Techniques in Entomology Spring 2007 Introduction

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Techniques in Entomology Spring 2007


The most import consideration for how to properly collect, kill, and prepare specimens is what types of analyses will be done. Molecular analyses, such as DNA sequencing, require particular considerations that preserve DNA and are only considered briefly here. However, even in the context of morphological analysis there is no ‘one-size-fits-all’ method for treating specimens. The following sections offer several alternatives for each step of the process and address reasons to apply each. Schauff (1986) offers a good general overview of techniques in collecting and preserving arthropods and is available online.

Killing agents

  1. Freezing: Easiest and non-toxic way of killing arthropods. Of course, this means having a freezer available near the collection site. Sometimes specimens can be kept cool and calm enough (eg. portable cooler) until they are frozen.

    1. Liquid nitrogen: To preserve sequestered toxins and other chemicals for subsequent assays it may be necessary to freeze specimens with nitrogen to prevent break down of compounds.

  2. Cyanide (sodium, potassium, calcium cyanide): Cyanide salts are among the most rapidly acting of all known poisons. In a killing jar cyanide knocks down insects quickly and lasts for a long time. Cyanide binds to cytochrome oxidase which is a catalyst in the oxygenation of bodily tissues, but when cyanide is bound the oxidase can no longer function and the cells of the body are deprived of oxygen. Caution: Cyanide may change the color of some bees and wasps.

  3. Ethyl Acetate (EtOAc): Ethyl acetate is non-hygroscopic and keeps insects soft. EtOAc volatilizes quickly and so needs to be replenished often. It will also cause a killing jar to “sweat” inside sometimes destroying specimens. It is relative non-toxic to humans. However, exposure to high concentrations has a narcotic effect and may cause mucous membrane irritation, and liver and kidney damage.

  4. Ammonium carbonate: As a fumigant killing agent can be obtained fairly cheaply and easily. In small collecting jars it can last for a long time, but may deplete too quick for practical use in large traps. Ammonium carbonate also keeps specimens relaxed until they are pinned.

  5. Alcohol: Various concentrations of ethyl alcohol are common as a killing agent and for subsequent preservation (see below). Some insect taxa and stages, however, don’t do well in alcohol. For example, some larvae will discolor (darken) if killed in alcohol; plant bugs (Miridae) often lose their legs; and Lepidoptera lose their scales. Using a syringe, alcohol can also be injected into the body of large specimens. High concentrations of ethanol tend to “harden” specimens more than high concentrations of Isopropanol, although this is not universally true. However, high concentrations of ethanol will keep the wings from twisting/folding, hairs from matting, and soft organs from shriveling.

  6. Ethylene glycol, Propylene glycol: Ethylene glycol, commonly used as antifreeze, can be used as a killing agent (eg. pitfall traps). However, because of its attractiveness and toxicity to mammals (causes kidney damage), it is slowly being replaced by propylene glycol (also used in cosmetics, food, and medicines). Propylene glycol is hygroscopic (absorbs moisture) and therefore may ‘dry out’ specimens, (ethylene glycol less so). See ‘Preserving Specimens’ below regarding preserving DNA and tissues.

  7. Water: Killing larvae with sub-boiling water has two particular advantages – there is no discoloration as with alcohol, and it leaves larvae ‘inflated’ making the study of setal patterns more convenient. Surfactants in ‘soapy water,’ as is commonly used in pan traps, break the surface tension of the water causing insects/spiders to drown. Absorption of water often results in distended organs, however. DNA in these specimens is most often too degraded to be useful.

  8. Formalin (Formaldehyde): Various fluids (Kahle’s, Bouin’s) containing formalin are used to kill and preserve larvae. These fluids do not result in the discoloration of larvae and fix tissues for future histological work, but may leave specimens too stiff for morphology work.

Handling specimens in the field

  1. Field boxes: Many collectors have elaborate systems for handling specimens in the field including pill boxes, glassine envelops, and spreading boards. Some specimens can be effectively pinned in the field. Others that will later be pointed can be secured in vials or pill boxes with cotton. For Lepidoptera it is preferable to either paralyze them (ie. ‘Pinching’ butterflies) or at least partly spreading the wings of specimens killed in the field. Specimens can also be placed between slightly damp layers of tissue or blotting paper to keep them soft enough for spreading later (addition of chlorocresol will prevent molding). “Papering” large Lepidoptera or Odonata may be a temporary or long-term storage solution whereby specimens are inserted into glassine or paper envelopes with their wings held together dorsally. Paper envelopes will absorb body moisture and help avoid “greasy” specimens. Specimens may also be rehydrated for later spreading.

  2. Alcohol: Alcohol is an easy way to deal with most taxa, but there are aspects to consider. First, alcohol is not allowed on airplanes. Second, if an alcohol vial has air in it the specimen will be sloshed around like it is in a washing machine. Third, some taxa, particularly Lepidoptera and Miridae, can loose important features (scales and legs, respectively). Forth, specimens that will later be mounted dry may not prepare very well (eg. Depending on the alcohol concentration, organs may be distended or, conversely, fixed; wings may be difficult to fully extend).

Preserving specimens

  1. Dry specimens: Except for very small taxa and soft-bodied taxa, most museum specimens are maintained long-term as pinned (or pointed) dry specimens. If concerned only with the chitinous features of the specimen (eg. sclerites, genitalia), dried specimens are adequate. Other tissues, however, may require particular storage techniques. Dean & Ballard (2001) found that pinned, dry Drosophila museum specimens were adequate for DNA sequence analysis after two years, and that exposure to naphthalene had no ill effect of DNA preservation. The factor most important for DNA sequence analysis of dry specimens is how quickly the specimens dry (before DNA degradation) which is dependent on how/where specimens dry and how large is the specimen.

  2. Freezing: Specimens can be frozen alone or in one of the solutions below. The typical household freezer (-20˚C) will preserve internal tissues, but at -80˚C cells and internal tissues can rupture unless stored in ethylene or propylene glycol. External structures (exoskeleton) are generally not effective by a deep freeze.

  3. Alcohol: Alcohols, particularly at higher concentrations, will fix tissues. This may be irrelevant for molecular work, but has some implications for morphological work. 70-80% ethanol is most common for preserving specimens for morphological work, but has been shown to result in degradation of DNA for molecular work (see Dean & Ballard 2001; Vink et al. 2005). When fixing tissues for slide mounting, progressively higher concentrations of ethanol are used to dehydrate tissues because of the hygroscopic nature of alcohol. Before long-term storage the alcohol should be changed from what the specimen was collected in since the alcohol will draw-out the body moisture, thus diluting the alcohol. The alcohol level should also be monitored over time with compensation for evaporation.

  4. Ethylene & Propylene glycol: Ethylene & Propylene glycol are gaining popularity for preserving specimens because they preserve tissue integrity (cells don’t rupture in a -80˚C freezer) and still allow for successful DNA extraction and amplification (see Leal-Klevezas et al. 2000; Vink et al. 2005).

  5. Formalin (Formaldehyde): Formaldehyde fixes tissues or cells by irreversibly cross-linking proteins. It was (is?) commonly used for preserving vertebrate specimens. For insects and spiders, formulations containing formaldehyde, including Kahle’s fluid, Bouin’s fluid, are used to fix tissues for histological work. However, specimens preserved this way may become too hardened for morphological work. Methods of removing formalin from specimens have been developed for analysis of vertebrate DNA (see Kearney & Stuart 2004).

  6. Acetone and other liquids: I mention these here because some Odonata specialists try to maintain specimen color by storing them in acetone.

Handling specimens “back at the lab”

  1. Pinning liquid-preserved specimens: Specimens with hard exoskeletons (eg. Beetles, some bugs, etc.) can be pinned/pointed directly without any more treatment. Others can be treated as follows: 3+ hours in cellosolve to remove moisture; up to 1 hour in xylene to remove cellosolve; air-dry specimen (under a fume hood) occasionally dab with tissue to absorb excess moisture. This treatment will leave hairs, bristles, etc. in an unmatted, natural condition. Pinned specimens may require a little glue since liquid-preserved specimens often don’t ‘grab’ the pin very well. Caution should be observed when using xylene as it carcinogenic, adversely affects the brain, and causes irritation of mucous membranes. HistoClear is a non-toxic alternative to xylene. Also, prolonged exposure to cellosolve can affect the kidneys and liver.

  2. Rehydration: Specimens can be rehydrated for pinning/spreading using a relaxing chamber. Damp paper towels or sand provide moisture that, in a closed container, will relax specimens for manipulation. If specimens require more than a day or two consider adding chlorocresol, phenol, or naphthalene to prevent molding. Very hot water may also be injected into the wing muscles with a very fine syringe. Particular body parts, like legs, may be “softened” independently using a drop of Barber’s fluid or household ammonium.

  3. Critical Point Drying: For this technique specimens are immersed in 100% EtOH which is exchanged (in a critical point drying machine) with liquid CO2. The CO2 is then warmed to a ‘critical point’ and then bled off. By removing moisture from the tissues in the way tissues ‘freeze’ in a more natural position than by air drying.

  4. Freeze-drying: This technique requires freezing the specimen in a natural pose, then removing moisture in a desiccating vacuum. Special expensive equipment is normally required, but “low tech” methods, such as placing a specimen in a desiccating jar in a freezer can also be used, (but take longer).

  5. HMDS (Hexamethyldisilazane): For this technique specimens are soaked in 100% EtOH to remove moisture. Next, specimens are passed through several baths of HMDS. Finally the HMDS is allowed to evaporate (under a fume hood). HMDS is very toxic (can cause severe eye and skin burns) – Acetone is a possible replacement.

  6. Pinning: Be sure to use only insect pins as others will rust or otherwise degrade specimens. Be sure specimens are adequately relaxed before inserting pins to avoid damage to the specimen. There are commonly accepted standards as to how (orientation and location of pin) particular groups of insects should be pinned (consult any entomology text or field guide). What is most important is that all characters for identification and analysis are unobstructed. There is no objective limit to what size specimen should be pinned, (versus double mounted). However, many entomologists prefer not to use size 1 or smaller (0, 00, 000) pins because they are difficult to insert through labels and into pinning surfaces, and often result in abdomens (or other body parts) getting “flinged” across the room (due to their lack of stiffness). If specimens require these smaller size pins consider double-mounting.

  7. Double mounting: Double mounting includes mounting insects on triangular points or minuten pins and foam/cork blocks that are subsequently mounted with size 2 or 3 pins. Points should be made from stiff, acid-free paper, such as is used for labels, so that they do not become yellow and brittle over time. The tip of a point may be folded to create a “shelf” on which the specimen sits. Hide glue is commonly used to glue specimens to points. However, white (Elmer’s®) glue, clear acetate cement, or clear finger nail polish may also be used. Minuten pins come in several lengths and thicknesses. Double mounting strips can be made from several materials including cork, balsa wood, fine foam, or silicone rubber. It is important that the “grain” size of cork or plastic be fine enough that minutens can be inserted straight and are “grabbed” well by the material. Course grain materials may not hold pins well and often result in slanted mounts. Small insects that should have their wings spread, such as Lepidoptera, Trichoptera, and Neuroptera, should be mounted with minutens rather than points. There are commonly accepted standards as to how (orientation) insects should be double mounted (point facing forward, insect facing right – ie. right side of insect stuck to point). Workers in some groups (like Tipulidae and Culicidae), however, may have different preferences (like legs facing towards pin to keep them from getting damaged).


Rearing specimens provide ‘pristine’ specimens as well as access to life stages often not collected, associates larvae with adults, and reveals parasitoid relationships. The most important considerations are containment, proper food, and proper environmental conditions. Some insects also require refugia for part of the day or for pupation.

  1. Acquiring specimens for rearing: Eggs can sometimes be obtained from gravid females. Some female moths will oviposit on the walls of a collecting vial or container. Sometimes it is necessary to have the proper host plant material available to stimulate oviposition. If larvae are collected from host material, be sure to collect extra material.

  2. Rearing containers: Small plants can be grown in screen cages or enclosed in clear plastic bags. However, it could be difficult to collect reared specimens from these containers. Dark cages with collection jars make it easier to collect reared specimens (which are attracted to the jars as an exit). Mason jars with screen or gauze lids work for smaller situations. Be sure to include adequate space for emerging specimens to expand their wings. Also be sure that specimens are not able to chew their way out of the container.

  3. Food: If possible, host plant material should be exchanged regularly with fresh material. For internal feeders, such as leaf miners or seed borers this is not an option. In these cases, great care should be taken to avoid desiccation or molding of the host material (see Environmental Conditions). If the collected specimen was not associated with a host plant a “salad” of plant materials from the surrounding environment should be provided then inspected to see which plant is the preferred host. Carnivorous species can often be maintained on easily massed-reared insects such as waxworms, mealworms, maggots, mosquito larvae, or fruit flies. Some may even take (an appropriately-sized piece of) raw meat dangled from a thread.

  4. Artificial medium: Artificial rearing media have been formulated for many agricultural and other economic pests and work for many related groups. Most consist of an agar base with nutrients and feeding stimuli added and can be obtained from biological supply houses.

  5. Environmental conditions: Temperature, humidity, and day-length should simulate natural conditions as much as possible. Environmentally controlled rearing facilities are preferred, but low tech substitutions work nearly as well. A damp cotton ball, sponge, or paper towel can maintain elevated humidity, but particular humidity requirements can be maintained with saturated salt solutions (Winston & Bates, 1960). Pupae may be placed on damp sawdust or moss for eclosion. For larger cages with plants, occasional misting with a spray bottle may be appropriate. Excess moisture can lead to mold or fungus growth or can trap insects in the surface tension. Direct sunlight may cause over-heating and moisture build up in containers, indirect sunlight is preferred. Some species may require a “cold shock” that cues the insect that winter has passed and to break dormancy. This can be simulated with refrigeration. “Grow lights” may also be used to control day-length.

  6. Pupation: Some species require particular microhabitats for pupation. Layered paper towels or corrugated cardboard can be substituted for loose soil and leaf litter as pupation sites. Moisture is crucial at eclosion – use moist saw dust, moss, sponge, cotton, or paper towel to maintain elevated humidity, but take care that pupae are not in direct contact with water.

  7. Sanitation: Waste products, such as frass and the remains of eaten host material, should be removed regularly to avoid build up of unsanitary conditions. Occasionally wiping down the rearing container with a dilute bleach solution or ethanol will help prevent the growth of mold and bacteria. Good ventilation/air circulation within the rearing setup will also prevent moldy conditions.

  8. Mating studies: Place males and females in a vial for a day or two. Host (oviposition) material may help stimulate mating and oviposition.

  9. Culturing: Maintaining cultures of particular species is beyond the scope of this discussion. However, the above considerations will apply in most cases. Consult specific references for the particular needs of the species of interest.

References Cited

Dean MD, Ballard JWO. 2001. Factors affecting mitochondrial DNA quality from museum preserved Drosophila simulans. Entomologia Experimentalis Et Applicata 98: 279-83

Kearney M, Stuart B. 2004. Repeated evolution of limblessness and digging heads in worm lizards revealed by DNA from old bones. Proceedings of the Royal Society B: Biological Sciences 271: 1677-83

Leal-Klevezas DS, Martinez-Vazquez IO, Cuevas-Hernandez B, Martinez-Soriano JP. 2000. Antifreeze Solution Improves DNA Recovery by Preserving the Integrity of Pathogen-Infected Blood and Other Tissues. Clinical and Diagnostic Laboratory Immunology 7: 945-6 { continued as Clinical and Vaccination Immunology }

Schauff ME. 1986 (revised 200?). Collecting and preserving insects and mites. USDA Miscellaneous Publication no. 1443. 68 p. {also available online at}

Vink CJ, Thomas SM, Paquin P, Hayashi CY, Hedin M. 2005. The effects of preservatives and temperature on arachnid DNA. Invertebrate Systematics 19(2): 99-104

Winston PW, Bates DH. 1960. Saturated solutions for the control of humidity in biological research. Ecology 41(1): 232-237


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