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Collecting and preserving chalcidoids

Introduction | Collecting methods | Separating material | Relaxing specimens | Drying specimens | Card mounting | Slide mounting | Storing and preserving | Preparing for S.E.M. | Mailing to specialists | Shipping in alcohol | References

Collecting methods - continued

(2) Beating
This method entails holding a beating tray (white canvas stretched over a 1-metre square frame or even an inverted umbrella) under a bush, branch or other suitable habitat, and then sharply hitting the branch with a heavy stick to dislodge any insects. These fall directly into the tray and can be collected using an aspirator or fine paint brush dampened with alcohol. This method has an advantage over collecting with a sweep net in that it may be used for thick heavy vegetation unsuitable for sweeping (eg very thorny bushes, or even bare branches). The disadvantages are obvious. The catch is substantially smaller because many insects escape, and it can only be used if the vegetation is reasonable high. Beating is best done in cool weather or early or late in the day when insects are least active

(3) Pyrethrum spraying
This is a method of obtaining insects from dead or rotten wood or from habitats unsuitable for sampling by other means. It can also be used to collect insects from small bushes or single branches. A polythene sheet is spread under a piece of vegetation or rotten wood which is then sprayed with a pyrethroid (sold for domestic purposes as an aerosol fly-killer, eg RAID®). Dying insects, as well as those falling out of crevices or holes in bark, etc. will fall onto the sheet and can be collected using an aspirator, fine brush or forceps, or funneled into a into a suitable receptacle. This operation must obviously be undertaken when there is no breeze. Other disadvantages are that smaller individuals may get trapped in minute droplets of spray on leaves, bark, etc. and specimens may become stiff and difficult to relax, which hinders mounting.

(4) Canopy fogging
This is a development of the aerosol spraying method described above. It is especially successful for sampling taller trees or the canopy of tropical forests. It is one of the most productive methods of collecting many groups of chalcidoids, eg Eupelmidae, Encyrtidae, Aphelinidae, Signiphoridae and Trichogrammatidae). A variation of the method has been described by Davis & Stork (1996) and involves knocking down insects from the canopy using a pyrethroid insecticide transported by means of a heated oil carrier. This is produced using a two-stroke insecticide fogging machine. As the carrier is warmer than the surrounding air rises by convection into the canopy even if released from ground level and the canopy is 40m high. Falling insects are collected by means of canopy funnels or sheets slung between trees or from ropes. A suitable collecting pot is positioned in the centre of the funnel or sheet. This is separated from the funnel or sheet by a 1cm wide strip of fine mesh that is fine enough to allow alcohol to pass through but not the smallest Hymenoptera. After an hour or so the insects that have fallen onto the funnels are washed into the collecting bottles by means of a fine spray of alcohol administered by a knapsack sprayer or alcohol squirted from wash bottles. To prevent wastage, a low concentration of alcohol (10-20%) can be used for this operation, but it must be replaced as soon as possible with 70% alcohol in the collecting bottles.

(5) Rearing
This is probably the most rewarding method of collecting chalcidoids, since the biological information gathered may prove of great value to the taxonomist as well as to those working in biological control. The main drawback is the considerable time and effort that is required to locate suitable hosts. Also a lower diversity of species can be collected in any given time compared with that obtained by sweeping or other sampling techniques.
Most immature and mature stages of the higher orders of insects provide hosts for chalcids, eg Hemiptera, Coleoptera, Diptera, Lepidoptera. Some chalcids are phytophagous, and many are associated with the reproductive organs of pines, fruits of various flowering plants, flowering heads and stems of various grasses and also fig fruit (Ficus spp.). The latter usually harbour a particularly rich chalcid fauna that can be reared easily (see Boucek, et al. 1981). Whatever likely hosts are collected it is best to put them in a suitable receptacle to await the emergence of the parasitoid(s), eg glass tube with a cotton wool plug, brown paper bag, gelatin capsule, polythene bag. Parasitized hosts can often be distinguished from healthy hosts by their slightly different colouration or behaviour. Parasitized hosts may be darker than healthy ones or may move at a different pace or in a different way from healthy ones. On rare occasions it may be advisable to put vegetation harbouring hosts into an emergence box.The emerging insects are attracted to the light and collected into a glass tube placed over an inverted glass filter-funnel placed on the roof or sides of the box. This prevents return of the insects into the box. One disadvantage of the emergence box is that a fairly large proportion of the parasitoids may not find their way into the collecting tubes and therefore it should only be used where a high parasitoid infestation is likely. Another disadvantage is that there may be several possible hosts in a sample from which the parasitoids are reared. It is essential that wherever possible suspected parasitized insects are individually segregated to reduce the risk of specimen being labelled with incorrect and often misleading biological information. It is very easy to put an aphid mummy, attached to a small piece of leaf, inside a glass tube and assume that a chalcidioid found wandering around the inside of the tube has come from the aphid whereas it actually emerged from undetected agromyzid puparium in the piece of leaf. Parasitoids found in a tube containing coccids on a twig may actually have come from lepidopteran or heteropteran eggs intermixed with the coccids. Where possible, parasitized hosts must be isolated from plant or other material to prevent erroneous records. Also the remains of the host must be retained together with the resulting parasitoid, perhaps kept in a suitable gelatin capsule pinned with the specimen or even glued on the same card as the specimen.

(6) Malaise trap
Together with the advent of Critical Point Drying to dry specimens from alcohol (see below), the use of Malaise traps for collecting chalcidoids has probably revolutionised the study of chalcidoids.

A Malaise trap is used to collect large number of flying and occasionally flightless insects. A suitable design for collecting chalcidoids has been described by Townes (1972). The mesh of the net must be fine enough to prevent smaller chalcidoids from passing through easily. A Malaise trap, if correctly constructed and sited, will provide a representative sample of chalcids to be sound in an area and will collect even the smallest insects, eg Megaphragma spp. The trap may be run dry with an insecticidal fumigant in the collecting pot but in this instance the material may be damaged by other dying insects. They may also be covered in moth scales and nany insects may not even reach the collecting pot. However, if 70-80% alcohol is used as the killing agent, the size of the catch will be greatly increased, possibly because there is a certain amount of attraction to the alcohol itself (many insects are associated with rotting or fermenting fruit). Two advantages of the Malaise trap are that it need only be visited once every week or two weeks for emptying and it can be serviced by a non entomologist. It is, however, wise to empty the trap more frequently than once a week to ensure that spiders have not spun webs in the entrance to the collecting head, and to check that the trap has not been fallen down or damaged. In hotter areas smaller insects may deteriorate rapidly in warm alcohol (>25°C) for more than a few days. The fact that the catch of chalcidoids collected into alcohol is usually larger, more diverse and cleaner than that obtained using a fumigant, outweighs the preference for dry-collected specimens (see section on preservation).

(7) Flight intercept trap
This is a type of trap much favoured by coleopterists (see Peck & Davis, 1980; Masner & Goulet , 1981; Davis & Stork, 1996), but it is also an excellent way of collecting microhymenoptera. The trap consists of a 1m high, 2-3m wide length of black terylene netting slung vertically between two trees or posts. It is protected from rain by a transparent polythene roof that extends about 0.5m either side of the vertical netting for at least its length. A number of pans are placed beneath the netting for the whole of its length. The pans contain water with a drop or two of detergent, a saturated salt solution, a 50/50 ethylene glycol/water mix or some other suitable collecting medium (see comments below under yellow pan traps. Flying insects hit the vertical net and fall into the pans. They can then be collected at regular intervals using the same method described below for yellow pan traps. The catch can be increased by spraying the netting with a long-lived contact insecticide such as Ambush®.

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Last updated 19-Aug-2003 Dr B R Pitkin