Collecting and Preserving Insects and Mites: Techniques and Tools

2.2.1 - Refrigeration and Freezing ............................................................................ 25 2.2.2 - Dry preservation ............................................................................................ 26 2.2.3 - Papering ......................................................................................................... 26 2.2.4 - Liquid Preservation ........................................................................................ 26 2.3- Preservation for Molecular Studies ................................................................................ 27 Part 3. Mounting Specimens ............................................................................................................ 27 3.1 - Preparing Dry Specimens for Mounting........................................................................ 28 3.2 - Preparing Liquid-Preserved Specimens ........................................................................ 30 3.3 - Direct Pinning ............................................................................................................ 31 3.4 - Double Mounts ............................................................................................................ 32 3.5 - Spreading Boards and Blocks ........................................................................................ 35 3.5.1 - Construction of Spreading Boards. ................................................................ 35 3.5.2 - Using the Spreading Boards .......................................................................... 36 3.5.3 - Construction of Spreading Blocks. ................................................................ 37 3.6 - Riker Mounts ............................................................................................................ 38 3.7 - Inflation of Larvae ......................................................................................................... 38 3.8 - Artificial Drying ............................................................................................................ 38 3.9 - Embedding ............................................................................................................ 39 3.10 - Mounting Specimens for Microscopic Examination ................................................... 39 Part 4. - Sample Procedures 43 4.1 - Preparation and Storage of Genitalia. ............................................................................ 43 4.2 - Mounting Wings ............................................................................................................ 44 4.3 - Mounting Larvae of Diptera, Coleoptera, Lepidoptera, and Other Groups. ................. 45 Part 5 - Labeling ............................................................................................................ 46 5.1 - Paper ............................................................................................................ 46 5.2 - Ink ............................................................................................................ 46 5.3 - Lettered and Printed Labels ........................................................................................... 46 5.4 - Size of Labels ............................................................................................................ 46 5.5 - Label Data ............................................................................................................ 47 Introduction The Class Arthropoda, which includes insects, spiders, mites, and their relatives, is without question the most successful group of organisms on the planet. Insects alone account for nearly 55% of all species known to science (Barrowclough 1992). Spiders, mites and insects inhabit every terrestrial habitat on the planet and play a major role in the evolution and maintenance of biotic communities. They are the primary pollinators of flowering plants; they are important consumers and recyclers of decaying organic matter; and they are integral components in the foodwebs of vertebrates and other invertebrates. For these reasons, and many others, the study of insects and their relatives is of increasing importance as society faces increased challenges to preserve and enhance environmental quality, reduce pesticide usage, increase crop productivity, control food costs, and increase trade in the global community. Pest species are responsible for enormous economic losses annually, attacking crops and ornamental plants, causing damage to our food and clothing, and vectoring diseases that effect cultivated plants, our pets and livestock, and ourselves. The damage cause by pests species is far outweighed by the positive effects of beneficial species. Pollinators ensure the production of fruit, parasitoids and predators help control pest species, some species contain chemicals of pharmaceutical value, and a large number of species contribute to the decomposition and recycling of dead and decaying matter. Because of the damage inflicted by pest species, increased knowledge of these organisms has the potential to save lives and money. Correct identification of a newly detected pest or disease vector is of utmost importance because the scientific name of an organism is the key to all known information about its morphology, its behavior and life history, and its potential threat to human welfare. The behavior of insects and mites can be observed most easily in their natural environments. However, many species, especially the smaller ones, must be collected and properly preserved before they can be identified. Because correct identification seldom is easy, it is important that specimens be preserved in the best condition possible. The identification of a particular insect or mite usually requires examination of minute details of its anatomy with the aid of a hand lens or microscope. Some specimens may require dissection or even study with the electron microscope. If these details on a specimen are concealed, missing, or destroyed because of improper handling or preservation, identification is made difficult or impossible, and information about the species to which it belongs cannot be made available. Therefore, adequate preservation and proper labeling of specimens are essential to their identification. The methods used to collect insects and mites are dictated by the ultimate goal of the samples collected. Insects may be collected as a hobby for personal enjoyment of their diversity and beauty. They may be collected in conjunction with school courses on biology or entomology. Specific insects groups may be sampled to assess or measure biodiversity * This manual is an updated and modified version of the USDA Misc. Publication no. 1443 published by the Agricultural Research service in 1986 and Edited by George C. Steyskal, William L. Murphy, and Edna M. Hoover. 5.6 - Placing the Labels.......................................................................................................... 47 5.7 - Bar Coding ............................................................................................................ 48 5.8 - Labeling Vials ............................................................................................................ 48 5.9 - Labeling Microscope Slides .......................................................................................... 48 5.10 - Identification Labels .................................................................................................... 48 Part 6 - Care of the Collection ........................................................................................................... 48 6.1 - Housing the Collection .................................................................................................. 48 6.2 - Protecting Specimens From Pests and Mold ................................................................. 49 Part 7 - Packing and Shipping Specimens .......................................................................................... 50 7.1 - Packing Materials. ......................................................................................................... 50 7.2 - Pinned Specimens. ......................................................................................................... 50 7.3 - Specimens in Vials......................................................................................................... 51 7.4 - Loading Cartons. ........................................................................................................... 51 7.5 - Shipping Microscope Slides .......................................................................................... 52 7.6 - Shipping Live Specimens. ............................................................................................. 52 4 Collecting and Preserving Insects and Mites to help identify appropriate areas to be included in reserves. Aquatic species may be used to detect changes in water quality. Pest species may be sampled to assess presence/absence or abundance in order to determine whether control measures are necessary. Specific groups or species may be collected to acquire material for biological, physiological, ecological, molecular, and systematic studies. This manual provides a summary of the methods and techniques used by professionals and amateurs alike to collect and preserve specimens for study. While many of the methods covered here, such as pinning, have changed very little in the last hundred years, other techniques have become available only in the last few years or decades with advancing technologies. Older manuals such as Steyskal et al. (1986), Martin (1977) and Upton (1991) while still useful will not cover such these as preservation for molecular studies. In addition, most of these older publications are now out of print and may be difficult to find. What to Collect Because of their incredible diversity, insects, mites, and other related groups vary widely as to their proper collecting requirements and methods. In the following sections, we will explore some of the many recommended techniques and look at the varied equipment used by collectors. The emphasis will be on insects and mites, but much of what is included here will also pertain to other related groups such as spiders. Which species and how many specimens to collect depends on the purpose for which the material is intended. For hobbyists and students, small samples are usually adequate. However, when important pest insects and mites need to be identified, they should be collected in series if at all feasible. A sample of 20 specimens should be considered the minimum, and even larger numbers may be desirable. If adults and immatures are present, specimens should be col- lected of all life stages. Excess specimens can be discarded or exchanged, but it is not always possible to collect additional speci- mens when needed. Frequently insects and mites cannot be identified accurately from immature stages, and it is then necessary to rear them to the adult stage to obtain a precise identifi- cation. Photographers should collect the specimens they photograph if positive identification is desired; minute, critical diagnostic characters often are not depicted in photographs. If specimens are destined for display cases that portray them in their natural habitats, it may be important to collect a sample of the host plant for the display. Many persons starting a collection attempt to collect every specimen they find. Biology students in high school and college are often required to collect specimens from as many orders or groups as possible. The experience and knowledge gained in making a general collection are of value in helping the collector decide on a specialty. However, with so many different kinds of insects from which to choose—over 100,000 described species in North America alone—most persons find that as their skills and interests increase, concentrating eventually on 1 or 2 of the major insect or mite groups is desirable. Specimens other than those in a chosen group may still be collected for exchange with other collectors. References: Lewis & Taylor 1965; Seber 1973; Barrowclough 1992. Part 1. Equipment and Collecting Methods 1.1 Basic Equipment Collecting methods may be divided into two broad categories. In the first the collector actively searches out Fig. 1. A field collecting kit. 5 Techniques and Tools the insects, using nets, aspirators, beating sheets, or whatever apparatus suits his or her particular needs. In the second, the collector participates passively and permits traps to do the work. Both approaches may be used simultaneously, and both are discussed in the following pages. Using a variety of collecting methods will help to maximize the number of specimens taken, especially when briefly visiting an interesting area. While picking up insects by hand is simple and sometimes effective, their size, mobility, and the possibil- ity of being bitten or stung usually dictates that various kinds of equipment and special methods are needed. Those described here have general application; it is expected that the collector will make some adaptations to fit his or her own purposes and resources. In fact, as experience collecing increases, or the target group becomes more focussed, the use of specialized techniques increases. For additional information, especially concerning the use of specialized techniques, consult the list of references. References (general): Arnett, 1985; Balogh 1958; Banks 1909; Banks et al. 1981; Bland & Jacques 1978; Borror et al.; British Museum 1974; Cantrall 1939-40; Cantrall 1941; Chu 1949; Edmunds & McCafferty 1978; Foote 1948; Klots 1932; Knudsen 1966; Knudsen 1972; Kogan & Herzog 1980; Lehker & Deay 1969; Lincoln & Sheals 1979; McNutt 1976; Martin 1977; Nicholls 1970; Norris 1966; Oldroyd 1958; Peterson 1964; Service 1976; Southwood 1979; Stein 1976; Upton, 1991; USDA 1970; USDA 1966-70; Urquhart 1965; Wagstaffe & Fidler 1955. The equipment used to assemble a general insect or mite collection need not be elaborate or expensive. In many instances, a collecting net (see below) and several killing bottles (see p. 5) will suffice; however, additional items will permit more effective sampling of a particular fauna. Many collectors carry a bag (fig. 1) or wear a vest in which they store equipment. The following items usually are included in the general collector’s bag: (1) Forceps. Fine, lightweight forceps are recommended; if sharp-pointed forceps are used, care must be taken not to puncture specimens. If possible, grasp specimens with the part of the forceps slightly behind the points. (2) Vials containing alcohol or other preservatives (see p. 21). (3) Killing bottles of various sizes. (4) Small boxes or containers for storing specimens after their removal from killing bottles. These may be made of cardboard, plastic, or metal and should be partly filled with soft tissue or cloth to keep specimens from rolling about. Do not use cotton because specimens become entangled in the fibers and may become virtually impossible to extricate without damage. (5) Small envelopes for temporary storage of delicate specimens and/or gelcaps for tiny specimens. (6) One or more aspirators (see p. 7-8). (7) Absorbent tissue for use in killing bottles and aspira- tors. (8) Notebook and writing equipment for jotting down notes and label data. (9) A strong knife for opening galls, seed pods, twigs, etc and a pair of scissors for cutting labels. (10) A small, fine brush (camel’s hair is best) for picking up minute specimens. Moisten the tip; tiny specimens will adhere to it and may be transferred to a killing bottle or vial. (11) Bags for storing plant material, rearing material, or Berlese samples. For collecting much plant material, a botanist’s vasculum or tin box is advisable. (12) A hand lens. This list may be modified according to the special kinds of insects or mites to be collected. A small digging tool or trowel may be useful for collecting insects from soil or for gathering Berlese samples and a heavy knife or small hatchet for searching under bark or in decaying logs. A plant press should be available to prepare plant speci- mens for determination or as voucher specimens, espe- cially when leaf-mining insects are being studied. When collecting at night, have a flashlight or headlamp; the latter is especially useful because it leaves the hands free. Much of the equipment listed above may be obtained from around the home or from ordinary sources like a drug store, but equipment especially designed for insect collecting often must be bought from special supply houses. If there is a local company, their address may usually be found in the yellow pages of telephone directo- ries under “Biological Laboratory Supplies” or “Labora- tory Equipment and Supplies.” The faculty members of a local university’s biology or entomology department or curators at a nearby museum are usually willing to help and in the best position to recommend a supplier in the area. Professional journals also sometimes carry advertise- ments for equipment suppliers. 1.2 Collecting Nets Collecting nets come in three basic forms: Aerial, sweeping, and aquatic. The first is designed especially for 6 Collecting and Preserving Insects and Mites collecting butterflies and other flying insects. Both the bag and handle are relatively lightweight. The sweeping net is similar to the aerial net but is stronger and has a more durable bag to withstand being dragged through dense vegetation. Aquatic nets are used for gathering insects from water and are usually made of metal screening or heavy scrim with a canvas band affixed to a metal rim. A metal handle is advisable because wooden ones may deteriorate after repeated wetting. The net you choose depends on the kind of insects or mites you wish to collect. Several kinds of nets, including collapsible models with interchangeable bags, are available from biological supply houses, but anyone with a little mechanical ability can make a useful net. The advantage of a homemade net is that the size and shape can be adapted to the needs of the user, to the kind of collecting intended, and to the material available, which need not be expensive. These materials include— (1) Piece of heavy (8- gage) steel wire for the rim, bent to form a ring 30-38 cm in diameter (fig. 2, A). Small nets 15 cm or so in diameter sometimes are useful, but nets larger than 38 cm are too cumbersome for most collect- ing. (2) Dacron or other strong, light fabric through which air can flow freely. Brussels netting is best but may be difficult to obtain; otherwise nylon netting, marquisette, organdy, or good quality cheesecloth can be used, but the last snags easily and is not durable. The material should be folded double and should be 1.5-1.75 times the rim diameter in length (fig. 2, B). The edges should be double-stitched (French seams). (3) Strip of muslin, light canvas, or other tightly woven cloth long enough to encircle the rim. The open top of the net bag is sewn between the folded edges of this band to form a tube through which the wire rim is inserted (fig. 2, C). (4) Straight hardwood dowel about 19 mm in diameter and 105-140 cm long (to suit the collector). For attachment of the rim to the handle, a pair of holes of the same diam- eter as the wire are drilled opposite each other to receive the bent tips of the wire, and a pair of Fig. 2. Collecting Net Fig. 3. A truck equipped with a large net. 7 Techniques and Tools grooves as deep and as wide as the wire are cut from each hole to the end of the dowel to receive the straight part of the wire (fig. 2, D). (5) Tape or wire to lash the ends of the rims tightly into the grooves in the end of the handle. This may be electrician’s plastic tape or fiber strapping tape commonly used for packaging. If wire is used, the ends should be bound with tape to secure them and to keep them from snagging. A close-fitting metal sleeve (ferrule) may be slipped over the rim ends and held in place with a small roundheaded screw instead of tape or wire lashing. After the net has been placed on the rim, the ends of the band should be sewn together and the rim ends fastened to the handle. The other end of the handle should be filed to remove sharp edges. The net is then ready for use (fig. 2, E). Efficient use of a net is gained only with experience. Collection of specimens in flight calls for the basic stroke—swing the net rapidly to capture the specimen, then follow through to force the insect into the very bottom of the bag. Twist the wrist as you follow through so the bottom of the bag hangs over the rim; this will entrap the specimen. If the insect alights on the ground or other surface, it may be easier to use a downward stroke, quickly swinging down on top of the insect. With the rim of the net in contact with the ground to prevent the specimen from escaping, hold the tip of the bag up with one hand. Most insects will fly or crawl upward into the tip of the bag, which can then be flipped over the rim to entrap the specimen. Sweeping the net through vegetation, along the sand and seaweed on beaches, or up and down tree trunks will catch many kinds of insects and mites. The aerial net may be used in this way, but the more durable sweeping net is recommended for such rough usage. After sweeping with the net, a strong swing will bring anything in the bag to the bottom, and then by immediately grasping the middle of the net with the free hand, the catch will be confined to a small part of the bag. Only the most rugged sweeping net may be used through thistles or brambles. Even some kinds of grasses, such as sawgrass, can quickly ruin a net. Burs and sticky seeds are also a serious problem. The catch may be transferred from the bag to a killing jar in one of several ways. Single specimens are transferred most easily by lightly holding them in a fold of the net with one hand while inserting the open killing jar into the net with the other. While the jar is still in the net, cover the opening until the specimen is stupefied; other- wise, it may escape before the jar can be removed from the net and closed. To prevent a butterfly from damaging its wings by fluttering in the net, squeeze the thorax gently through the netting when the butterfly’s wings are closed. Experience will teach you how much pressure to exert; obviously, pinching small specimens of any kind is not recommended. When numerous specimens are in the net after prolonged sweeping, it may be desirable to put the entire tip of the bag into a large killing jar for a few minutes to stun the insects. They may then be removed and desired specimens placed separately into a killing jar, or the entire mass may be dumped into a killing jar for later sorting. These methods of mass collecting are especially adapted to obtaining small insects not readily recognizable until the catch is sorted under a microscope. Removal of stinging insects from a net may be a problem. They will often crawl toward the rim of the bag and may be made to enter a killing jar held at the point where they crawl over the rim. However, many insects will fly as soon as they reach the rim, and a desired specimen may be lost. A useful method is to trap the insect in a fold of the net, carefully keeping a sufficient amount of netting between fingers and insect to avoid being stung. This fold of the net can then be inserted into the killing jar to stun the insect. After a few moments, it should be safe to remove the insect from the net and transfer it to a killing jar. If the stunned insect clings to the net and does not fall readily into the jar, use forceps or pry the insect loose with the jar lid or a small stick—not with your fingers. Aerial nets made of dacron or nylon may be used to sweep insects from water if an aquatic net is not at hand. The netting will dry quickly if swept strongly through the air a few times; however, it should not be used again until thoroughly dry, or other specimens, especially butterflies, may be ruined. A number of special modifications are necessary to adapt a net for aquatic collecting. For specialized collecting, nets can be attached to the ends of beams that are rotated about their midlength by a motor drive. Nets also can be adapted to be towed by or mounted on vehicles (fig. 3) (Peck and Cook, 1992). References: Dresner 1970; Johnson 1950; Rogers & Smith 1977; Rudd & Jensen 1977; Takeda et al. 1962; Williams & Miine 1935; vehicle-mounted net: Almand et al. 1974; Barnard 1979; Grigarick 1959; Harwood 1961; Hill 1971; Holzapfel et al. 1978; Kronblad & Lundberg 1978; Landin 1976; McNutt 1976; Noyes, 1982; Rudd & Jensen 1977; Torre-Bueno 1937; Traver 1940. 1.3 Killing Jars or Bottles Effective collecting of insects and related groups usually requires that the specimens be killed so that they may be properly mounted and studied. The most widely 8 Collecting and Preserving Insects and Mites employed method for killing collected specimens is the killing jar (bottle). Any heavy, wide-mouthed glass jar or bottle with a tight-fitting stopper or metal screw top may be used. Tops that may be removed with only a quarter turn often are preferred but may not be obtained readily. The killing agent used may be any of various liquids or solids. Liquid killing agents generally are considered to be slower acting but safer to use than solids such as cyanide, but some of them are known to accumulate in human tissue after repeated or prolonged exposure. Despite its extreme toxicity, cyanide is a noncumulative poison, and brief exposure to the fumes, as inevitably occurs when opening jars to insert or remove specimens, is not believed to reault in any permanent harm. Never deliberately inhale the fumes, even momentarily. All killing agents are to some extent hazardous to human health. All killing jars or bottles should be clearly labeled “POISON” and should be kept away from children or persons who may be unaware of their potential danger. When not in use, killing jars should be stored in a safe place away from children and pets where they are not liable to accidental breakage. Cyanide jars should not be stored in an area such as a bedroom where any accidental leakage could expose someone to fumes. Remember that killing agents can be as effective against humans as they are against insects and that care and caution in their construction and use are essential. 1.4 Liquid Killing Agents Jars for use with liquid killing agents are prepared in one of two ways. One way (fig. 4, A) is to pour about 2.5 cm of plaster of paris mixed with water into the bottom of the jar and allow the plaster to dry. Enough of the killing agent is then added to saturate the plaster; any excess should be poured off. This kind of jar can be recharged merely by adding more killing agent. The second method is to place a wad of cotton or other absorbent material in the bottom of a jar, pour enough liquid killing agent into the jar to nearly saturate the absorbent material, and then press a piece of stiff paper on it or a cardboard cut to fit the inside of the jar tightly. The paper or cardboard acts as a barrier between the insect and the killing agent, keeping the latter from evaporating too rapidly and also preventing the specimen from becoming entangled in loose fibers. Among the liquid killing agents are ethyl acetate (CH3CO2 • C2H5), ether (diethyl ether, C2H5 • O • C2H5), chloroform (CHCI3), and ammonia water (NH4OH solution). Ethyl acetate is most widely used. All of these chemicals are extremely volatile and flammable and should never be used near fire. Children should only use them under adult supervision. Ethyl acetate is regarded by many as the most satisfactory liquid killing agent. Its fumes are less toxic to humans than those of the other substances. Although it usually stuns insects quickly, it kills them slowly. Speci- mens that appear dead may revive if removed from the killing jar too soon, but a compensating advantage is that most specimens may be left in an ethyl acetate killing jar for several days and still be limp. If the ethyl acetate is allowed to evaporate from the specimens, they will harden. Killing jars with ethyl acetate are preferred by many entomologists, especially for infrequent use. Ether and chloroform are both extremely volatile and Fig. 4. Killing jars 9 Techniques and Tools flammable and should not be used near an open flame or lighted cigarette. Their high volatility makes them service- able in a killing jar for only a short time. Perhaps the greatest hazard with chloroform is that even when stored in a dark-colored jar, it eventually forms the extremely toxic gas phosgene (carbonyl chloride, COCI2). Chloro- form, however, is useful when other substances cannot be obtained. It stuns and kills quickly but has the disadvan- tage of stiffening specimens. Ethyl Alcohol (ethanol or ETOH) is widely used to kill small Coleoptera adults, small Hymenoptera, and many immature insects and soft-bodied insects. It is most commonly used at 70-80% concentration and many workers add 5% glacial acetic acid ("acetic alcohol") which helps penetration of the alcohol into the specimen and leaves specimens more relaxed. Isopropyl alcohol (rubbing alcohol) may also be used, and may be easier to find and purchase than Ethanol. However, Ethanol is preferred for most applications. Ethanol is used com- monly in Berlese funnels and similar traps. Liquid ammonia is irritating to humans, and in general is not a particularly effective killing agent for most insects. However, it is highly recommended for use in small vials for dispatching microlepidoptera, and it has been used with variable success in blacklight traps, again for Lepidoptera. Specimens killed in ammonia tend to stay in a relaxed condition much longer than those killed by cyanide, allowing greater ease of spreading. Ammonia is readily available from many sources. Ammonium carbonate, a solid but volatile substance, also can be used. 1.5 Solid Killing Agents The solid killing agents most often used in killing jars are the cyanides—potassium cyanide (KCN), sodium cyanide (NaCN), or calcium cyanide [Ca(CN)2]. Handle all cyanides with extreme care. They are dangerous, rapid- acting poisons with no known antidote. If even a single grain touches the skin, wash immediately with water. To avoid handling the cyanide and having to find a safe place to store or dispose of surplus crystals, you may be able to find a chemist, pharmacist, or professional entomologist to make the killing jar for you. If this is not feasible, use utmost care in following the instructions given here. To make a cyanide killing jar or bottle, place a layer (about 15 mm) of cyanide crystals in the bottom (fig. 4, B). Potassium cyanide is best; sodium cyanide is as effective but is hygroscopic, that is, it absorbs water and makes the jar wet; and calcium cyanide is seldom avail- able. Cover the crystals with about 10 mm of sawdust and then add about 7 mm of plaster of paris mixed with water to form a thick paste, working quickly before the plaster solidifies. Then add crumpled absorbent paper to prevent water condensation on the inside glass surface. Instead of the plaster of paris, a plug of paper or cardboard may be pressed on top of the sawdust. Be sure that it fits tightly. When ready to use after a few hours, place several drops of water on the plaster or paper plug. In an hour or so, enough fumes of hydrocyanic acid will have been pro- duced to make the jar operative. Do not test this by sniffing the open jar. Every killing jar or bottle should be clearly and prominently labeled “POISON”. The bottom must be covered with tape, preferably cloth, plastic, or clinical adhesive tape, to cushion the glass against breakage and to keep its dangerous contents from being scattered if the container breaks. Killing jars or bottles will last longer and give better results if the following simple rules are observed: (1) Place a few narrow strips of absorbent paper in each jar or bottle to keep it dry and to prevent specimens from mutilating or soiling each other. Replace the strips when they become moist or dirty. This method is useful for most insects except Lepidoptera, which are too difficult to disentangle without damage. (2) Do not leave killing jars in direct sunlight as they will sweat and rapidly lose their killing power. (3) If moisture condenses in a jar, wipe it dry with absorbent tissue. (4) Keep delicate specimens in separate jars so that larger specimens will not damage them. (5) Do not allow a large number of specimens to accumulate in a jar unless it is to be used specifically for temporary storage. (6) Do not leave insects in cyanide jars for more than a few hours. The fumes will change the colors of some insects, especially yellows to red, and specimens will generally become brittle and difficult to handle. (7) If it is necessary to keep insects in killing jars for more than several hours, place the specimens in another container and store them in a refrigerator. (8) Keep butterflies and moths in jars by themselves so that their hairs and scales will not ruin other kinds of insects. (9) Never test a killing jar by smelling its contents. (10) Old jars that no longer kill quickly should be recharged or disposed of by burning or burying. A cyanide 10 Collecting and Preserving Insects and Mites jar that has become dry may be reactivated by adding a few drops of water. Spray-dispensed insecticides may be used, if not to kill specimens, to at least ‘knock them down’ into a container from which they may be picked up. If they are directed into a container topped with a funnel, they may be allowed to revive and treated further as desired (see Clark & Blom 1979). References: Banks et al. 1981; Clark & Blom 1979; Frost 1958; Lindroth 1957; Pennington 1967; Preiss et al.; White 1964. 1.6 Aspirators and Suction Devices The aspirator (fig. 5, A), known in England as a ‘pooter,’ is a convenient and effective device for collecting small insects and mites. The following materials are needed to construct an aspirator: (1) Vial 2.5-5 cm in diameter and about 12 cm long. (2) Two pieces of glass or copper tubing about 7 mm in diameter, one piece about 8 cm long and the other about 13 cm long. (3) Rubber stopper with two holes in which the tubing will fit snugly. (4) Piece of flexible rubber or plastic tubing about 1 meter long, with diameter just large enough to fit snugly over one end of shorter piece of stiff tubing. (5) Small piece of cloth mesh, such as cheesecloth, and rubberband. To make an aspirator, bend the glass or copper tubes as in figure 5, A. In bending or cutting glass tubes, always protect your fingers by holding the glass between several layers of cloth. Obtain the advice of a chemist or labora- tory technician for cutting and bending glass. Moisten one end of the longer tube and insert it through one of the holes in the rubber stopper. Moisten one end of the shorter tube, insert it through the other hole in the stopper, and using a rubberband fasten the cloth mesh over the end that was inserted through the stopper; this will prevent speci- mens from being sucked into the collector’s mouth when the aspirator is used. Attach one end of the flexible tubing to the free end of this tube. The length, size, and amount of bend in the tubing will vary according to the user’s needs. To complete the assembly, insert the rubber stopper into the vial. To use the aspirator, place the free end of the flexible tubing in the mouth, move the end of the longer glass tube close to a small specimen, and suck sharply. The specimen will be pulled into the vial. Instead of using a vial, some workers prefer a tube (fig. 5, B). In either method, it is well to keep small pieces of absorbent tissue in the vial or tube at all times to prevent moisture from accumulating. Be cautioned that there is some danger of inhaling harmful substances or organisms when using a suction-type aspirator (see Hurd 1954). Either the vial- or tubing-type aspirator (fig. 5, B) may be converted into a blow-type aspirator by removing the 13-cm glass tube (see fig. 5, A) and substituting a T— shaped attachment (fig. 5, B). The flexible tubing is attached to one arm of the ‘T,’ the opposite arm is left open, and the stem of the ‘T’ is inserted into the vial and covered with mesh. Upon blowing through the flexible tubing, a current of air passes across the ‘T’ and creates a partial vacuum in the vial, which produces the suction needed to draw specimens into the vial. This kind of aspirator eliminates the danger of inhaling small particles, fungus spores, or noxious fumes. Aspirators with a squeeze bulb may sometimes be purchased, or if a valved bulb can be obtained, they may be constructed for use with either pressure or suction. Collection traps also have been devised with the suction feature applied on a much larger scale than with the usual aspirator. Suction produced by a fan has been employed in traps in conjunction with light or other attractants. Some of these traps are described in the following references and in the section on Traps. Suction is created by a piston in a ‘slurp-gun’ described for aquatic collecting. This principle could be adapted for use in air to gather insects and to deposit them in a vial attached to the side of the piston. References: Azrang 1976; Barnard & Mulla 1977; Fig. 5. Aspirators 11 Techniques and Tools Bradbury & Morrison 1975; Clifford et al. 1977; Evans et al. 1964; Galtsoff et al. 1937; Hurd 1954; Johnson 1950; Johnson & Taylor 1955; Johnson et al. 1957; Lumsden 1958; Minter 1961; Mulhern 1942; Scholdt & Neri 1974; Taylor 1962a; Turnbull & Nicholls 1966; White 1964; Weins & Burgess 1972; Williams 1973; Woke 1955. 1.7 Beating Sheets A beating sheet should be made of durable cloth, preferably white, attached to a frame about 1 meter square, with two pieces of doweling or other light wood crossing each other and fitted into pockets at each corner of the cloth. An ordinary light-colored umbrella also may be used as a beating sheet. Place the beating sheet or umbrella under a tree or shrub and sharply beat the branches or foliage with a club or stick. Specimens will fall onto the sheet and may be removed from the light-colored material by hand or with forceps, a moistened brush, or an aspira- tor. Locating specimens on the sheet is sometimes a problem because of leaves or other unwanted material dropping onto the sheet. Watching for movement will help locate specimens, as well as tilting the sheet so that the debris is displaced or even allowed to fall off, with the insects and mites left clinging to the cloth. Beating sheets are especially useful in collecting beetles, true bugs, and larval Lepidoptera. Beating may be the best collecting technique when the weather has turned cold, or early and late in the day, when normally active insects seek shelter in vegetation and are otherwise difficult to detect. A ‘ground cloth’ also is used in sampling crop fields (see Rudd & Jensen 1977). 1.8 Sifters Sifters are used to collect insects and mites that live in ground litter, leaf mold, rotting wood, mammal and bird nests, fungi, shore detritus, lichens, mosses, and similar material. Sifters are especially useful for winter collecting to pick up hibernating specimens. Almost any container with a wire-mesh screen bottom will serve as a sifter. The size of the mesh depends on the size of the specimens sought. For general purposes, screening with 2.5-3 meshes per centimeter is satisfactory. To use the sifter, place the material to be sifted into the container and shake it gently over a white pan or piece of white cloth. As the insects and mites fall onto the cloth, they may be collected with forceps, a brush, or an aspirator. A similar method is used chiefly to collect mites from foliage. Using a sifter of 20-mesh screen (about 8 per centimeter) with a funnel underneath that leads to a small vial, beat pieces of vegetation against the screen to dislodge the mites, which will fall through the screen and into the vial below. Another type of sifter employs two hoops of heavy metal, each with a handle. A long (3-4 ft.) canvas bag is sewn to the top hoop. The bag is left open at the end and secured with a cord or twist-tie. About 1 foot down in the bag, the second hoop is sewn to the canvas and to this is attached a metal screen. Coarse debris is loaded into the top and sifted down to the end of the canvas bag. Sifted debris is then ready to be processed by one of the follow- ing separators or extractors. References: Martin 1977. 1.9 Separators and Extractors Somewhat similar to the sifter are various devices designed to separate or extract live specimens from substances in which they may be found, such as leaf mold and other kinds of vegetable matter, shore detritus, dung, even net sweepings that include so much foreign matter that it is difficult to pick out the insects. These devices usually depend on some physical aid such as light, heat, or dryness to impel the insects to leave the foreign matter. One of the simplest such devices is the sweeping separator (fig. 7). This is simply a carton or wooden box with a tight-fitting lid. Near the top of the box on one side is inserted a glass jar. If the jar is made with a screw top, a hole of proper diameter cut in the side of the carton will permit the jar to be screwed onto it. The cover ring, Fig. 6. Separation bag 12 Collecting and Preserving Insects and Mites without the lid, from a home-canning jar may be nailed to the periphery of a hole in a wooden box and the jar then screwed onto the ring. The sweepings are dumped into the box and the cover is quickly closed. The insects in the darkened box soon will be attracted to the lighted glass jar. When all the insects appear to have entered the jar, it can be removed and its contents put into a killing jar. Alternatively, a jar cover containing a piece of blotting paper soaked with xylene may be placed over the jar for awhile to stun the insects, which may then be sorted. A more sophisiticated version of this separtor is made of alumnium with a clear plastic top and cloth collecting bag (fig. 6). Sitting on three legs, this separator allow the collector to dump the catch into the bag, place the lid, which is lined with a magnet, on top and insert an aspirator through a small hole in the side of the bag. As insects are attracted to the top and collect on the plastic, the aspirator can be moved about to suck up the insects of interest. Nets can also be modified to help keep plant material contacted during sweeping away from the insects. In most cases, hardware cloth or some other screening material with fairly large holes (ca. 1cm in diameter) is placed across the net opening and fastened to the net ring. This works well to keep out larger pieces of plant debris but will not be effective in excluding seeds and other small plant parts (Noyes 1988; Zolnerowich et al. 1990). Insects collected into alcohol can also be separated from plant debris by the use of screens. In this method, a screen of 1/4 inch diameter galvanized hardware cloth is fastened over a frame. Below this is another screen made from a very fine mesh material such as organdy or a small section of panty hose. The insect/plant material collection is poured over the coarser screen and alcohol is added. When agitated, the insect will sink through the larger screen while plant material will float or be stopped by the screen mesh. A similar method uses a set of three stacked screens of decreasing diameter and specimens are washed from one layer to another using a gentle spray of water. Care must be taken so that the washing does not damage the specimens. The Berlese or Tullgren funnel (Upton 1991) (fig. 8) and its modifications are cleaner and more efficient than sifting to separate insects and mites from leaf mold and similar materials. The sample (usually presifted to remove large debris) is placed on a screen near the top of a funnel. A light bulb can be placed above the sample to produce heat and light, which drive the insects downward into the funnel, or heated coils or a jacket around the funnel can be used to dry the sample and make it inhospitable. The insects and mites are directed by the funnel into a con- tainer, sometimes containing alcohol at the bottom of the funnel. Care should be taken not to dry the sample so rapidly that slow-moving specimens are immobilized before they can leave the sample. To prevent large amounts of debris from falling into the container, place the sample on the screen before the container is put in place. A similar separator is the photoeclector or Winkler/Moczarski Elector. This device is similar to the Berlese funnel except that no light bulb or other heat source is used to drive the insect to the bottom. Instead, an open jar, with a most cloth or tissue inside, is attached to the bottom of the funnel or canvas bag and insects are Fig. 7. Sweeping separator Fig. 8. Berlese funnel 13 Techniques and Tools attracted to the light and humidity. An added advantage of this device is that it requires no electricity and so may be more readily used in the field. References: Besuchet et al. 1987; Brown 1973; Everett & Lancaster 1968; Finch & Skinner 1974; Gui et al. 1942; Kempson et al. 1962; Kevan 1955; Kevan 1962; Lane & Anderson 1976; Martin 1977; Masner & Gibson 1979; Murphy 1962; Newell 1955; Norton and Kethley 1988; Salmon 1946. 1.10 Traps Since a trap is defined as anything that impedes or stops the progress of an organism, this subject is extensive, including devices used with or without baits, lures, or other attractants. Besides its construction, the performance of a trap depends on such factors as its location, time of year or day, weather, temperature, and kind of attractant used, if any. A little ingenuity coupled with knowledge of the habits of the insects or mites sought will suggest modifications or improvements in nearly any trap or may even suggest new traps. Only a few of the most useful traps are discussed here, but the following references describe many more, especially Martin 1977; Peterson 1964; Southwood 1979. References: A'Brook 1973; Banks 1959; Banks et al. 1981; Barber 1931; Bidlingmayer 1967; Broadbent 1949; Broadbent et al. 1948; Dunn & Reeves 1980; Evans 1975; Flaschka & Floyd 1969; Ford 1973; Glasgow & Duffy 1961; Golmeric & Davenport 1971; Granger 1970; Hafraoui et al. 1980; Hanec & Bracken 1964; Hansens et al. 1971; Hargrove 1977; Hartstack et al. 1968; Hathaway 1981; Heathcote et al. 1969; Hienton 1974; Hollingsworth et al. 1963; Howell 1980; Kimerle & Anderson 1967; Klein et al. 1973; Martin 1977; Meyerdirk et al. 1979; Morris 1961; Peterson 1964; Pickens et al. 1972; Southwood 1979; Sparks et al. 1980; Taylor 1962b; Thorsteinson et al. 1965; Weseloh 1974; Whittaker 1952; Williams 1951; Woke 1955. 1.10.1 Effects of Elevation One of the external factors affecting the performance of traps, especially light traps, has been specially studied, namely the effect of the elevation (above sea or ground level) at which the trap is placed when in use. The subject is complex, with many variables related to kinds of insects, locality, and so forth, which are discussed in the following references. References: Blakeslee et al. 1959; Callahan et al. 1972; Cooke 1969; Frost 1957; Glick 1939; Glick 1957; Goma 1965; Meyers 1959; Roling & Kearby 1975; Stewart & Lam 1968. 1.10.2 Windowpane Traps One of the simplest and cheapest traps is a barrier consisting of a windowpane held upright by stakes in the ground or suspended by a line from a tree or from a horizontal line. A trough filled with a liquid killing agent is so placed that insects flying into the pane drop into the trough and drown. They are removed from the liquid, washed with alcohol or other solvent, then preserved in alcohol or dried and pinned. The trap is not recommended for adult Lepidoptera or other insects that may be ruined if collected in fluid. A modification of this trap uses the central "pane" of a malaise trap instead of a pane of glass. The malaise trap pane covers more space than glass, is easier to transport, and, of course, is not breakable. Various mesh sizes if cloth can also be used depending on the insects targeted. These traps may also be referred to as flight intercept traps. References: Chapman & Kinghorn 1955; Corbet 1965; Kato et al. 1966; Lehker & Deay 1969; Masner and Goulet 1981; Nijholt & Chapman 1968; Peck and Davies 1980; Roling & Kearby 1975; Wilson 1969. 1.10.3 Interceptions Nets and Barriers A piece of netting, 1.8 meters or more in height, can be stretched between three trees or poles to form a V- shaped trap with the wide end of the V open. A triangular roof should be adjusted to slope gently downward to the broad open side of the V. A device of this type will intercept many kinds of flying insects, particularly if the trap is situated with the point of the V toward the side of maximum light and in the direction of air movement. A pair of such nets set in opposite directions, or a single net in a zigzag shape, will intercept specimens from two directions. Since insects flying into such a net tend to gather at the pyramidal apex, they are easy to collect. In one variant of this trap the cloth is sprayed with a sythetic pyrethroid insecticide and the insects which are killed by contact with the cloth then fall into a long pan trap at the bottom. The so-called ‘funnel’ or ‘ramp’ traps are inter- ception devices that direct insects to a central point, where a retaining device or killing jar may be placed. More complex arrangements have been described in the litera- ture, primarily for migrating butterflies. References: Gillies 1969; Graham et al. 1961; Hocking & Hudson 1974; Jonasson 1954; Leech 1955; Masner and Goulet 1981; Merrill & Skelly 1968; Nielsen 1960; Parman 1931, 1932; Steyskal 1981; Walker and Lenczewski 1989; Walker and Whitesell 1993, 1994. 14 Collecting and Preserving Insects and Mites 1.10.4 Malaise Traps One of the most widely used insect traps was developed by the Swedish entomologist René Malaise and that now bears his name. Several modifications of his original design have been published, and at least one is available commercially. The trap, as originally designed, consists of a vertical net serving as a baffle, end nets, and a sloping canopy leading up to a collecting device (fig. 9). The collecting device may be a jar with either a solid or evaporating killing agent or a liquid in which the insects drown. The original design is unidirectional or bidirec- tional with the baffle in the middle, but more recent types include a nondirectional type with cross baffles and with the collecting device in the center. Malaise traps have been phenomenally successful, sometimes collecting large numbers of species that could not be obtained otherwise. Attractants may be used to increase the efficiency of the traps for special purposes. References: Butler 1966; Townes 1972; Steyskal 1981 (bibliography). 1.10.5 Pitfall and Dish Traps Another simple but very effective and useful type of interception trap consists of a jar, can, or dish sunk in the earth (fig. 10). A cover must be placed over the open top of the jar to exclude rain and small vertebrates while allowing insects and mites to enter. A piece of bark, wood, or flat stone will serve this purpose. Pitfall traps may be baited with various substances, depending on the kind of insects or mites the collector hopes to capture. Although most that fall into the trap will remain there, it should be inspected daily, if possible, and desired specimens removed and placed in alcohol or in a killing bottle while they are in their best condition. Also in the pitfall category is the cereal dish trap, which is a simple but effective device for obtaining insects attracted to dung. It consists of a small dish, preferably Figs. 10-11. 10, Pitfall Trap (Top). 11, Cereal Dish Trap (Bottom). Fig. 9. Malaise trap. 15 Techniques and Tools with a rim, set in the earth (fig. 11) and partly filled with 70 percent ethanol, or, if available, with ethylene glycol, which does not evaporate. A piece of stout wire, such as a coathanger, is bent as shown, with a loop at one end to hold the bait receptacle. A few zigzag bends in the other end of the wire will keep the looped end from swinging after the wire is pushed into the earth. The bait receptacle may be a small plastic or metal cup such as is often used for medicine doses, or a coffee creamer, or a cup formed from aluminum foil. When baited with animal or human feces, this trap attracts beetles, mostly of the families Scarabaeidae and Staphylinidae, springtails, ants, earwigs, some parasitic Hymenoptera, and, rather surprisingly, several families of flies, especially Phoridae, Sepsidae, and Muscidae. The larger, strong-flying calliphorid and sarcophagid flies seldom fall into the liquid, although they are attracted to the bait. The alcohol fumes probably cause the smaller flies to drop into it. The trap is made of easily obtained materials, is easily transported, and provides excellent results. It deserves wide use. References: Adlerz 1971; Barber 1931; Beaudry 1954; Briggs 1971; Clark and Blom 1992; Dethier 1955; Fichter 1941; Gist & Crossley 1973; Golding 1941; Greenslade 1973; Greenslade & Greenslade 1971; Greenslade 1964; Gressitt et al. 1961; Grigarick 1959; Heathcote 1957; Houseweart et al. 1979; Joosse 1975; Loschiavo 1974; Luff 1968, 1975; Masner & Huggert 1979; Morrill 1975 (bibliography); Muma 1975; Newton & Peck 1975; Reeves 1980; Schmid et al. 1973; Shubeck 1976; Smith 1976; Smith et al. 1977; Thomas & Sleeper 1977; Tretzel 1955; Van den Berghe 1992; Welch 1964. 1.10.6 Moericke Traps and Other Color Traps Moericke traps or yellow pan traps are used extensively by some collectors. An aluminum or plastic pan is painted yellow and placed on the ground (or a depression may be dug and the pan set in the depres- sion) and filled about 1/3 full with salt water, or some other non-toxic fluid. A few drops of detergent of some other surfactant is added to the water to break the surface tension. Insects attracted to the pan fall into the fluid and perish. The trap is then strained periodically (one favorite strainer is a small aquarium fish net). Pan traps such as this, are often placed under malaise traps and flight interception traps to catch insects that may hit the trap and fall to the ground. Yellow seems to be the best color for traps, but various kinds of insects react differently to different colors. Some recent research indicates that certain parasitic wasps respond most strongly to blue. Colored sticky traps are also used to sample insects in various habitats. One of these, the Manitoba trap (fig. 15) has a black sphere to attract horse flies (family Tabanidae), which are then captured in a canopy-type trap. References: Beroza 1972; Granger 1970; Gurney et al. 1964; Hottes 1951; Kieckhefer et al. 1976; Kring 1970; Marshall, 1994; Moericke 1951, 1955 (in german); Prokopy 1973; Weseloh, 1986. 1.10.7 Emergence and Rearing Traps An emergence trap is any device that prevents adult insects from dispersing when they emerge from their immature stages in any substrate, such as soil, plant tissue, or water. A simple canopy over an area of soil, over a plant infested with larvae, or over a section of stream or other water area containing immature stages of midges, may- flies, and other arthropods will secure the emerging adults. If it is equipped with a retaining device, as in the Malaise trap, the adults can be killed and preserved shortly after emergence. It must be remembered, however, that many insects should not be killed too soon after emergence because the adults are often teneral or soft bodied and incompletely pigmented and must be kept alive until the body and wings completely harden and colors develop fully. Emergence traps and rearing cages (fig. 12) enable Fig. 12. Emergence and rearing traps. 16 Collecting and Preserving Insects and Mites the insects to develop naturally while insuring their capture when they mature or when larvae emerge to pupate. References: Adkins 1972; Akar and Osgood 1987; Banks et al. 1981; Barber & Mathews 1979; Butler 1966; Catts 1970; Cheng 1975; Coon & Pepper 1968; Davidson & Swan 1933; Debolt et al. 1975; Doane 1961; Gerking 1957; Glen 1976; Harwood & Areekul 1957; Hollis 1980; Kimerle & Anderson 1967; Krombein 1967; LaGasa & Smith 1978; Lammers 1977; Langford & Daffern 1975; Levin 1957; Lindeberg 1958; Macan 1964; McCauley 1976; Masteller 1977; Merritt & Poorbaugh 1975; Morgan et al. 1963; Morrill & Whitcomb 1972; Mundie 1956, 1964, 1966, 1971; Murray & Charles 1975; Needham 1937; Nielson 1974; Smith et al. 1977; Thompson & Gregg 1974; Turnock 1957; Yates 1974. 1.10.8 Lobster or Eel Trap This category includes any container that has its open end fitted with a truncated cone directed inward, as in a lobster or eel trap, known as a ‘Reuse’ in German. An ordinary killing jar with a funnel fastened into its open end is an example. When the funnel is placed over an insect, the specimen will usually crawl or fly toward the light and enter the jar through the funnel. Modified traps of this type include the Steiner and McPhail traps, which are used primarily in fruit fly surveys but are suitable for many other purposes. The inside of the Steiner trap usually has a sticky material containing a pheromone or other lure. Both traps, as well as similar devices, may be used with different attractants to collect diverse kinds of insects. References: Bellamy & Reeves 1952; Broce et al. 1977; Brockway et al. 1962; Doane 1961; Hollis 1980; Jacobson & Beroza 1964; Morrill & Whitcomb 1972; Nakagawa et al. 1975; Nicholls 1960; Nielson 1974; Reierson & Wagner 1975; Steyskal 1977. 1.10.9 Light Traps With light traps, advantage is taken of the attraction of many insects to a light source. Using various wave- lengths as the attractant, a great variety of traps can be devised, a few of which are described here. Many traps can be constructed easily from materials generally available around the home. All wiring and electrical connections should be approved for outdoor use. Funnels can be made of metal, plastic, or heavy paper. Traps can be used with or without a cover, but if they are to be operated for several nights, covers should be in- stalled to keep out rain. The New Jersey trap (fig. 13) includes a motorized fan to force insects attracted to the light into a killing jar. It has been especially useful for collecting small, non-scaly insects such as midges and gnats. This type of light trap, in which the insects fall directly into a killing jar, is not recommended for use with moths because such delicate Fig. 13. New Jersey Trap Fig. 14. Wilkinson Trap 17 Techniques and Tools specimens may be badly rubbed or torn. If only small insects are desired, they may be protected from damage by larger insects by placing a screen with the proper sized mesh over the entrance. The Minnesota trap is very similar to the New Jersey trap, but it does not include a fan or any motorized method of draft induction. The Wilkinson trap (fig. 14) requires somewhat more effort to construct than the preceding traps, but it has the advantage of confining, not killing, the trapped insects. Moths, therefore, can be collected in good condition if the trap is inspected frequently and desirable specimens are removed quickly through the hinged top and placed in a killing jar. Several highly effective but more elaborate devices have been made for collecting moths and other fragile insects in good condition. Basically, they all use the principle of a funnel with a central light source above it and vanes or baffles to intercept the approaching insects that are dropped through the funnel into the container beneath, which may or may not hold a killing agent. The nature of the container and the type of killing agent affect the quality of the specimens obtained. Some traps catch the insects alive in a large collection chamber, such as a garbage can, which is filled or nearly filled with loosely arranged egg cartons. Most moths will come to rest in the cavities between the egg cartons and will remain there until removed in the morning. Other traps are designed to kill the insects by means of high concentrations of fumes from a liquid killing agent, such as tetrachloroethane or calcium cyanide. A heaping tablespoon or more of calcium cyanide is placed in each of four to six brown paper bags, which are hung in a large garbage can or other large container. A dampened cloth, such as a washcloth, is also hung inside the can to humidify the air and activate the cyanide. This is espe- cially necessary in dry weather. The concentration of the gas inside the can is so great that insects are inactivated almost instantly on entering, and even the most delicate specimens are damaged very little. The bags containing the calcium cyanide powder should be replaced as needed. If two of the oldest bags are replaced with two fresh ones each successive night, the trap can be run as long as the collector desires. Handle cyanide outdoors, facing downwind, and with extreme caution. During the day, when the trap is not in use, store the cyanide bags in an airtight container. All forms of cyanide used as killing agents react and break down quickly when exposed to air and moisture; neverthe- less dispose of the residue carefully. To prevent rainwater from accumulating in the trap, place a screen-covered funnel inside the collection chamber to drain the water out through a hole in the bottom of the trap. Sometimes a system of separators is added to guide beetles and other heavy, hard-bodied insects into a different part of the container than the moths and other delicate specimens. The most efficient light traps use lamps rich in their output of ultraviolet light. The British-made Robinson trap employs an intense, blue-white, 125-watt mercury vapor lamp of a type used for street lighting. This, the most effective insect attractant commercially marketed, is widely used in many kinds of light traps because it has some special advantages over other kinds of attractants. For example, this type of lamp is the only one that emits the kind of light that attracts large numbers of Catocala (underwing) moths, a colorful group popular with many collectors. Many traps are equipped with 15-watt ultraviolet fluorescent tubes, which emit a highly visible bluish-white light, although blacklight tubes emitting deep purple light are similarly effective. Ultraviolet tubes of lower or higher wattage also may be used and are all highly effective. A 15-watt ultraviolet tube has been estimated to attract about 10 times as many insects as a 500 candlepower gasoline lantern or incandescent lamp. The advantage of the fluorescent tube over the mercury vapor lamp is that it is less expensive and much more portable. A 15-watt tube is easily powered by an ordinary automobile battery by using an inverter to change 6- or 12-volt direct current to 120- volt alternating current. Also, its ultraviolet output is not strong enough to cause any significant eye damage. The safety factor of the mercury vapor lamp at close range is less certain, although entomologists who have used the Robinson trap for many years seem to have suffered no ill effects. A new, lightweight, spillproof 12-volt battery, in which the acid electrolyte is a gel rather than a liquid, is far superior to the standard automotive battery for power- ing light traps, but it is fairly expensive and requires a special charger. Special lightweight, nickel-cadmium battery packs, used to power blacklights for collecting, are marketed by some dealers of entomological equipment. 1.10.10 Light Sheets Another highly effective method of using light to attract moths and other nocturnal insects is with a light sheet (fig. 15). This is simply a cloth sheet, usually a white bedsheet, hung outdoors at night with an appropriate light source or combination of sources such as ultraviolet fluorescent tubes, gasoline lanterns, or automobile headlights placed a few feet in front of it. As insects are attracted and alight on the sheet, they are easily captured in cyanide bottles or jars by the collector who stands in attendance or at least checks the sheet frequently. The 18 Collecting and Preserving Insects and Mites sheet may be pinned to a rope tied between two trees or fastened to the side of a building, with the bottom edge spread out on the ground beneath the light. Some collec- tors use supports to hold the bottom edge of the sheet several centimeters above the ground so that no specimens can crawl into the vegetation under the sheet and be overlooked. Other collectors turn up the edge to form a trough into which insects may fall as they strike the sheet. The light sheet remains unsurpassed as a method of collecting moths in flawless condition or of obtaining live females for rearing purposes. Its main disadvantage is that species that fly very late or those that are active only in the early morning hours may be missed unless one is prepared to spend most of the night at the sheet. Many other insects besides moths are attracted to the sheet, and collectors of beetles, flies and other kinds of insects would do well to collect with this method. It should be emphasized that the phases of the moon may influence the attraction of insects to artificial light. A bright moon may compete with the light source resulting in a reduced catch. The best collecting period each month extends from the fifth night after the full moon until about a week before the next full moon. References (light traps and sheets): Andreyev et al. 1970; Apperson & Yows 1976; Barr et al. 1963; Barrett et al. 1971; Bartnett & Stephenson 1968; Belton & Kempster 1963; Belton & Pucat 1967; Blakeslee et al. 1959; Breyev 1963; Burbutis & Stewart 1979; Carlson 1971; Carlson 1972; Clark & Curtis 1973; Davis & Landis 1949; DeFoliart 1972; Freeman 1972; Frost 1952, 1964; Graham et al. 1961; Gurney et al. 1964; Hardwick 1968; Hathaway 1981; Hollingsworth & Hartstack 1972; Hollingsworth et al.; Howell 1980; Kovrov & Monchadskii 1963; Lowe & Putnam 1964; McDonald 1970; Meyers 1959; Miller et al. 1970; Morgan & Uebel 1974; Mulhern 1942; Nantung Institute of Agriculture 1975; Onsager 1976; Powers 1969; Pratt 1944; Smith et al. 1974; Stanley & Dominick 1970; Stewart & Payne 1971; Stewart & Lam 1968; Tedders & Edwards 1972; USDA 1961; White 1964; Wilkinson 1969; Williams 1948; Zimmerman 1978. 1.10.11 Sticky Traps In this type of trap, a board, piece of tape, pane of glass, piece of wire net, cylinder, or other object, often painted yellow, is coated with a sticky substance and suspended from a tree branch or other convenient object. Insects landing on the sticky surface are unable to extricate themselves. The sticky material is later dissolved with a suitable solvent, usually toluene, xylene, ethylacetate, or various combinations of these, and the insects are washed first in Cellosolve and then in xylene. This type of trap should not be used to collect certain specimens, such as Lepidoptera, which are ruined by the sticky substance and cannot be removed without being destroyed. Various sticky-trap materials are available commer- cially, some with added attractants. However, use caution in selecting a sticky substance because some are difficult to dissolve. References: Buriff 1973; Chiang 1973; Dominick 1972; Edmunds et al. 1976; Evans 1975; Gillies & Snow 1967; Golding 1941, 1946; Goodenough & Snow 1973; Harris et al. 1971; Harris & McCafferty 1977; Heathcote 1957; Johnson 1950; Lambert & Franklin 1967; Mason & Sublette 1971; Maxwell 1965; Moreland 1955; Murphy 1962 (pp.226-227), 1985; Prokopy 1968; Still 1960; Taylor 1962b; Williams 1973. 1.10.12 Snap Traps Two kinds of traps designed for quantitative sam- pling may be termed “snap traps.” One of them (see Menzies & Hagley 1977) consists of a pair of wooden or plastic discs, slotted to the center so as to fit on a tree branch and connected to each other by a pair of rods. A cloth cylinder is affixed at one end to one of the discs and at the other end to a ring sliding on the rods. After the cloth cylinder has been pulled to one end and has been secured in place, the ring is held by a pair of latches. When insects have settled on the branch, its leaves, or flowers, the latches are released by pulling on a string from a distance, and the trap is snapped shut by a pair of springs on the rods, capturing any insects present. One of the canopy traps (see Turnbull & Nichols 1966) operates in a similar fashion. When a remotely controlled latch is Fig. 15. A light sheet in the field 19 Techniques and Tools pulled, a spring-loaded canopy is snapped over an area of soil, and insects within the canopy are collected by suction or a vacuum device. This trap was designed for use in grasslands. 1.10.13 Artificial Refuges Many insects, especially beetles, are successfully found under stones, planks, or rotten logs. Providing such refuges, as pieces of wood, card board, or even complex traps, is also a form of trapping. Lepidoptera larvae, for example, will congregate under burlap tied in a band around the trunks of trees. This technique has even been used to help control some pest species such as the gypsy moth. References: Campion 1972; Shubeck 1976. 1.10.14 Electrical Grid Traps In recent years, electrocuting pest insects has been used extensively in control work. The insects are attracted to a device by a pheromone or other lure placed in a chamber protected by a strongly charged electrical grid. The method deserves study for other purposes, such as surveying the arthropod fauna of an area. References: Goodenough & Snow 1973; Hartstack et al. 1968; Mitchell et al. 1972, 1973, 1974; Rogers & Smith 1977; Stanley et al. 1977. 1.11 Baits, Lures, and Other Attractants Any substance that attracts insects may be used as a bait. Natural products, chemicals derived therefrom or synthesized, and secretions of the insects themselves may all be used as attractants. Mere exposure of the substance may be considered as setting up a trap, and attractive substances are used in many constructed traps. Sugaring for moths, one of the oldest collecting methods, involves the use of a specially prepared bait in which some form of sugar is an essential component. The bait may be refined or brown sugar, molasses, or sirup. Such substances often are mixed with stale beer, fermented peaches, bananas, or some other fruit— there is no standard formula. Each lepidopterist has his or her own favorite recipe. One particularly satisfactory recipe uses fresh, ripe peaches; culls or windfalls are suitable. Remove the seeds but not the skins, mash the fruit, then place it in a 4-liter (1-gal) or larger container of plastic, glass, stainless steel, enamelware, or crockery with a snugly fitting but not tight cover. Avoid using metal containers that may rust or corrode. Fill each container only onehalf to two-thirds full to allow space for expansion. Add about a cup of sugar and place in a moderately warm place for the mixture to ferment. The bubbling fermentation reaction should start in a day or so and may continue for 2 weeks or more, depending on the temperature. During this time, check the fermentation every day or every other day and add sugar until fermentation appears to have subsided completely. As the added sugar is converted to alcohol, the growth of yeast slows and eventually ceases. After fermentation ceases, the bait should remain stable and should then be kept in tightly sealed containers to prevent contamination and evaporation. If the mixture is allowed to run low in sugar during the fermentation process, vinegar will be produced instead of alcohol. It is therefore important to smell the bait periodically and to add plenty of sugar to avoid this. The amount of sugar consumed will be surprising, usually over 0.4 kg per liter (3.3 lb per gal). The bait should have a sweet, fruity, winelike fragrance. A trace of vinegar is not objectionable but is better avoided. Canned fruit, such as applesauce, may also be used to make the bait, but inasmuch as such products are completely sterile, a small amount of yeast must be added to start fermentation. Although the bait may seem troublesome to prepare, it keeps for years and is thus available at any time, even when fruit is not in season. Immediately before use, the bait may be mixed with 30 to 50 percent molasses or brown sugar or a mixture of these. This thickens the bait so that it will not dry out so quickly, and it makes the supply last longer. The best time to set out the sugar bait is in the early evening before dark. It may be applied with a paint brush Fig. 15. A Manitoba trap. 20 Collecting and Preserving Insects and Mites in streaks on tree trunks, fenceposts, or other surfaces. Choose a definite route, such as along a trail or along the edge of a field, so that later you can follow it in the dark with a lantern or flashlight. Experienced collectors learn to approach the patches of bait stealthily with a light in one hand and a killing jar in the other to catch the moths before they are frightened off. Some collectors prefer to wear a headlamp, leaving both hands free. Although some moths will fly away and be lost, a net usually is regarded as an unnecessary encumbrance, because moths can be directed rather easily into the jar. Sugaring is an especially useful way to collect noctuid moths, and the bait applied in the evening often will attract various diurnal insects on the following days. The peach bait previously described has been used in butterfly traps with spectacular results. However, collecting with baits is notoriously unpredict- able, being extremely productive on one occasion and disappointing on another, under apparently identical conditions. 1.11.1 Baiting With Feces. Animal and human feces attract many insects. A simple but effective method of collecting such insects is to place fresh feces on a piece of paper on the ground and wait a few minutes. When a sufficient number of insects have arrived, a net with its bag held upward can be brought carefully over the bait about 1 meter above it. This will not disturb the insects, nor will they be greatly disturbed when the net is lowered gently about two- thirds of the distance to the bait. At this point, the net should be quickly lowered until its rim strikes the paper. The insects, mostly flies, will rise into the net, which may then be lifted a short distance above the bait and quickly swung side- ways, capturing the insects in the bottom of the bag. In about half an hour, many flies can be caught, virtually all that have come to the bait. Because of this, the ‘baiting with feces’ method may be used for quantitative studies (see Steyskal 1957). Feces are most attractive to insects during the first hour after deposition, but insects coming for a more extended period may be captured by placing a canopy trap over the feces or by using the feces with the cereal dish trap (see p. 12). Emergence traps placed over old feces will capture adult insects emerging from immature forms feeding there. The same methods also may be used with other baits, such as decaying fruit, small carcasses, and a wide variety of other substances. 1.11.2 The Oatmeal Trail Hubbell (1956) showed that dry oatmeal scattered along a path will attract such insects as crickets, camel crickets, cockroaches, and ants. Some of these insects feed only at night and may be hand-collected by flashlight or by light from a headlamp. A killing bottle is used, and the specimens are collected with fingers, an aspirator, or a net. 1.11.3 Pheromones and Other Attractants Substances naturally produced by insects to attract others of their own kind are known as pheromones. They are often used in traps to aid in controlling pest species. Most pheromones are highly specific, attracting only one species or a group of closely related species. “Spanish Fly” (cantharidin) has recently come into use as an extremely effective attractant for various beetles, such as pedilids, and bugs, such as bryocerines. Female specimens of certain insects, such as cicadas and silkworm moths, may be placed alive in a trap and used as a bait with their pheromones and the sounds they produce attracting males. Female saturniids (silkworm moth) may be used to attract males which may come from great distances. The phero- mones of sesiid moths are commercially available and can be attached to the collector's net or hung over a dish with ethylene glycol. Host animals likewise may be used as bait for various bloodsucking insects, with or without constructed traps. Carbon dioxide in the form of “Dry Ice,” cylinder gas, or marble chips treated with an acid such as vinegar serves as an attractant for certain insects and has been very successful in attracting horse flies to Malaise and Manitoba traps. 1.11.4 - Sounds, etc. Sounds are produced by many insects to attract others of their own kind. These sounds are very specific in pitch, tempo, and duration. Recordings of such sounds, played at the proper volume, have been effective in luring grasshoppers, crickets, and other kinds of insects. Hesperiid moths (skippers) have been shown to be attracted to small pieces of wetted paper placed on vegetation (Lamas et al., 1993). References for attractants: General—Acree et al. 1968; Atkins 1957; Beavers et al 1972; Beroza 1970, 1972; Beroza & Green 1963; Bram 1978; Carestia & Savage 1967; Clinch 1971; Coffey 1966; Debolt et al. 1975; DeJong 1967; Fahy 1972; Golding 1941; Greenslade 1964; Hocking 1963; Howell 1980; Hubbell 1956; Jacobson & Beroza 1964; Laird 1981; Lee et al. 1982; LeSage & Harrison 1979; Luff 1975; Macleod & Donnelly 1956; Mason 1963; Morris & DeFoliart 1969; Nakagawa et al. 1971; Newhouse et al. 1966; Pinniger 1975; Rennison & Robertson 1959; Roberts 1972; Sanders & Dobson 1966; Shorey & McKelvey 1977; Steyskal 1957; Strenzke 1966; Walsh 1933; Wellso & Fischer 1971; Wilton 1963; carbon dioxide—Blume et al. 1972; Carestis 21 Techniques and Tools & Savage 1967; Davidson & Swan 1933; Debolt et al. 1975; Evans 1975; Fahy 1972; Gillies & Snow 1967; Hoy 1970; Kato et al. 1966; Knox & Hays 1972; Morris & DeFoliart 1969; Newhouse et al. 1966; Reeves 1951, 1953; Rennison & Robertson 1959; Roberts 1972; Snoddy & Hays 1966; Stryker & Young 1970; Takeda et al. 1962; Whitsel & Schoeppner 1965; Wilson et al. 1972; phero- mones—Beaudry 1954; Bellamy & Reeves 1952; Beroza et al. 1974; Birch 1974; Campion 1972; Campion et al. 1974; Goonewardene et al. 1973; Hathaway 1981; Holbrook & Beroza 1960; Howell 1980; Howland et al 1969; Jacobson 1972; Jacobson & Beroza 1964; Mitchell et al. 1972; Neal 1979; Peacock & Cuthbert 1975; Shorey 1973; Shorey & McKelvey 1977; Sparks et al. 1980; Steck & Bailey 1978; Weatherston 1976; sound—Belton 1962; Cade 1975. 1.12 - Collecting Aquatic and Soil Insects and Ectoparasites Insects and mites emerging from water may be collected by the same means as terrestrial insects, but specialized equipment is required. Aquatic insects are of great importance to water quality and are being intensely investigated in biodiversity studies. The following references pertain to aquatic collecting. References: Apperson & Yows 1976; Carlson 1971; Coon & Pepper 1968; Coulson et al. 1970; Eastop 1955; Edmondson & Winberg 1971; Edwards et al. 1981; English 1987; Essig 1958; Gerking 1957; Hodgson 1940; Jonasson 1954; Kimerle & Anderson 1967; LaGasa & Smith 1978; Langford & Daffern 1975; Lawson & Merritt 1979; LeSage & Harrison 1979; Macan 1964; McCauley 1976; Mason & Sublette 1971; Masteller 1977; Merritt et al. 1978 (general); Morgan et al. 1963; Mundie 1956, 1964, 1966, 1971; Murray & Charles 1975; Pennak 1978 (general); Piecrynski 1961; Sladeckova 1962; Tarshis 1968a, 1968b; Waters 196; Welch 1848 (general); Wood & Davies 1966; Weber 1987; Wood et al. 1979. As with aquatic specimens, insects and mites that live on or under the soil surface require special techniques and equipment for their collection and study. Many soilinhabiting species are of great economic importance because they devour the roots of crops. Many spend their immature stages in soil but emerge and leave the soil as adults. A considerable amount of literature on soil insects has been published, the most useful of which is cited here. See also the references cited under Separators and Extrac- tors (p. 9) and Pitfall and Dish Traps (p. 12) References: Barnes 1941; Briggs 1971; Brindle 1963; Dethier 1955; Fessenden 1949; Kevan 1955, 1962; Kuhnelt et al. 1976; Lane & Anderson 1976; MacFayden 1962; Murphy 1962; Newell 1955; Paquin and Coderre 1996; Salt & Hollick 1944; Teskey 1962. Some ectoparasites, particularly those that fly, may be collected in some of the traps discussed,using their hosts as bait; others may be collected by means of the special devices described in the following references. References: British Museum 1974 (p. 152), Comstock 1940; Watson & Amerson 1967; Williamson 1954. 1.13 - Rearing Collectors should take every opportunity to rear insects and mites. Not only are reared specimens generally in the best possible condition, but rearing provides life stages that otherwise might be collected only rarely or with great difficulty. By preserving one or more specimens from each of the stages as they are reared, if sufficient material is available, the collector can obtain series of immature stages along with associated adults. Such series are extremely desirable, especially for species in which the adult is known but the immature stages are unknown or difficult to identify. The converse often is true also—some species of insects, such as stem-mining flies, are fairly abundant in the larval stage but have never been reared to the adult stage; consequently, one does not know whether they are stages of a species that has been described and named from an adult but whose life history is unknown. Since adults of these flies are seldom found, the easiest way to obtain the stage necessary for specific determina- tion is to rear the larvae or pupae. If only a few specimens are reared, the shed skins and pupal cases or puparia should be preserved, as they are of value if properly associated with the reared adult. Do not preserve a pupa or puparium with an adult unless you are positive that the association is correct. It is best to put pupae in separate containers so that adults or parasites that emerge are associated with certainty. If at all feasible, the parasite’s host should be preserved for identification. Keep careful notes throughout the rearing so that all data relative to the biology of the species are properly correlated. 1.13.1- Containers for Rearing To rear specimens successfully, simulate as closely as possible in the rearing cages the natural conditions under which the immatures werq found. Almost any container will serve as a temporary cage for living insects or mites. One simple temporary cage that is very handy on field trips is a paper bag. Plant material or a soil sample containing insects or mites is placed in the paper bag, which is then sealed. A paper bag also can be placed over the top of a plant on which insects or mites are found. The 22 Collecting and Preserving Insects and Mites bottom edge of the bag is tied tightly around the exposed stems, which are cut and placed in a jar of water. One disadvantage of using a paper bag is that it is not transpar- ent, so it must be removed to observe the specimens or to determine when the foliage needs to be changed. Clear plastic bags are better suited to such viewing. Plastic bags with a paper towel placed in the bottom are extrememly efficient rearing containers for leaf mining and other small moths. They require frequent inversion to minimize condensation. Another simple temporary cage is a glass jar with its lid replaced by a piece of organdy cloth or gauze held in place by a rubberband. A few such jars in a collecting kit are useful for holding live insects. For aquatic species, using a watertight lid on the jars is advisable. If aquatic insects are to be transported over a considerable distance, fewer will die if the jar is packed with wet moss or leaves than if the specimens are allowed to slosh around in water alone. After arrival at your destination, release the insects into a good rearing container (fig. 12). Aquatic insects can be reared in their natural habitat by confining them in a wire screen or gauze cage, part of which is submerged in water. Be sure to anchor the cage securely. The screen used in aquatic cages should be coarse enough to allow food to flow through, yet fine enough to retain the insects being reared. Certain aquatic insects may be reared readily indoors in an aquarium or even in a glass jar. The main goal is to try to duplicate their natural habitat. If the specimen was collected from a rapidly flowing stream, it is unlikely to survive indoors unless the water is aerated. Other insects do well in stagnant water. Aquatic vegetation usually should be provided in the aquarium even for predaceous specimens, such as dragonfly nymphs, which often are found clinging to underwater stems. Keep sufficient space, which will vary according to the insect being reared, between the surface of the water and the screen or gauze cover over the aquarium to allow the adult insect to emerge. A dragonfly, for example, needs considerable space, plus a stick, rock, or other object on which to perch after emerging so that the wings will develop fully. Most adult insects, both terrestrial and aquatic, are teneral when they first emerge and should not be killed until the exoskeleton and wings harden and the colors develop fully. This may be a matter of minutes, hours, or even days. It is advisable to keep even small flies alive for 1 full day after they emerge. Specimens killed while still teneral will shrivel when mounted. Some insects, if kept in cages too long after emerging, especially butterflies and moths, will beat their wings against the cage and lose many scales or tear their wings. Providing adequate space in which emerging insects may expand their wings fully and move about slightly is therefore critical in the design of rearing cages. Beetles and other boring insects often are abundant in bark and wood. If pieces of such material are placed in glass or metal containers, excellent specimens of the adults may be obtained, although sometimes not for a considerable time. Cages made of wood or cardboard are not suitable for such insects because those found in wood or bark usually are well equipped, both in immature and adult stages, to chew their way through a cage made of such material and thus escape. A flowerpot cage is one of the best containers for rearing plant-feeding species over an extended period. The host plant, if its size and habitat permit, is placed in a flowerpot, and a cylinder of glass, plastic, or wire screen is placed around the plant (fig. 12, lower left). Another type of flowerpot cage is made by inserting a cane or stick, taller than the plant, into the soil in the pot. One end of a net or muslin tube is fitted over the edge of the pot and is held in place by a string. The other end of the tube is tied around the top of the stick. An advantage of the flowerpot cage is that the plant is living, and fresh plant material need not be added daily. Plant-feeding mites will not wander far as long as suitable host material is available for them. Because mites are wingless even as adults, they can be confined in an open rearing container by making a barrier around the top edge or upper inner sides of the container with Vaseline or talcum powder. Emergence cages are essentially rearing cages that are used when it is impractical or impossible to bring specimens indoors. Emergence cages may also be consid- ered as traps and are discussed under that heading (see p. 13). With plant- feeding insects, a sleeve consisting of a muslin tube with open ends is slipped over a branch or plant and tied at one end. The insects are then placed in the tube, and the loose end of the tube is tied. This cloth tube can be modified to allow observation of the insects by replacing the midsection with a “window” of clear plastic or wire screen. If the insects in the tube require duff or debris in which to pupate, the tube should be placed perpendicular to the ground and duff or debris placed in the lower end. 1.13.2 - Rearing Conditions and Problems 1.13.2.1 - Moisture The moisture requirements of insects and mites are varied. Examination of the habitat from which specimens were collected should provide clues about their moisture requirements in captivity. Many insects in the pupal stage are resistant to drought. Species that normally infest stored 23 Techniques and Tools foods also require very little moisture; in fact, many produce their own water. Most species found outdoors require higher levels of humidity than are generally found indoors. Additional moisture can be added to indoor rearing cages in several ways. To increase the humidity in a cage, keep a moist pad of cotton on top of the screen cover of the cage, or place a moist sponge or a small glass vial filled with water in the cage. The mouth of the vial is plugged with cotton and the vial laid on its side so the cotton remains moist. Pupae may be held for long periods in moist sawdust, vermiculite, sphagnum, or peat moss. In a flowerpot cage, the water used to keep the plant alive should provide sufficient moisture for the plant feeding insects and mites. Spraying the leaves daily also may supplement moisture requirements in rearing cages. Too much moisture may result in water condensation on the sides of the cage, which may trap the specimens and damage or kill them. Excess moisture also enhances the growth of mold and fungus, which is detrimental to the development of most insects and mites. A 2-3 percent solution of table salt sprayed regularly in the cage will help prevent mold and fungus growth. 1.13.2.2 - Temperature Of all the environmental factors affect ing the development and behavior of insects and mites, tempera- ture may be the most critical. Since arthropods are cold blooded, their body temperatures are usually close to the temperature of the surrounding environment, and their metabolism and development are directly affected by increases and decreases in temperature. Each stage of an insect or mite species has a low and a high point at which development ceases. These are called threshold tempera- ture levels. Most species that are collected and brought indoors for rearing can be held at normal room tempera- ture; the optimum temperature for rearing will vary from species to species and with different stages of the same species. As with all rearing techniques, every attempt should be made to duplicate natural conditions. Specimens that normally would overwinter outdoors should be kept during the winter in rearing cages placed in an unheated room, porch, or garage. Never place an enclosed rearing cage in direct sunlight; the heat becomes too intense and may kill the specimens. 1.13.2.3 - Dormancy and Diapause Insects and mites are unable to control the tempera- ture of their environment; instead, they make physiological adjustments that allow them to survive temperature extremes. In regions with freezing winters, insects and mites have at least one stage that is resistant to low temperatures. The resistant form may be any stage—egg, larva, nymph, pupa, or adult. When winter arrives, only the resistant form survives. Dormancy is the physiological state of an insect or mite during a period of arrested development, whereas diapause is the prolonged period of arrested development brought about by such adverse conditions as heat, drought, or cold. This condition can be used to advantage in rearing. For example, if leaving rearing cages unattended for several days or longer is unavoidable, many (but unfortunately not all) specimens can be refrigerated temporarily to slow their activity and perhaps force diapause. This measure should be used with caution since the degree and duration of cold tolerated by different species will vary. The reverse situation, that of causing diapause to end, is equally useful. Overwintering pupae that normally would not develop into adults until spring can be forced to terminate diapause early by chilling them for several weeks or months, then bringing them to room temperature so normal activity will resume. Often mantid egg cases are brought indoors accidentally with Christmas greenery. The eggs, already chilled for several months, hatch when kept at room temperature, often to the complete surprise and consternation of the unsuspecting homeowner. 1.13.2.4 - Light Most species of insects and mites can be reared under ordinary lighting conditions; however, artificial manipulation of the light period will control diapause in many species. If the light requirements of the species being reared are known, it may be possible to adjust the period of light so that the specimens will continue to develop and will remain active instead of entering dia- pause, for exam