FNZ 16 - Nepticulidae (Insecta: Lepidoptera) - Methods and Conventions
Donner, H; Wilkinson, C 1989. Nepticulidae (Insecta: Lepidoptera). Fauna of New Zealand 16, 92 pages.
(
ISSN 0111-5383 (print),
;
no.
16.
ISBN 0-477-02538-2 (print),
).
Published 28 Apr 1989
ZooBank: http://zoobank.org/References/9BE5D9B7-27E2-46F0-8A35-4D4912BA0D99
Methods and Conventions
Collecting. In the field the larva is the easiest life stage to collect, by searching for its mine - usually in leaves, but also in fruits and bark. More useful information is gleaned from a reared moth than from one collected at light.
The larvae are collected by picking the leaves they mine, and if the leaves are very small a piece of stem is taken as well. The leaves are put into plastic bags, which are closed and labelled. Each sample should be kept in a separate bag to avoid confusion should the larva vacate the mine before being sorted for rearing. In older collections one may often see series of mixed species, resulting from the collector not keeping his samples separate. Different plants of the same species, or even different parts of the same plant, may house different species of miner, so whenever in doubt separate plastic bags should be used. It is useful to add some sphagnum moss to the bags to keep the leaves fresh and free from mould, especially if the specimens are likely to remain in the bags for some days.
As soon as possible after returning from the field the living larvae should be separated from those which are dead or parasitised. Healthy larvae are bright green or yellow, in contrast with their brown or black colour when dead. Parasitised specimens become decreasingly active and stop mining. As the percentage of parasitised specimens can be very high (50%), allowance should be made for this when collecting. Under the microscope the parasite can often be seen under the skin of the larval host. In our work, parasitised specimens are retained for study of the parasite and the living larvae are divided; some are deep-frozen for biochemical studies, but usually the majority are used for larval examination and rearing to adults.
Rearing. Rearing Nepticulidae normally provides an accurate means of identification which may be otherwise impossible unless the male genitalia are dissected out. Rearing further provides additional data regarding food plant, mine, immature stages, and which males and females belong together. Similarity of external appearance amongst species, as well as the ease with which scales are lost during flight, makes identification from wing markings much less reliable.
We try to keep reared larvae in the same conditions (e.g., of temperature, day length) as in the wild, thus aiming at the natural time of adult emergence, especially if breeding is hoped for. However, if we know nothing of the climatic tolerances and life cycles of our material, which is so often the case with new species brought from other hemispheres and continents, we divide the larval samples and try different regimes.
Leaves containing larvae are placed in glass jars with slightly damp earth and sphagnum moss to keep them fresh. When the larvae have left their mines and spun cocoons, the leaves are taken out to prevent mould from forming. If they are still in good condition they are then dried for the herbarium. Temperature is often an important factor. In summer broods, adults may emerge within a couple of weeks owing to warm temperatures. The pupal stage is lengthened by colder temperatures, and with over-wintering pupae sometimes sub-zero temperatures are essential to initiate adult development (CW). Pupae can be 'forced' by reducing the period of time they are kept in the cold, whether outside or in the refrigerator, but normally 1 month is optimal; the temperature can then be gradually increased.
To rear cocoons in natural conditions they are enclosed in stainless steel tins with gauze lids at both ends. These are buried just below the surface of the soil in autumn and dug up again in spring. Black polythene sheeting is placed over each end of the container to make it totally dark inside. A 3 x 1 cm glass tube connected to a hole in the side of the tin provides the only light source. On emergence the adults make for the light inside the tube, where they can be easily removed. This reduces the risk of losing specimens, which if the lid had to be removed would certainly occur.
Preparation. It is important to set the moths as soon as possible after emergence, because they quickly lose their scales when flying. The moths should be anaesthetised but not killed before mounting; after death they stiffen too quickly, and cannot easily be relaxed. We enclose the moth in a small tube and anaesthetise it by dampening the cork with a little ethyl acetate.
The small size of nepticulid moths demands special mounting materials and techniques. The setting boards used by us are small wooden blocks approximately 10 × 3 × 2 cm with a central groove 0.8 mm or 1.0 mm wide filled with polystyrene and balsa strips glued on either side. The moth's body is pinned into the groove, by means of a minuten pin (size 0.10 mm, 0.15 mm, or 0.20 mm) placed through the thorax. The wings are spread by means of gentle blowing and fine needles. They are held in place with strips of transparent paper 5 mm wide and pinned with the same fine pins outside the edge of the wings.
The board is placed in a killing jar with ethyl acetate vapour until the pinned moth is dead. When removed from the jar, the moth is left on the board to dry and set for about 10 days at 30°C. It is then taken off the board, placed on a strip of polyporus with a No. 3 pin, and labelled.
Genitalia preparations. For removing the abdomen we found it advisable to place the moth in a small box, the sides of which are higher than the pin. This prevents the abdomen from flicking away while it is being loosened. With a needle or fine forceps the tip is gently pushed downwards until the abdomen breaks off between the thorax and first abdominal segment. It is then placed in a small test tube with a little 10% potassium hydroxide solution. The tube is labelled with a number corresponding to that of the specimen - especially important if several slides are prepared at the same time.
A loose-fitting cap placed over the tube prevents loss of the specimen whilst allowing changes in pressure during heating. The tube is placed in a water bath and heated to 85°C for 25 - 50 minutes to disintegrate the internal membranous structures. The abdomen is then rinsed in distilled water. While still immersed, unwanted internal structures (e.g., gut, trachea) are cleaned away by drawing them out of the abdomen with hooked or straight minuten needles mounted in holders. The outer surface of the abdomen is cleaned of its scales to reveal abdominal structures which may be present.
A complete dissection of the male genitalia involves removal of the capsule from the abdomen and withdrawal of the aedeagus anteriorly from the capsule. However, a number of workers find this prolonged and exacting process unnecessary. For identification purposes, at least, the aedeagus can often be left in situ, and some workers feel that it is only necessary to evert the capsule from the surrounding abdominal membranes. In the female the membranes between segments 8 and 9 are carefully parted with forceps, and the bursa copulatrix and accessory sac are drawn out with the terminalia from the ensleeving abdomen, which can now be further cleaned. The bursa may also have to be emptied through a small cut.
There are several suitable stains. For males an aqueous solution of haemaluin is very good. Staining time varies according to the strength of the solution, but 3 minutes is usually sufficient. Females are often stained in Chlorazol Black E, which imparts colour to the preparation ranging from black through green and blue to yellow, depending on the degree of sclerotisation. Care must be taken not to overstain, or detail will be obliterated by too much black. Chlorazol can be used in aqueous solution, but more often it is dissolved in 70% ethanol. An aqueous stain is used after cleaning and before passing the genitalia through increasing concentrations of alcohol. In the case of Chlorazol in 70% ethanol staining is carried out when the 70% stage is reached in the alcohol series; the preparation is washed in 70% alcohol before and after staining.
After two washes in absolute alcohol the preparation is ready for mounting. Our mounting medium is Euparal, two separate drops of which are placed on the glass slide. The abdomen and genitalia are placed in them with fine forceps, ventral side up. The abdomen is normally placed to the right of the genitalia, and the aedeagus below the male capsule. The slide is left flat to start drying for 12 hours, after which time the genitalia and abdomen will stay in place, so another drop of Euparal can be added and the coverslip placed on top. A good way to avoid catching air bubbles is to place the coverslip on the edge of the table and tilt the slide down on to it. The slide is now placed in the oven to dry.
Slide labels should carry essential information from the data label, together with the slide number, species name and/or number, sex, host plant, mounting medium, date, and name or initials of the preparator. Often two labels will be needed for each slide; the layout should therefore be standardised.