The Insects - Outline of Entomology 3th Edition - Chapter 17 docx

Chapter 17 METHODS IN ENTOMOLOGY: COLLECTING, PRESERVATION, CURATION, AND IDENTIfiCATION Alfred Russel Wallace collecting butterflies. (After various sources, especially van Oosterzee 1997; Gardiner 1998.) TIC17 5/20/04 4:39 PM Page 427 428 Methods in entomology For many entomologists, questions of how and what to collect and preserve are determined by the research project (see also section 13.4). Choice of methods may depend upon the target taxa, life-history stage, geo- graphical scope, kind of host plant or animal, disease vector status, and most importantly, sampling design and cost-effectiveness. One factor common to all such studies is the need to communicate the information unambiguously to others, not least concerning the identity of the study organism(s). Undoubtedly, this will involve identification of specimens to provide names (section 1.4), which are necessary not only to tell others about the work, but also to provide access to previously published studies on the same, or related, insects. Identification requires material to be appropri- ately preserved so as to allow recognition of morpho- logical features which vary among taxa and life-history stages. After identifications have been made, the speci- mens remain important, and even have added value, and it is important to preserve some material (vouch- ers) for future reference. As information grows, it may be necessary to revisit the specimens to confirm iden- tity, or to compare with later-collected material. In this chapter we review a range of collecting methods, mounting and preservation techniques, and specimen curation, and discuss methods and principles of identification. 17.1 COLLECTION Those who study many aspects of vertebrate and plant biology can observe and manipulate their study organ- isms in the field, identify them and, for larger animals, also capture, mark, and release them with few or no harmful effects. Amongst the insects, these techniques are available perhaps only for butterflies and dragon- flies, and the larger beetles and bugs. Most insects can be identified reliably only after collection and preserva- tion. Naturally, this raises ethical considerations, and it is important to: • collect responsibly; • obtain the appropriate permit(s); • ensure that voucher specimens are deposited in a well-maintained museum collection. Responsible collecting means collecting only what is needed, avoidance or minimization of habitat destruc- tion, and making the specimens as useful as possible to all researchers by providing labels with detailed collec- tion data. In many countries or in designated reserve areas, permission is needed to collect insects. It is the collector’s responsibility to apply for permits and fulfill the demands of any permit-issuing agency. Further- more, if specimens are worth collecting in the first place, they should be preserved as a record of what has been studied. Collectors should ensure that all speci- mens (in the case of taxonomic work) or at least repres- entative voucher specimens (in the case of ecological, genetic, or behavioral research) are deposited in a recognized museum. Voucher specimens from surveys or experimental studies may be vital to later research. Depending upon the project, collection methods may be active or passive. Active collecting involves search- ing the environment for insects, and may be preceded by periods of observation before obtaining specimens for identification purposes. Active collecting tends to be quite specific, allowing targeting of the insects to be collected. Passive collecting involves erection or installation of traps, lures, or extraction devices, and entrapment depends upon the activity of the insects themselves. This is a much more general type of collect- ing, being relatively unselective in what is captured. 17.1.1 Active collecting Active collecting may involve physically picking indi- viduals from the habitat, using a wet finger, fine-hair brush, forceps, or an aspirator (also known in Britain as a pooter). Such techniques are useful for relatively slow-moving insects, such as immature stages and sedentary adults that may be incapable of flying or reluctant to fly. Insects revealed by searching particu- lar habitats, as in turning over stones, removing tree bark, or observed at rest by night, are all amenable to direct picking in this manner. Night-flying insects can be selectively picked from a light sheet – a piece of white cloth with an ultraviolet light suspended above it (but be careful to protect eyes and skin from exposure to ultraviolet light). Netting has long been a popular technique for capturing active insects. The vignette for this chapter depicts the naturalist and biogeographer Alfred Russel Wallace attempting to net the rare butterfly, Graphium androcles, in Ternate in 1858. Most insect nets have a handle about 50 cm long and a bag held open by a hoop of 35 cm diameter. For fast-flying, mobile insects such as butterflies and flies, a net with a longer handle and a wider mouth is appropriate, whereas a net with a narrower mouth and a shorter handle is sufficient for TIC17 5/20/04 4:39 PM Page 428 small and/or less agile insects. The net bag should always be deeper than the diameter so that the insects caught may be trapped in the bag when the net is twisted over. Nets can be used to capture insects whilst on the wing, or by using sweeping movements over the substrate to capture insects as they take wing on being disturbed, as for example from flower heads or other vegetation. Techniques of beating (sweeping) the vegetation require a stouter net than those used to intercept flight. Some insects when disturbed drop to the ground: this is especially true of beetles. The tech- nique of beating vegetation whilst a net or tray is held beneath allows the capture of insects with this defen- sive behavior. Indeed, it is recommended that even when seeking to pick individuals from exposed posi- tions, that a net or tray be placed beneath for the inevitable specimen that will evade capture by drop- ping to the ground (where it may be impossible to locate). Nets should be emptied frequently to prevent damage to the more fragile contents by more massive objects. Emptying depends upon the methods to be used for preservation. Selected individuals can be removed by picking or aspiration, or the complete contents can be emptied into a container, or onto a white tray from which targeted taxa can be removed (but beware of fast fliers departing). The above netting techniques can be used in aquatic habitats, though specialist nets tend to be of different materials from those used for terrestrial insects, and of smaller size (resistance to dragging a net through water is much greater than through air). Choice of mesh size is an important consideration – the finer mesh net required to capture a small aquatic larva compared with an adult beetle provides more resistance to being dragged through the water. Aquatic nets are usually shallower and triangular in shape, rather than the cir- cular shape used for trapping active aerial insects. This allows for more effective use in aquatic environments. 17.1.2 Passive collecting Many insects live in microhabitats from which they are difficult to extract – notably in leaf litter and similar soil debris or in deep tussocks of vegetation. Physical inspection of the habitat may be difficult and in such cases the behavior of the insects can be used to separate them from the vegetation, detritus, or soil. Particularly useful are negative phototaxic and thermotaxic and positive hygrotaxic responses in which the target insects move away from a source of strong heat and/or light along a gradient of increasing moisture, at the end of which they are concentrated and trapped. The Tullgren funnel (sometimes called a Berlese funnel) comprises a large (e.g. 60 cm diameter) metal funnel tapering to a replaceable collecting jar. Inside the funnel a metal mesh supports the sample of leaf litter or vegetation. A well-fitting lid containing illuminating lights is placed just above the sample and sets up a heat and humidity gradient that drives the live animals downwards in the funnel until they drop into the collecting jar, which contains ethanol or other preservative. The Winkler bag operates on similar principles, with drying of organic matter (litter, soil, leaves) forcing mobile animals downwards into a collecting chamber. The device consists of a wire frame enclosed with cloth that is tied at the top to ensure that speci- mens do not escape and to prevent invasion by scav- engers, such as ants. Pre-sieved organic matter is placed into one or more mesh sleeves, which are hung from the metal frame within the bag. The bottom of the bag tapers into a screw-on plastic collecting jar con- taining either preserving fluid or moist tissue paper for live material. Winckler bags are hung from a branch or from rope tied between two objects, and operate via the drying effects of the sun and wind. However, even mild windy conditions cause much detritus to fall into the residue, thus defeating the major purpose of the trap. They are extremely light, require no electric power and are very useful for collecting in remote areas, although when housed inside buildings or in areas subject to rain or high humidity, they can take many days to dry com- pletely and thus extraction of the fauna may be slow. Separating bags rely on the positive phototaxic (light) response of many flying insects. The bags are made from thick calico with the upper end fastened to a supporting internal ring on top of which is a clear Perspex lid; they are either suspended on strings or supported on a tripod. Collections made by sweeping or specialized collections of habitat are introduced by quickly tipping the net contents into the separator and closing the lid. Those mobile (flying) insects that are attracted to light will fly to the upper, clear surface, from which they can be collected with a long-tubed aspirator inserted through a slit in the side of the bag. Insect flight activity is seldom random, and it is pos- sible for the observer to recognize more frequently used routes and to place barrier traps to intercept the flight path. Margins of habitats (ecotones), stream lines, and Collection 429 TIC17 5/20/04 4:39 PM Page 429 430 Methods in entomology gaps in vegetation are evidently more utilized routes. Traps that rely on the interception of flight activity and the subsequent predictable response of certain insects include Malaise traps and window traps. The Malaise trap is a kind of modified tent in which insects are inter- cepted by a transverse barrier of net material. Those that seek to fly or climb over the vertical face of the trap are directed by this innate response into an uppermost corner and from there into a collection jar, usually containing liquid preservative. A modified Malaise trap, with a fluid-filled gutter added below, can be used to trap and preserve all those insects whose natural reaction is to drop when contact is made with a barrier. Based on similar principles, the window trap consists of a window-like vertical surface of glass, Perspex, or black fabric mesh, with a gutter of preserving fluid lying beneath. Only insects that drop on contact with the window are collected when they fall into the preserving fluid. Both traps are conventionally placed with the base to the ground, but either trap can be raised above the ground, for example into a forest canopy, and still function appropriately. On the ground, interception of crawling insects can be achieved by sinking containers into the ground to rim-level such that active insects fall in and cannot climb out. These pitfall traps vary in size, and may feature a roof to restrict dilution with rain and preclude access by inquisitive vertebrates (Fig. 17.1). Trapping can be enhanced by construction of a fence-line to guide insects to the pitfall, and by baiting the trap. Specimens can be collected dry if the container con- tains insecticide and crumpled paper, but more usually they are collected into a low-volatile liquid, such as pro- pylene glycol or ethylene glycol, and water, of varying composition depending on the frequency of visitation to empty the contents. Adding a few drops of detergent to the pitfall trap fluid reduces the surface tension and prevents the insects from floating on the surface of the liquid. Pitfall traps are used routinely to estimate species richness and relative abundances of ground active insects. However, it is too rarely understood that strong biases in trapping success may arise between compared sites of differing habitat structure (density of vegetation). This is because the probability of capture of an individual insect (trappability) is affected by the complexity of the vegetation and/or substrate that sur- rounds each trap. Habitat structure should be meas- ured and controlled for in such comparative studies. Trappability is affected also by the activity levels of insects (due to their physiological state, weather, etc.), their behavior (e.g. some species avoid traps or escape from them), and by trap size (e.g. small traps may exclude larger species). Thus, the capture rate (C) for pitfall traps varies with the population density (N) and trappability (T ) of the insect according to the equation C = TN. Usually, researchers are interested in estimat- ing the population density of captured insects or in determining the presence or absence of species, but such studies will be biased if trappability changes between study sites or over the time interval of the study. Similarly, comparisons of the abundances of different species will be biased if one species is more trappable than another. Many insects are attracted by baits or lures, placed in or around traps; these can be designed as “generic” to lure many insects, or “specific”, designed for a single target. Pitfall traps, which trap a broad spectrum of mobile ground insects, can have their effectiveness increased by baiting with meat (for carrion attraction), dung (for coprophagous insects such as dung beetles), fresh or rotting fruit (for certain Lepidoptera, Coleop- tera, and Diptera), or pheromones (for specific target insects such as fruit flies). A sweet, fermenting mixture of alcohol plus brown sugar or molasses can be daubed on surfaces to lure night-flying insects, a method termed “sugaring”. Carbon dioxide and volatiles such as butanol can be used to lure vertebrate-host-seeking insects such as mosquitoes and horseflies. Colors differentially attract insects: yellow is a strong lure for many hymenopterans and dipterans. This behavior is exploited in yellow pan traps which are simple yellow dishes filled with water and a surface- Fig. 17.1 A diagrammatic pitfall trap cut away to show the inground cup filled with preserving fluid. (After an unpublished drawing by A. Hastings.) TIC17 5/20/04 4:39 PM Page 430 tension reducing detergent and placed on the ground to lure flying insects to death by drowning. Outdoor swimming pools act as giant pan traps. Light trapping (see section 17.1.1 for light sheets) exploits the attraction to light of many nocturnal flying insects, particularly to the ultraviolet light emitted by fluorescent and mercury vapor lamps. After attraction to the light, insects may be picked or aspirated indi- vidually from a white sheet hung behind the light, or they may be funneled into a container such as a tank filled with egg carton packaging. There is rarely a need to kill all insects arriving at a light trap, and live insects may be sorted and inspected for retention or release. In flowing water, strategic placement of a stationary net to intercept the flow will trap many organisms, including live immature stages of insects that may otherwise be difficult to obtain. Generally, a fine mesh net is used, secured to a stable structure such as bank, tree, or bridge, to intercept the flow in such a way that drifting insects (either deliberately or by dislodgement) enter the net. Other passive trapping techniques in water include emergence traps, which are generally large inverted cones, into which adult insects fly on emergence. Such traps also can be used in terrestrial situations, such as over detritus or dung, etc. 17.2 PRESERVATION AND CURATION Most adult insects are pinned or mounted and stored dry, although the adults of some orders and all soft- bodied immature insects (eggs, larvae, nymphs, pupae or puparia) are preserved in vials of 70–80% ethanol (ethyl alcohol) or mounted onto microscope slides. Pupal cases, cocoons, waxy coverings, and exuviae may be kept dry and either pinned, mounted on cards or points, or, if delicate, stored in gelatin capsules or in preserving fluid. 17.2.1 Dry preservation Killing and handling prior to dry mounting Insects that are intended to be pinned and stored dry are best killed either in a killing bottle or tube con- taining a volatile poison, or in a freezer. Freezing avoids the use of chemical killing agents but it is important to place the insects into a small, airtight container to pre- vent drying out and to freeze them for at least 12–24 h. Frozen insects must be handled carefully and properly thawed before being pinned, otherwise the brittle appendages may break off. The safest and most readily available liquid killing agent is ethyl acetate, which although flammable, is not especially dangerous unless directly inhaled. It should not be used in an enclosed room. More poisonous substances, such as cyanide and chloroform, should be avoided by all except the most experienced entomologists. Ethyl acetate killing con- tainers are made by pouring a thick mixture of plaster of Paris and water into the bottom of a tube or wide- mouthed bottle or jar to a depth of 15–20 mm; the plaster must be completely dried before use. To “charge” a killing bottle, a small amount of ethyl acetate is poured onto and absorbed by the plaster, which can then be covered with tissue or cellulose wadding. With frequent use, particularly in hot weather, the container will need to be recharged regularly by adding more ethyl acetate. Crumpled tissue placed in the container will prevent insects from contacting and damaging each other. Killing bottles should be kept clean and dry, and insects should be removed as soon as they die to avoid color loss. Moths and butterflies should be killed separately to avoid them contaminating other insects with their scales. For details of the use of other killing agents, refer to either Martin (1977) or Upton (1991) under Further reading. Dead insects exhibit rigor mortis (stiffening of the muscles), which makes their appendages difficult to handle, and it is usually better to keep them in the killing bottle or in a hydrated atmosphere for 8–24 h (depending on size and species) until they have relaxed (see below), rather than pin them immediately after death. It should be noted that some large insects, espe- cially weevils, may take many hours to die in ethyl acetate vapors and a few insects do not freeze easily and thus may not be killed quickly in a normal household freezer. It is important to eviscerate (remove the gut and other internal organs of ) large insects or gravid females (especially cockroaches, grasshoppers, katydids, man- tids, stick-insects, and very large moths), otherwise the abdomens may rot and the surface of the specimens go greasy. Evisceration, also called gutting, is best carried out by making a slit along the side of the abdomen (in the membrane between the terga and sterna) using fine, sharp scissors and removing the body contents with a pair of fine forceps. A mixture of 3 parts talcum powder and 1 part boracic acid can be dusted into the body cavity, which in larger insects may be stuffed carefully with cotton wool. Preservation and curation 431 TIC17 5/20/04 4:39 PM Page 431 432 Methods in entomology The best preparations are made by mounting insects while they are fresh, and insects that have dried out must be relaxed before they can be mounted. Relaxing involves placing the dry specimens in a water-saturated atmosphere, preferably with a mold deterrent, for one to several days depending on the size of the insects. A suitable relaxing box can be made by placing a wet sponge or damp sand in the bottom of a plastic con- tainer or a wide jar and closing the lid firmly. Most smaller insects will be relaxed within 24 h, but larger specimens will take longer, during which time they should be checked regularly to ensure they do not become too wet. Pinning, staging, pointing, carding, spreading, and setting Specimens should be mounted only when they are fully relaxed, i.e. when their legs and wings are freely movable, rather than stiff or dry and brittle. All dry- mounting methods use entomological macropins – these are stainless steel pins, mostly 32–40 mm long, and come in a range of thicknesses and with either a solid or a nylon head. Never use dressmakers’ pins for mounting insects; they are too short and too thick. There are three widely used methods for mounting insects and the choice of the appropriate method depends on the kind of insect and its size, as well as the purpose of mounting. For scientific and professional collections, insects are either pinned directly with a macropin, micropinned, or pointed, as follows. Direct pinning This involves inserting a macropin, of appropriate thickness for the insect’s size, directly through the insect’s body; the correct position for the pin varies among insect orders (Fig. 17.2; section 17.2.4) and it is important to place the pin in the suggested place to avoid damaging structures that may be useful in identification. Specimens should be positioned about three-quarters of the way up the pin with at least 7 mm protruding above the insect to allow the mount to be gripped below the pin head using entomological forceps (which have a broad, truncate end) (Fig. 17.3). Speci- mens then are held in the desired positions on a piece of polyethylene foam or a cork board until they dry, which may take up to three weeks for large specimens. A desiccator or other artificial drying methods are re- commended in humid climates, but oven temperature should not rise above 35°C. Fig. 17.2 Pin positions for representative insects: (a) larger beetles (Coleoptera); (b) grasshoppers, katydids, and crickets (Orthoptera); (c) larger flies (Diptera); (d) moths and butterflies (Lepidoptera); (e) wasps and sawflies (Hymenoptera); (f ) lacewings (Neuroptera); (g) dragonflies and damselflies (Odonata), lateral view; (h) bugs, cicadas, and leaf- and planthoppers (Hemiptera: Heteroptera, Cicadomorpha, and Fulgoromorpha). TIC17 5/20/04 4:39 PM Page 432 Micropinning (staging or double mounting) This is used for many small insects and involves pinning the insect with a micropin to a stage that is mounted on a macropin (Fig. 17.4a,b); micropins are very fine, headless, stainless steel pins, from 10 to 15 mm long, Preservation and curation 433 Fig. 17.3 Correct and incorrect pinning: (a) insect in lateral view, correctly positioned; (b) too low on pin; (c) tilted on long axis, instead of horizontal; (d) insect in front view, correctly positioned; (e) too high on pin; (f ) body tilted laterally and pin position incorrect. Handling insect specimens with entomological forceps: (g) placing specimen mount into foam or cork; (h) removing mount from foam or cork. ((g,h) After Upton 1991.) Fig. 17.4 Micropinning with stage and cube mounts: (a) a small bug (Hemiptera) on a stage mount, with position of pin in thorax as shown in Fig. 17.2h; (b) moth (Lepidoptera) on a stage mount, with position of pin in thorax as shown in Fig. 17.2d; (c) mosquito (Diptera: Culicidae) on a cube mount, with thorax impaled laterally; (d) black fly (Diptera: Simuliidae) on a cube mount, with thorax impaled laterally. (After Upton 1991.) TIC17 5/20/04 4:39 PM Page 433 434 Methods in entomology and stages are small square or rectangular strips of white polyporus pith or synthetic equivalent. The micropins are inserted through the insect’s body in the same positions as used in macropinning. Small wasps and moths are mounted with their bodies parallel to the stage with the head facing away from the macropin, whereas small beetles, bugs, and flies are pinned with their bodies at right angles to the stage and to the left of the macropin. Some very small and delicate insects that are difficult to pin, such as mosquitoes and other small flies, are pinned to cube mounts; a cube of pith is mounted on a macropin and a micropin is inserted horizontally through the pith so that most of its length protrudes, and the insect then is impaled ventrally or laterally (Fig. 17.4c,d). Pointing This is used for small insects that would be damaged by pinning (Fig. 17.5a) (but not for small moths because the glue does not adhere well to scales, nor flies because important structures are obscured), for very sclerot- ized, small to medium-sized insects (especially weevils and ants) (Fig. 17.5b,c) whose cuticle is too hard to pierce with a micropin, or for mounting small speci- mens that are already dried. Points are made from small triangular pieces of white cardboard which either can be cut out with scissors or punched out using a special point punch. Each point is mounted on a stout macropin that is inserted centrally near the base of the triangle and the insect is then glued to the tip of the point using a minute quantity of water-soluble glue, for example based on gum arabic. The head of the insect should be to the right when the apex of the point is directed away from the person mounting. For most very small insects, the tip of the point should contact the insect on the vertical side of the thorax below the wings. Ants are glued to the upper apex of the point, and two or three points, each with an ant from the same nest, can be placed on one macropin. For small insects with a sloping lateral thorax, such as beetles and bugs, the tip of the point can be bent downwards slightly before applying the glue to the upper apex of the point. Carding For hobby collections or display purposes, insects (especially beetles) are sometimes carded, which involves gluing each specimen, usually by its venter, to a rectangular piece of card through which a macropin passes (Fig. 17.5d). Carding is not recommended for adult insects because structures on the underside are Fig. 17.5 Point mounts: (a) a small wasp; (b) a weevil; (c) an ant. Carding: (d) a beetle glued to a card mount. (After Upton 1991.) TIC17 5/20/04 4:39 PM Page 434 obscured by being glued to the card; however, carding may be suitable for mounting exuviae, pupal cases, puparia, or scale covers. Spreading and setting It is important to display the wings, legs, and antennae of many insects during mounting because features used for identification are often on the appendages. Specimens with open wings and neatly arranged legs and antennae also are more attractive in a collection. Spreading involves holding the appendages away from the body while the specimens are drying. Legs and antennae can be held in semi-natural positions with pins (Fig. 17.6a) and the wings can be opened and held out horizontally on a setting board using pieces of tracing paper, cellophane, greaseproof paper, etc. (Fig. 17.6b). Setting boards can be constructed from pieces of polyethylene foam or soft cork glued to sheets of plywood or masonite; several boards with a range of groove and board widths are needed to hold insects of different body sizes and wingspans. Insects must be left to dry thoroughly before removing the pins and/or setting paper, but it is essential to keep the collection data associated correctly with each specimen during drying. A permanent data label must be placed on each macropin below the mounted insect (or its point or stage) after the specimen is removed from the drying or setting board. Sometimes two labels are used – an upper one for the collection data and a second, lower label for the taxonomic identification. See section 17.2.5 for information on the data that should be recorded. 17.2.2 Fixing and wet preservation Most eggs, nymphs, larvae, pupae, puparia, and soft- bodied adults are preserved in liquid because drying usually causes them to shrivel and rot. The most com- monly used preservative for the long-term storage of insects is ethanol (ethyl alcohol) mixed in various con- centrations (but usually 75–80%) with water. How- ever, aphids and scale insects are often preserved in lactic-alcohol, which is a mixture of 2 parts 95% Preservation and curation 435 Fig. 17.6 Spreading of appendages prior to drying of specimens: (a) a beetle pinned to a foam sheet showing the spread antennae and legs held with pins; (b) setting board with mantid and butterfly showing spread wings held in place by pinned setting paper. ((b) After Upton 1991.) TIC17 5/20/04 4:39 PM Page 435 436 Methods in entomology ethanol and 1 part 75% lactic acid, because this liquid prevents them from becoming brittle and facilitates subsequent maceration of body tissue prior to slide mounting. Most immature insects will shrink, and pig- mented ones will discolor if placed directly into ethanol. Immature and soft-bodied insects, as well as specimens intended for study of internal structures, must first be dropped alive into a fixative solution prior to liquid preservation. All fixatives contain ethanol and glacial acetic acid, in various concentrations, combined with other liquids. Fixatives containing formalin (40% formaldehyde in water) should never be used for speci- mens intended for slide mounting (as internal tissues harden and will not macerate), but are ideal for speci- mens intended for histological study. Recipes for some commonly employed fixatives are: KAA – 2 parts glacial acetic acid, 10 parts 95% ethanol, and 1 part kerosene (dye free). Carnoy’s fluid – 1 part glacial acetic acid, 6 parts 95% ethanol, and 3 parts chloroform. FAA – 1 part glacial acetic acid, 25 parts 95% ethanol, 20 parts water, and 5 parts formalin. Pampel’s fluid – 2–4 parts glacial acetic acid, 15 parts 95% ethanol, 30 parts water, and 6 parts formalin. AGA – 1 part glacial acetic acid, 6 parts 95% ethanol, 4 parts water, and 1 part glycerol. Each specimen or collection should be stored in a separ- ate glass vial or bottle that is sealed to prevent evapora- tion. The data label (section 17.2.5) should be inside the vial to prevent its separation from the specimen. Vials can be stored in racks or, to provide greater pro- tection against evaporation, they can be placed inside a larger jar containing ethanol. 17.2.3 Microscope slide mounting The features that need to be seen for the identification of many of the smaller insects (and their immature stages) often can be viewed satisfactorily only under the higher magnification of a compound microscope. Specimens must therefore be mounted either whole on glass microscope slides or dissected before mounting. Fur- thermore, the discrimination of minute structures may require the staining of the cuticle to differentiate the various parts or the use of special microscope optics such as phase- or interference-contrast microscopy. There is a wide choice of stains and mounting media, and the preparation methods largely depend on which type of mountant is employed. Mountants are either aqueous gum-chloral-based (e.g. Hoyer’s mountant, Berlese fluid) or resin-based (e.g. Canada balsam, Euparal). The former are more convenient for prepar- ing temporary mounts for some identification purposes but deteriorate (often irretrievably) over time, whereas the latter are more time-consuming to prepare but are permanent and thus are recommended for taxonomic specimens intended for long-term storage. Prior to slide mounting, the specimens generally are “cleared” by soaking in either alkaline solutions (e.g. 10% potassium hydroxide (KOH) or 10% sodium hydroxide (NaOH)) or acidic solutions (e.g. lactic acid or lactophenol) to macerate and remove the body con- tents. Hydroxide solutions are used where complete maceration of soft tissues is required and are most appropriate for specimens that are to be mounted in resin-based mountants. In contrast, most gum-chloral mountants continue to clear specimens after mounting and thus gentler macerating agents can be used or, in some cases, very small insects can be mounted directly into the mountant without any prior clearing. After hydroxide treatment, specimens must be washed in a weak acidic solution to halt the maceration. Cleared specimens are mounted directly into gum-chloral mountants, but must be stained (if required) and dehydrated thoroughly prior to placing in resin-based mountants. Dehydration involves successive washes in a graded alcohol (usually ethanol) series with several changes in absolute alcohol. A final wash in propan- 2-ol (isopropyl alcohol) is recommended because this alcohol is hydrophilic and will remove all trace of water from the specimen. If a specimen is to be stained (e.g. in acid fuchsin or chlorazol black E), then it is placed, prior to dehydration, in a small dish of stain for the length of time required to produce the desired depth of color. The last stage of mounting is to put a drop of the mountant centrally on a glass slide, place the specimen in the liquid, and carefully lower a cover slip onto the preparation. A small amount of mountant on the underside of the cover slip will help to reduce the likeli- hood of bubbles in the preparation. The slides should be maintained in the flat (horizontal) position during drying, which can be hastened in an oven at 40–45°C; slides prepared using aqueous mountants should be oven dried for only a few days but resin-based moun- tants may be left for several weeks (Canada balsam mounts may take many months to harden unless oven dried). If longer-term storage of gum-chloral slides is required, then they must be “ringed” with an insulat- TIC17 5/20/04 4:39 PM Page 436 [...]... after death) Pin through the mid-line of the thorax between the wings, with the pin emerging between the first and second pair of legs (Fig 17. 2g); set the wings with the front margins of the hind wings at right angles to the TIC17 5/20/04 4:39 PM Page 439 Preservation and curation body (a good setting method is to place the newly pinned odonate upside down with the head of the pin pushed into a foam... groups, of the insects that are encountered; • no specialist with knowledge of the insects from the area in which your study takes place – as seen in Chapter 1, entomologists are distributed in an inverse manner to the diversity of insects; • no specialists able or prepared to study the insects collected because the condition or life-history stage of the specimens prevents ready identification There is... This guarantees the security of valuable specimens, and enters them into the broader scientific arena by facilitating the sharing of data, and the provision of loans to colleagues and fellow scientists 17. 3 IDENTIFICATION Identification of insects is at the heart of almost every entomological study, but this is not always recognized Rather too often a survey is made for one of a variety of reasons (e.g... series of questions, concerning the presence, shape, or color of a structure, which are presented in the form of choices For example, one might have to determine whether the 441 specimen has wings or not – in the case that the specimen of interest has wings then all possibilities without wings are eliminated The next question might concern whether there is one or two pairs of wings, and if there are... (phasmatids, phasmids, stick -insects or walking sticks) These are found on vegetation, usually nocturnally (sometimes attracted to light) Rear the nymphs to obtain adults, and remove the gut from all but the smallest specimens Pin through the base of the mesothorax with the pin emerging between bases of the mesothoracic legs, spread the left wings, and fold the antennae back along the body Phthiraptera (lice)... in another fluid prior to preservation (section 17. 2.2) Generally, 75– 80% ethanol is suggested for liquid storage, but the preferred strength often differs between collectors and depends on the kind of insect For detailed instructions on how to collect and preserve different insects, refer to the publications in the further reading list at the end of this chapter Archaeognatha (bristletails) These... pairs, whether one pair of wings is modified in some way relative to the other pair This means of proceeding by a choice of one out of two (couplets), thereby eliminating one option at each step, is termed a “dichotomous key” because at each consecutive step there is a dichotomy, or branch One works down the key until eventually the choice is between two alternatives that lead no further: these are the terminals... that the pin emerges between the mid and hind legs (Figs 17. 2a, 17. 3, & 17. 6a) Mount smaller specimens on card points with the apex of the point bent down slightly (Fig 17. 5b) and contacting the posterior lateral thorax between the mid and hind pair of legs Immature stages are preserved in fluid (stored in 85–90% ethanol, preferably after fixation in KAA or Carnoy’s fluid) 437 Collembola (springtails) These... if possible, most or part of the actual specimens from which the DNA is extracted For example, DNA can be extracted from a single leg of larger insects or, for smaller insects, such as thrips and scale insects, there are methods for obtaining DNA from the whole specimen while retaining the relatively intact cuticle as the voucher FURTHER READING Regional texts for identifying insects 443 Development;... ventilated, fireproof areas Collections of glass slides preferably are stored horizontally, but with major taxonomic collections of groups preserved on slides, some vertical storage of well-dried slides may be required on grounds of costs and space-saving Other than small personal (“hobby”) collections of insects, it is good scientific practice to arrange for the eventual deposition of collections into . when the apex of the point is directed away from the person mounting. For most very small insects, the tip of the point should contact the insect on the vertical side of the thorax below the wings base of the triangle and the insect is then glued to the tip of the point using a minute quantity of water-soluble glue, for example based on gum arabic. The head of the insect should be to the right. to avoid them contaminating other insects with their scales. For details of the use of other killing agents, refer to either Martin (1977) or Upton (1991) under Further reading. Dead insects exhibit