival paper is essential permanent and has a neutralized acid content. Index card paper is frequently of low quality and has a high acid content which increases pin corrosion, and yellowing and decomposition of the paper.

Glues of adhering specimens to points, or repairing broken specimens, provide a constantly changing debate for entomologists. Unfortunately, the "perfect" glue seems not to exist. Many people use common "white" glue. This is a convenient, inexpensive and easily obtained alternative, but such glues do not dry quickly enough in humid situations and will loosen their hold on smooth beetles after a few years. Chemically, these are too unstable for permanent use.

A popular and traditional alternative used by many entomologists is shellac gel. This is made by boiling shellac to change it from a thin liquid into a thicker nearly gel-like substance, and adding alcohol for dilution when necessary. Shellac is a good alternative to white glue. With an alcohol base it will dry faster than white glue, but will still have some problems in very humid conditions. It is more permanent, but is difficult or impossible to dissolve when needed for studying beetles in detail.

One substance that may have good potential for use in tropical environments is a translucent polyvinyl acetate material with the tradename of "Gelva." Gelva is alcohol based and dries quickly. It is easy and safe to work with, is easily thinned with 95% ethanol when necessary, and seems to be permanent. Subsequent study of beetles is easily done as dry Gelva will dissolve in alcohol or ammonia.

Other glues commonly used but with many drawbacks include the commercial instant-glues, epoxies, cements, and fingernail polish. None of these have properties that are acceptable for permanent care and study of beetles, and many have essential chemicals that are toxic to people. Common fingernail polish falls into this category; although it dries fast, is inexpensive, and is easy to use, it contains many solvents, is not permanent in collections, and will not dissolve when specimens need to be studied.

 

Relaxing

When the parataxonomist is collecting a lot of materials, there may not be time for each lot of specimens to be properly pinned or pointed immediately. If specimens dry before preparation they will require relaxation to prevent breakage during preparation. Also, specimens that may be coated with soil, grease, or other contaminants may require cleaning. Both the relaxation and cleaning of specimens can be achieved with simple materials.

Relaxation of specimens can be done passively within a humidity chamber or by immersion of the specimens into a water-based liquid. A humidity chamber can be nothing more complicated than a plastic container with a lid. The container should be at least 4-5 cm deep to allow room for large specimens and development of a good atmosphere. Place an absorbant material, such as a sponge or multiple layers of paper towel, on the bottom of the container. Dampen this material with water. Place the beetle specimens on a tissue or paper towel section and place this section and the beetles on a small dish or other non-absorbant surface that is then placed on top of the absorbant material. Close the container and allow the specimens to absorb moisture for 1-2 days. Check the specimens frequently. If small beetles soften quickly, prepare them as they become relaxed. Never keep the specimens in a container for more than 2 days because mold will begin to grow and quickly damage the beetles, or the body parts will begin to separate by deterioration of the connecting membranes. Also, too long of time in the chamber may change some colors on some beetles. When specimens are relaxed, this is the time to take a fine brush and gently clean them of contaminants.

An alternative to the passive humidity chamber is the use of hot water. Many beetles can be quickly relaxed by being placed into hot water for a few minutes. Simply bring a small pan of water to near boiling, remove the pan and water from the heat, and place the beetles into the water until they are relaxed. Steaming the beetles by placing them on paper towel resting on a screen over the hot water is also effective. In either case, beetles relaxed in this manner must be prepared quickly. Do not steam more specimens than you can prepare in a short period of time, as repeated steaming will damage the beetles and cause the body parts to separate.

One simple, fast, and effective way to relax and clean specimens is to use common household ammonia solutions. Ammonia is a common cleaning solution available in markets. For already clean specimens, simply prepare a humidity chamber as normal, but instead of water use ammonia to wet the absorbant material. The ammonia atmosphere will relax the specimens more quickly and will retart the growth of mold. However, do not keep the specimens in an ammonia chamber for more than 2-3 days as the body disarticulation may occur.

An alternative use of ammonia for even more rapid relaxation and cleaning of specimens is to immerse the beetles into a small dish of ammonia. The body will absorb moisture and the ammonia will cause adhering soil to fall away or being easily removed with a fine brush. When the specimen is relaxed, gently place it onto an absorbant tissue to draw the liquid off the beetles' body, and prepare normally. Ammonia is also good for dissolving most greasy films from liquified body fats. Immersion relaxation of beetles is a good method for rapid relaxation of small lots of specimens, or cleaning of individual specimens.

 

Spreading and Arrangement

Many beetles do not require extensive manipulation when preparing them. However, to help protect body parts, save space in collection boxes, and to provide a neat and orderly appearance, the arrangement of legs and antennae should be considered. The spreading of wings on beetles, as is done with butterflies and some other insects, is not normally done. Methods for arranging beetles body parts will be demonstrated by a curator or specialist.

Immediately following the pinning or pointing of a specimen, it should be placed on a foam board that has a sheet of paper pinned down. This paper helps to provide a smooth surface to prevent breakage of tarsi and other body parts, and to absorb moisture. Push the specimen pin through the paper and into the foam board until it just touches the paper. You may now arrange the legs, antennae, elytra, or mouthparts as desired, using insect pins to hold structures in place until the specimen is dry.

All specimens must be dried. Drying is necessary to remove body moisture from the specimens in order to retain positions of arranged appendages, and to inhibit decay and loss of specimens. In air conditioned work areas, specimens can usually be placed into cabinets or on shelves, away from ants, wasps, and birds, and allowed to dry under ambient conditions. In non-air conditioned spaces, particularly when the humidity is high, the specimens may require heat drying. An effective method is the use of a gas-fired oven on its lowest setting and with the door slightly open.

 

Labelling

All specimens for inventories and scientific use lose value if they lack essential data for understanding their geography and ecology. Essential information for each specimen is the location, date of capture, and collector name. Location data includes the political units of country, province, district, distance and direction from a town or landmark (if appropriate), longitude and latitude, and altitude (if available). The date of collection is vital and should never be compromised. The name of the collector is important for attributing responsibility, or glory, for each specimen. Additional information that is pertinent and should always be provided when available is lot numbers, trap type or collecting method, and host or habitat information. Other information, such as temperature, time of day or night, weather conditions, or other factors that may be important for a particular specimen or situation should also be recorded. All of this information will be databased for retrieval and provides the scientific power of good inventory projects.

Field labels, when used, are simple notes that lead to more detailed records and information on each specimen. However, field labels should always provide a minimum set of information in case notebooks are misplaced or lost. Minimum information would be a location, the date of capture, and collector name. All specimens, or series of specimens in a collection box should have at least field labels clearly and unambiguously associated with them.

 

Liquid Preservation

The liquid preservation of beetles is usually done either for bulk storage of numerous specimens, or the immature stages. Specific treatment of beetle immatures was discussed above, but storage and chemical solution information is provided here.

Vials

Usually, specimens are kept in glass vials, these being small glass bottles with either screwcap or stopper-type closures. The choice between size of vial and style of closures is usually preferential to the user, but longterm storage necessitates prevention of preservative leakage. Accessibility of specimens requires a vial style and closure type that can properly seal as well as be repeatedly functional. As such, materials availability may change depending on curatorial needs and suppliers.

The parataxonomist will usually need relatively small volume vials with ease of use being priorities. For most purposes, vials with screwcap closures bearing a flexible polyvinyl or Teflon-type liner are best for reusability and prevention of leaks. Such closures provide excellent short-term storage and are also useful for longterm storage of specimens. Vial size is highly variable and sizes should be chosen for a specific use. Commonly used sizes include those measuring 17x60 mm and 21x70 mm. Scintillation vials of 20 ml volume and measuring 28x61 mm with polyvinyl cone inserts in the closures are an excellent choice for both fieldwork and storage of few specimens per vial.

Shell vials with snap-caps are inexpensive and easy to use for field and temporary use. However, these should not be used for permanent storage of specimens. The caps lose their seal after 2-3 uses.

Plastic Bags

The temporary preservation of large or bulk samples is sometimes necessary. Without large and heavy bottles and jars, the saving and preserving of such samples can be problematic. Laboratory quality zip-lock style or while-pak style plastic bags are excellent alternatives to heavy glass or bulky rigid containers. These bags can be used for temporary storage of alcohol preserved specimens. As with all samples, alcohol solutions should be changed frequently when samples are fresh to compensate for body fluid dilution. It is also recommended that for storage periods exceeding 2-3 weeks or shipment that the bags be doubled to help retart leaks and drying of samples, or dilution of solution. Double-check all closures for tightness before storing or shipping bags of specimens.

Preservatives

Both short-term and long-term liquid preservation of beetles requires use of appropriate solutions. The most commonly used and basic chemical is ethyl alcohol, or ethanol, in 75-80% solution. Stronger solutions may be used if the collector anticipates excessive dilution of field solutions, such as the collecting of immatures or aquatic beetles. Long-term storage solutions for beetles may be as low as 70% concentration.

Ethanol is preferable to either isopropyl or methyl (methanol) alcohols, as the latter types remove too much water from specimen tissues and causes them to become brittle. This inhibits subsequent dissection and study. Also isopropyl and methyl alcohols are less compatible with other chemicals and solutions, and methanol is highly toxic. Neither alcohol is good at longterm preservation of scientific quality specimens. Ethanol has a wider chemical compatibility, is relatively non-toxic, and is an excellent long-term preservative. Ethanol has a low skin absorbancy, so is not usually a concern in this regard, but drying of the skin can be a problem for some people. Ingestion of ethanol should be prevented as reagent grade ethanol is chemically dried with benzene, which is toxic.

Remember, as discussed above under immatures, ethanol is a preservative, not a fixative of tissues. This means that specimens should be killed in a method other than drowning in ethanol, or that the ethanol solution be changed 2-3 times over a 2-5 day interval following collecting, so that dilution from body fluids does not interfere with preservation.

Acetic ethanol is a commonly used substitute for plain ethanol solutions, particular for field use. This solution is made with 80% ethanol and glacial acetic acid, usually in a 19:1 ratio. The acetic acid quickly penetrates beetle tissues and quickly kills specimens. The ethanol in solution then acts as a preservative. However, as with plain ethanol solutions, field samples should have the fluid replaced to reduce the effects of body fluid dilution. Acetic ethanol is most commonly used for immatures, but is also useful for adult beetles. Caution must be used as the acetic acid will also quickly penetrate human skin and cause drying, or tissue damage with extensive exposure.

Another common solution for beetles, particularly immatures, is "KAA." This is a solution of acetic ethanol with clarified kerosene. A basic solution can be made with the acetic ethanol solution mentioned above, with the addition of 0.5 parts of kersosene. A good quality kerosene must be used to prevent chemical separation and ensure proper activity of the solution. KAA is more commonly used for preservation of Lepidoptera larvae, but is also used for beetle larvae. The solution acts much as acetic ethanol, but the kerosene helps to distend specimens so that small body parts are more readily studied. Kerosene imparts a strong odor to the solution and the combination of acetic acid and kerosene should not be allowed on the skin or clothing. Also, specimens initially killed in KAA should have the solution changed to acetic ethanol or plain ethanol within 2-4 days to compensate for body fluid dilution and for long-term preservation.

 

 

 

ACKNOWLEDGMENTS

 

This manual evolved to address needs and standardize inventory-related sampling protocols of the parataxonomists and curators at the Instituto Nacional de Biodiversidid (INBio). As such, numerous persons were involved at various levels and through the years of the Coleoptera invenory, and our gratitude is immense for the assistance of so many people. Alvaro Herrera, Jesus Ugalde, and Manuel Zumbado are thanked for their considerable efforts to keep the arthropod inventory efforts working smoothly and by considerable logistical and administrative support to us and other specialists. Angela Mora and Elena Ulate, and Carlos Hernandez and Carlos Guzmán, provided efforts and assistance that went well beyond their positions as curatorial technicians. The parataxonomists were a large group and we greatly appreciate the work of all of them, with special thanks to Wendy Porras, Wilfredo Arana, Alejandro Azofeifa, and Roger Gonzalez, who gave considerably of their time and dedication in the field to visiting specialists and took special effort to advance their training and knowledge of beetles and conservation biology. Paul Hanson and Carolina Godoy, provided generous collegiality, advice and friendship. The entire support staff of INBio is thanked for their friendship and professionalism during the course of the inventory and compilation of this manual.

 

 

REFERENCES

Aiken, Ronald B., and Robert E. Roughley. 1985. An effective trapping and marking method for aquatic beetles. Proceedings of the Academy of Natural Sciences of Philadelphia 137: 5-7.

Hilsenhoff, W. L. 1991. Comparison of bottle traps with a D-frame net for collecting adults and larvae of Dytiscidae and Hydrophilidae (Coleoptera). The Coleoperistst Bulletin 45(2):143-146.

Hilsenhoff, W. L. and B. H. Tracy. 1985. Techniques for collecting water beetles from lentic habitats. Proceedings of the Academy of Natural Sciences of Philadelphia, 137: 8-11.

Martin, J.E.H. 1977. Collecting, preparing and preserving insects, mites and spiders. The Insects and Arachnids of Canada. Part 1. Publication 1643. Canada Department of Agriculture, Ottawa. 182 pp.

Oldroyd, H. 1958. Collecting, preserving and studying insects. The Macmillan Company, New York, 327p.

Oman, P.W. 1946. Collection and preservation of insects. U.S. Department of Agriculture, Milscellaneous Publication No. 601, 42p.

Schauf, M.E. Undated. Collecting and preserving insect and mites: techniques and tools. U.S. Department of Agriculture, Systematic Entomology Laboratory.

Steyskal, G.C., W.L. Murphy, and E.M. Hoover. Editors. 1986. Insects and mites: techniques for collection and preservation. United States Department of Agriculture. Agricultural Research Service. Miscellaneous Publication, # 1443. 103 pp.

Spangler, P. J. 1991. A durable, lightweight net and a manual aspirator for collecting aquatic organisms. The Pan-Pacific Entomologist. 1981; 57: 245-250.


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