Landcare Research - Manaaki Whenua

Landcare-Research -Manaaki Whenua

FNZ 17 - Mymaridae (Insecta: Hymenoptera) - Collecting and Preservation

Noyes, JS; Valentine, EW 1989. Chalcidoidea (Insecta: Hymenoptera) - introduction, and review of smaller families. Fauna of New Zealand 18, 96 pages.
( ISSN 0111-5383 (print), ; no. 18. ISBN 0-477-02545-5 (print) ). Published 02 Aug 1989
ZooBank: http://zoobank.org/References/1D0A405D-6643-42DB-B911-80664BC6F853

Collecting and Preservation

Perhaps the main difficulty in the study of mymarids is their small size and thus their relative unpopularity with insect collectors. Therefore, a common complaint of the serious student of this group (or indeed of any microhymenoptera) is that there is not adequate material available for study. The following methods are the most rewarding for collecting and preserving mymarids. For greater detail on collecting and preservation, see Noyes (1982).

Collecting methods

Malaise Trapping. This is probably the best single method for collecting mymarids, since even species of small size that are rarely collected by any other means can be collected in quite large numbers, e.g., Alaptus spp. A design for an efficient trap for collecting chalcids has been described by Townes (1972). The advantage of a Malaise trap is that it can be set up and serviced by anyone. It can be emptied relatively infrequently - at fortnightly intervals if ethyl alcohol is used as a killing and preserving agent. The use of ethyl alcohol is slightly disadvantageous because material is collected wet (see 'Preservation', below), and the subsequent sorting of required specimens from the catch takes considerable time. It is possible to collect dry using fumigants such as Vapona®, but the catch is greatly reduced and the material is often covered with moth scales and damaged by dying insects.

Sweeping. Sweep-net collecting is a satisfactory method for collecting larger mymarids, since a large number of species may be collected in a short time. The insects can be removed from the net using an aspirator, and killed by placing a plug of paper tissue or cotton wool soaked in ethyl acetate in the entry tube of the aspirator. Before any insects are sucked into the aspirator it is advisable to place in it a crumpled sheet of soft tissue to prevent insects from sticking to the sides of the tube. It is important to keep the specimens as dry as possible.

The sweep-net is most efficient if the handle is about 1.2 m long and the head triangular, with the handle joining the head in the middle of one side of the triangle. The triangular head allows the net to be used most efficiently on grassland, i.e., more of the net will be in contact with the ground during sweeping. The relatively long handle allows the net to be held as far away from the body as possible, so that insects are not forewarned by footfalls, etc. It also allows for greater reach when collecting in forest habitats. The best material for the frame is aluminium, since this is sufficently rigid yet light enough in use. The net bag should be of a material which is strong and durable, yet allows easy passage of air; canvas is not suitable.

When sweeping grassland it is very important to sweep in long arcs, keeping the head of the net in contact with the ground for the entire arc by pressing down on the handle.

Yellow Pan Trapping. This is a good method of catching mymarids. The trap consists of a tray about 30 cm square and 5 - 8 cm deep, painted yellow on the inside. Insects are attracted to the yellow colour and drown in the collecting medium contained in the pan. The collecting medium may be water with a few drops of detergent to break the surface tension, or water plus ethylene glycol (1:1), or a dilute solution of picric acid, or a saturated solution of common salt. If water is used the trap must be emptied at least once a day or the material will deteriorate very badly. With other media the trap can be emptied weekly (weather permitting). Specimens must be rinsed well in clean water before transferring them to 70% ethanol. Ethylene glycol is not recommended because the subsequent slide-preparation of material may be difficult; the specimens tend to collapse badly on transfer to balsam. Picric acid causes the gaster to distend (this is not often important taxonomically).

Pan trapping is ideal for collecting from among trees and in forest where there is little undergrowth.

Rearing. Rearing is the most rewarding method of obtaining specimens, since much can be learned about their biology. A limitation is that much effort has to be put into locating possible host eggs and rearing mymarids from them. A good method for circumventing this is to use the eggs laid in the laboratory by a potential host species. These can then be placed in suitable habitats outdoors and returned to the laboratory after a suitable period of time to await the emergence of parasitoids. Potential host eggs should be kept in a container appropriate to the size of the sample and examined regularly for the emergence of mymarids. Care should be taken to ensure that the correct host is recorded. It is very easy to collect an egg on a small piece of leaf and assume that a parasitoid found wandering around or dead in the tube comes from the egg, whereas in fact it may have emerged from some other host such as an agromyzid pupa which may have been overlooked. Rearing from individually isolated hosts can overcome this problem.

Suction Trapping. The suction trap has a single advantage over a Malaise trap in that it actively sucks in small flying insects. It has many disadvantages, e.g., it needs an electric power source, is much more expensive, and fails to catch flightless insects. Even so, catches from a suction trap should not be overlooked since they may contain many interesting species.

Extracting from leaf litter and moss. This is a good method for collecting some Mymaridae which are rarely collected by other methods. Specimens can be collected from leaf litter or moss using a Berlese funnel or an emergence box.

Preservation

Material collected dry should be kept dry. Specimens that are not mounted within 2 hours of killing in ethyl acetate vapour should be relaxed before mounting, or layered between sheets of cellulose wadding (or even a couple of layers of soft toilet paper), and stored in a strong, dry, airtight box. A crystal or two of thymol should be added to inhibit the growth of mould. Material collected into alcohol, e.g., in Malaise traps or yellow pan traps, should be dried as soon as possible. This can be done by air-drying the material on absorbent card (see below under 'Mounting material'). Unfortunately, air-dried specimens usually collapse or shrivel upon drying. This can be prevented by dehydrating specimens using a critical-point drier (see Gordh & Hall 1979). Dried specimens can be stored between sheets of tissue paper or in gelatin capsules, held in place with finely teased cotton wool. Specimens which have been kept in alcohol for a long time (5 years or more for specimens less than 1 mm in length) are normally unsuitable for making slides.

Mounting Material. Mymarids are best mounted on rectangular cards using a water-soluble glue. The specimens should be mounted with the vertical axis through the thorax at about 45° to the plane of the card, preferably with the wings, legs, and antennae displayed and the wings and head free of glue - see Noyes (1982) for details. This method requires a good deal of practice, but has advantages over card-point mounting in that specimens are well protected, and the various parts are much easier to see against the white background of the card.

Specimens from alcohol should be dried on a piece of moderately absorbent card, e.g., Bristol board. The insect should be placed in a drop of alcohol on the card with its wings flat against the card. As soon as it is dry, in 5 - 25 seconds, it should be removed and mounted.

Relaxing specimens. Specimens that have been dry for some time become extremely brittle, and should be softened to prevent breakage of appendages. This is best done by placing them on a piece of tissue on a glass dish inside a plastic box. Put in a few drops of water or glacial acetic acid (not more than 1.5 ml per 0.75 1 of box) and leave for 8 - 24 hours. If material is layered, put it in the box still in layers, otherwise it may be damaged when removing the top layer. If the material was killed in ethyl acetate vapour it should be sufficiently relaxed to be mounted without any damage whatsoever.

Slide Mounting. It is necessary to make good slides of at least some specimens from a series in order to see characters of taxonomic value.

  1. Temporary slides. These usually entail mounting whole specimens in a water-soluble medium such as Hoyer's or Berlese. It is not recommended, and should be used only if there is a surfeit of material of the one species available. Its main (and probably only) advantage is that slides are relatively quick and easy to make.
  2. Permanent slides. It is essential that slides be made using Canada balsam if material is of taxonomic value, or if only a limited amount of material is available. This method is laborious and time-consuming, and requires a good deal of practice to master it. Preferably, body parts should be mounted (after clearing in 10% potassium hydroxide solution) under four or five separate 6 mm coverslips. For a detailed description of the method, see Noyes (1982).
Briefly, the method is as follows. Remove the wings and place them in balsam. Clear the specimen in 10% KOH at 20°C for 24 - 48 hours (if it has never been in alcohol), or at 20°C for 72 hours, or at 20°C for 24 hours followed by 40°C for 24 hours (if it has been in alcohol). Next, neutralise the preparation in glacial acetic acid for 10 minutes, followed by distilled water, then dehydrate through 35%, 70%, and 95% alcohols, (each for 10 minutes), and finally clear in clove oil (or terpineol warmed under a bench light) for 10 minutes. Position the body parts in individual drops of balsam on a slide and keep in an oven at 40°C for 4 weeks; this fixes them in position. Finally, add the requisite amount of balsam to each part (smallest amount possible for wings and antennae, and largest for thorax and gaster), and place a coverslip over each part of the preparation. NOTE: the balsam should be relatively thick, i.e., if a pin is put in it and pulled out the balsam should form 'strings'. If the balsam is thinner it will be difficult to position the coverslip flat; the balsam will also contract as it dries, crushing the part that it covers.

Data Labels. All material should be labelled adequately with at least collection locality, date, and host data (if reared). Never use code numbers alone and keep the data separate; data may be lost or mislaid in this way.

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