Tuatara: Volume 4, Issue 1, July 1951
The Collection and Preservation of Insects
The Collection and Preservation of Insects
Netting, Sweeping and Beating
An insect net has a more or less circular frame from which the net-bag itself depends and to one end of which is affixed a handle, usually detachable. For swift-flying insects such as dragon-flies, and the larger Lepidoptera, Hymenoptera and Diptera, the net should be of strong muslin or mosquito netting, about 18 in. in diameter across the frame and about 30 in. in depth, and with a long handle. For sweeping smaller and less strongly-flying insects from vegetation — grasses, sedges, bushes and creepers — a net made of more durable material is needed, and this need not be so wide or deep, nor the handle so long. For such work I have found very satisfactory a net 12 in. in diameter and 18 in. deep, made of white flour-bag cloth shaped and sewn together and atteached to a frame made of a metal rod about 3/16 in. in diameter bent to a circle and with the two free ends bent out to form a short handle (fig. 1). If desired, this can be fitted into the end of a wooden or even a metal tube handle. A net of this kind stands up to much rough use and can easily be mended if frayed or torn. It is particularly useful for insects like Hemiptera (bugs, leaf-hoppers), small beetles and weevils, and, among other Arthropods, bush- and grass-frequenting spiders, all of which are often caught in surprising numbers with a few sweeps. Instead of sweeping, the net may be held under the bush or tree, which is beaten with a stick. Such a net is well suited too for catching aquatic insects. Good hauls of water-bugs (Notonectids and Corixids) and water-beetles can be made by dragging it through the water or along the bottom of shallow pools. The tiny Veliids (Microvelia) can be scooped in large numbers with one edge of the net, and rapidly picked up with an aspirator (p. 15) as they run on to the drier parts of the net. If necessary, the inlet tube can be dried from time to time by pushing in a twisted cotton-wool plug. A mesh net has the advantage, particularly for large and active fliers, that the capture may be seen without opening the net, and secured by working a tube or killing-bottle down to it, but for rough sweeping in the capture of smaller and less active forms, the second type is extremely useful. Many small beetles ‘feign death’ and most bugs and plant-hoppers will walk up the net for some distance before attempting to fly or hop. If the net is gradually opened out from the top down and perhaps shaken down occasionally, these can be captured with an aspirator as they walk up.page break page 15
Nets of both kinds, especially if used for sweeping, should be attached to the frame by a fold of strong material, such as calico, to protect the edge of the net from wear. The net itself is cone-shaped, with the narrower closed end broadly rounded, and should hang freely without any constriction below the frame, and be deep enough so that when closed by twisting the handle at the completion of each sweep, there is sufficient overlap to imprison the insects. The frame may be of wood, metal rod, or spring steel (which can be looped to fold up the net into a very small compass) — the last material, however, tends eventually to snap; and it may be in one piece or of two or more pieces socketed, hinged, or screwed together so as to be disassembled or folded up. The handle may be screwed on or may be tubular at its end to fit around the end of the frame (e.g., a length of bamboo can be used in this way).
Many types of apparatus have been devised for catching specimens beaten from trees or bushes. One such consists of strong cloth sewn on to a wooden framework which can be folded up and when opened forms a rectangular tray, about 4 ft. by 2½ ft., sloping down gradually to the centre. A tarpaulin spread closely under the vegetation to be beaten will catch twigs and leaves shaken into the centre and concentrate the capture. An umbrella is a convenient and light instrument for catching beaten specimens.
Where possible, observations should be made and recorded of behaviour and biology. Once an insect has been located by sweeping, its host-plant should be searched for and an attempt made to observe feeding, and the part attacked (e.g., stem, leaf, seed-head) noted.
The Aspirator ‘Suck-Gun’
This consists of a tube or bottle with two holes bored in its cork (preferably rubber, to be quite airtight) for the insertion of two glass tubes as shown in fig. 2. Each tube is fitted with a length of rubber tubing to give range and flexibility; around the inner end of the suction tube is tied a small piece of fine gauze or muslin to prevent the entry of insects. The end of the inlet tube, which may be widened by splaying in a gas flame, is brought down close to the insect, which is sucked into the container by drawing air by the mouth through the suction tube. After a sufficiently large capture, the insects can be tapped to the bottom and an ordinary cork quickly fitted, the cork with tubes being put into a new container. The free end of the inlet tube is kept blocked between captures by a small cork, twig, or twisted plug of cotton-wool. Insects straying into this tube can be drawn back into the container by closing the inlet tube with a finger, then sucking on the aspirating tube and at the same time removing the finger. Such an aspirator is very useful for catching small insects in numbers from vegetation, the ground, or the sweeping net. Spring-tails (Collembola) and Thysanura should be quickly transferred to 95% alcohol.
The attraction of many insects to light can be utilized in their capture, e.g., a sheet of white material with a bright light behind it can be set up page 16 at dusk near the edge of the bush and the insects netted. A permanent light trap can be made having a bright source of light (electric bulb or fuel lamp) surrounded by sheets of glass forming an outer slope leading the insects to the light and an inner slope leading them into the killing-bottle, as shown in fig. 3. The plates of glass are supported by wooden triangles and the whole supported on a platform and covered by a roof. Details of construction of such a trap are given by Williams (1924, 1948), who recommends tetra-chlor-ethane in preference to potassium cyanide as a killing agent. A layer of Plaster of Paris is allowed to set on the bottom of the bottle and about a teaspoonful of the liquid poured on before use. The body of the jar should be as wide as possible; layers of crumpled paper help to separate the insects and minimize damage.
Miscellaneous Methods of Capture
Many beetles and their larvae inhabit timber, both rotting and sound, and flat-bugs (Aradids), beetles, cockroaches, etc. are to be found beneath loose bark. Turning over logs and stones reveals ground-beetles (Carabids), Staphylinids, earwigs, ants, etc. Leaf-mould is often rich in small beetles and bugs and Collembola. These can be caught by spreading out the debris on a tarpaulin or sheets of newspaper and collecting with a ‘suck-gun’, or by using a Berlese funnel (fig. 4). The heat, light, and drying out caused by the globe in the top of the funnel drive the insects down, to drop eventually into a tube of 95% alcohol. Under dry conditions, heating may not be necessary; if the layer of mould is thin, desiccation and positive geotropism will cause the insects to drop through the gauze. Salmon (1946) describes a convenient type of portable extraction apparatus for use in the field. The organic debris in large leaf-bases, such as those of nikau, usually reveals a variety of small beetles (Anthribids, Cryptophagids, Staphylinids, weevils, etc.), Collembola, and Anthocorids. Flower-heads house thrips and small beetles and attract numerous Diptera and Hymenoptera. On the surface of flood-water and in the debris stranded by floods are often found insects brought down from higher regions. Tufts of sedges, grasses and rushes in swampy areas shelter concentrations of insects after flooding. The bases of such tufts should be examined in any case, Hemiptera in particular being commonly found there. More or less open sandy or clay areas should be searched for tiger-beetles and Hymenoptera, which in warm weather dart about actively in search of prey, usually close to the ground. Piles of sea-weed and the sand beneath should be examined for beetles (Staphylinids, Histerids, Chaerodes), kelp-flies, and the littoral earwig (Anisolabis).
Killing agents commonly used include ether, ethyl acetate, amyl acetate, chloroform, and potassium or sodium cyanide. The first four are used in very small quantities — a single drop on the cork of a tube or on a wad of cotton-wool, and may conveniently be carried in the field in a tightly page 17 glass-stoppered drop-bottle. With small or delicate insects, like leaf-hoppers, leaf-bugs (Mirids), small flies, Psocoptera, care must be taken to avoid wetting. Exposure of the tubes to sunlight should be avoided as leading to evaporation and subsequent condensation of the killing agent and of water on the sides, to which the insects will stick, with probable damage to wings and other appendages. Such small insects are best either kept alive, with not too many together in a tube, and killed in the laboratory, or transferred soon after killing to dry containers between layers of cotton-wool or soft paper. Chloroform has the disadvantage of making many insects rigid and brittle. Insects killed with amyl acetate remain relaxed, and duplicates of heavily chitinised types, such as the larger beetles, may be kept stored almost indefinitely between cotton-wool impregnated with this agent. Small and soft-bodied insects, however, are liable to disintegrate with prolonged exposure. Potassium cyanide is used in a killing-bottle — a wide-mouthed, tightly stoppered jar with lumps of cyanide in the bottom covered with Plaster of Paris. Contact of the insects with this base is prevented by layers of filter-paper, while the addition of crumpled paper will serve to separate the specimens before death. Lepidoptera should preferably be killed in a separate jar. Cyanide has the disadvantages of being highly poisonous (the jar should be conspicuously labelled so in red) and of affecting the colours of many insects; the writer has long abandoned its use in favour of other killing agents. Young leaves of the bay laurel, usually grown as hedges, can be chopped up and used in a killing-bottle. The leaves emit hydrogen cyanide (the strong ‘bitter almond’ smell of which is apparent on crushing), and although comparatively slow-acting, have the advantages of lacking the danger of KCN and of keeping the insects relaxed. These must not, however, be left in long enough to develop mould. Larger Lepidoptera may be quickly killed or paralysed by squeezing the thorax; this prevents rubbing off of scales by movement against the container.
Insects from different localities or plant hosts should be kept separate and relevant observations kept in a note-book under numbers corresponding to those on the containers. On prolonged field trips the material should be transferred (at least once a day) to dry containers, preferably non-glass (cardboard or wood), between paper or cotton-wool, unless there are facilities for pinning-up in the field. Labelled match-boxes will hold large numbers of small insects; if there is danger of crushing, as in a pack, they can be stored in solid receptacles such as tins. A pack to accommodate collecting gear can easily be devised to meet individual requirements. General considerations are separate compartments for jars and tubes, to minimise breakage, and for full and empty tubes. It is an advantage to have a range of tubes of different sizes. Tubes in which many pharmaceutical and dental products are packed are useful.
Small and soft-bodied insects like Collembola, Thysanura, thrips, nymphs and larvae are killed and stored in 95% alcohol. Methylated spirits is not satisfactory, causing brittleness and eventual disintegration.page 18
The life-history and biology of comparatively few of our insects have been fully worked out, and there is plenty of scope here both for field observations and for rearing in the laboratory or insectary, and for the breeding out of parasites from such live material.
Mounting and Setting
Medium and large sized insects are mounted directly on pins. Entomological pins (anti-corrosive, of small diameter, and with round heads) are made in a range of sizes varying in length and thickness. The insect is best pinned while still relaxed. If it has hardened, it is placed in a tin or jar over a layer of damp felt, cotton-wool, or clean sand (with thymol or carbolic acid added) until relaxed. The insects are separated from the damp layer by card or a perforated zinc platform. The legs are extended and the pin inserted from above, through the middle thoracic segment (mesonotum) in most insects, through the anterior third of the right wing-case in beetles, and through the scutellum (thoracic shield) in Heteroptera. The insect should be mounted at about one-third of the distance from the top of the pin.
A setting board consists of a wooden base, some 14 in. long and 2 in. to 5 in. wide, covered with a layer of cork and this with two thicker cork layers leaving a groove ¼ in. to ½ in. wide between them, and the whole upper surface covered with white paper (fig. 5). The body of the insect lies in the groove and the legs are extended and held in place with pins or points (fig. 6). The wings are pulled into position with a fine needle or camel-hair brush and fastened down with card strips and points. The posterior margins of the fore wings should be at right angles to the body (fig. 7). The wings of beetles are not usually extended, nor are those of bugs, except that with the latter the wings of one side may be extended in some of the specimens of each species. Temporary labels should be added at the side, with details of date and place of capture, etc. The mounted insects are stored in a dry and well ventilated cupboard for 1-3 months, until completely set and hardened into position. The surplus pins and triangles are then removed and a small card label affixed below the insect, with date and locality of capture, name or initials of collector, host-plant, etc., written with a fine nib in Indian ink.
For insects too small to be mounted in this way, double mounting is resorted to. (1) The specimen is mounted on a micro-pin which is then stuck into a small strip of polyporus mounted on a larger pin, with the label affixed beneath (fig. 8). A number of specimens of the same species can be mounted on the one length of polyporus. These strips and the micro-pins (in a range of sizes) are obtainable from English dealers. (2) Alternatively, the insect may be stuck on its side to the apex of a small white card triangle by a drop of gum-arabic. The triangle and label are mounted as before on a pin (fig. 9). The older method, particularly for beetles, is to glue the insect to a card rectangle with the legs extended in position. Rows of such specimens can certainly add to the neatness of a page 19 collection, but the method has the disadvantage of obscuring ventral structures often used in classification, so that, if the initial identification is uncertain or needs revision later, the insects will have to be soaked off the cards and remounted. To a certain extent this can be overcome by mounting some specimens of each species on their backs. Double mounting has the advantage of exposing all aspects of every specimen to ready observation.
Genitalia and other mounts: The male genitalia are important features for specific identification in many groups. In such cases they should be dissected out and mounted. With larger beetles, the method described by Britton (1940, p. 473) can be used: ‘The male genitalia are perhaps most easily removed from dry pinned specimens by breaking the abdomen off by downward pressure at its apex. The abdomen is then placed in boiling water for a short time until the tissues are softened. An opening is made between the soft tergites of the upper side with a pair of needles, and the aedeagus removed through this. The aedeagus is then cleaned and mounted on a card attached to the pin, while the abdomen is gummed back in its place.’
With some insects the genitalia will need depigmenting by boiling for one to a few minutes in 10% KOH or NaOH solution. An easy clearing method, whether preceded or not by this treatment, is to place for a few minutes first in glacial acetic acid and then in clove oil, finally mounting in Canada balsam, either on a slide beneath a coverslip, the slides being labelled and stored, or, with larger genitalia, between two celluloid strips, which can be affixed directly on the pin beneath the specimen.
Small, delicate insects such as Collembola must be mounted whole on slides, after depigmenting, if necessary, by immersion in KOH or lactic acid. Into a gum arabic-chloral hydrate medium, such as that given by Womersley (1939, p. 291), they can be mounted directly or from water, acetic acid, or lactic acid. Salmon (1949) gives a detailed account of improved mounting methods for such small insects, including the use of media containing polyvinyl alcohol.
Storage and Preservation
Pinned specimens are best kept in specially made insect store-boxes or cabinets. The latter consist of a series of box trays, lined at the bottom with paper-covered cork and having removable glass lids. The former are hinged along one side, with both the lid and bottom section cork-lined and deep enough for insects to be pinned into both without the pins of one half damaging the specimens in the other. Failing either of these, or for temporary storage, cigar boxes or cardboard boxes can be used, lined with cork or other substance (such as pinex, cork linoleum) which will hold pins securely.
Before pinning in the insects, each box should be painted inside with a mixture to prevent ingress of insect pests (psocids, museum beetles, etc.), and allowed to dry in the closed position. Such a mixture is a saturated page 20 solution of camphor, naphthalene, or para- di- chloro-benzene in 1 part chloroform, 1 part colourless (medicinal) creosote, and 6 parts benzene (benzol). In addition, there can be pinned in one corner a small box of naphthalene or camphor, with thymol or carbolic acid added to prevent mould, and the contents held in place with cotton-wool. Many store-boxes have a small compartment for this purpose cut into one of the sides. If pests do gain entry, the box should be isolated and fumigated with carbon bisulphide. Insects infected with mould must be isolated and painted with an alcoholic solution of mercuric chloride. (Note that this substance is poisonous.)
When transferring pinned specimens, to prevent the pins from bending or springing they should be gripped below the insect with a pair of stout mounting forceps, with the ends broad, serrated, and bent out at an angle.
Each box or tray is to contain members of one group (order, family, subfamily, etc., depending on the size of the collection). The inside of the box is divided into a series of vertical rows and in each row the insects are arranged in species in transverse rows. A label with the name of the genus is affixed above the specimens and one bearing the species name beneath (in each case with the author's name), as in fig. 10. These labels are affixed by points — short headless pins — which can be made cheaply by snipping the heads of short, non-entomological pins.
A duplicate collection in 95% alcohol is useful. Small insects can be kept in tiny tubes of alcohol which are themselves immersed in larger containers of alcohol. This prevents drying out and obviates refilling. Larger insects, if required for subsequent dissection, may be kept in a form-alcohol solution such as Pampel's Fluid (95% alcohol, 15 parts; formalin, 6; glacial acetic, 4; and distilled water, 30 parts), or treated with Carls' Fixative (Tillyard, p. 488). Duplicates of dried material can be stored between cotton-wool or paper in labelled boxes well supplied with naphthalene, and these classified and packed into larger boxes. Insects such as dragon-flies and Lepidoptera can be stored in envelopes or folded paper triangles, large numbers of which will fit into one box.
For further details of collecting and preserving methods, the student is referred to the authors listed below and to the Auckland War Memorial Museum Educational Leaflets: Nos. 2 and 3.
Britton, E. B.— Trans. Roy. Soc. N.Z., 1940; 69 (4): 473. ‘The Carabidae (Coleoptera) of New Zealand (Part 1. Pterostichini).’
Hudson, G. V.— ‘New Zealand Beetles and Their Larvae’ (pp. 13-26), 1934 Wellington.
Salmon, J. T.— Dom. Mus. Records in Entomology, 1946; 1 (2): 13-18. ‘A Portable Apparatus for the Extraction from Leaf Mould of Collembola and other Minute Organisms.’
Salmon, J. T.— Trans. Roy. Soc. N.Z., 1949; 77 (5): 250-253. ‘New Methods in Microscopy for the Study of Small Insects and Arthropods.’
Tillyard, R. J.— ‘The Insects of Australia and New Zealand’ (pp. 485-500), 1926. Sydney.
Williams, C. B.— Bull. ent. Res., 1924; 15; 57-60. ‘An improved light trap for insects.’
Williams, C. B.— Proc. R. ent. Soc. Lond., 1948; 23: 67-85. ‘The Rothamsted light trap.’
Womersley, H.— ‘Primitive Insects of South Australia’, 1939. Adelaide.