Invented by the Swedish entomologist Rene Malaise, the design is remarkably simple and is based on the shape of an old-fashioned two-man tent. The legend goes that when he returned to his camp after a hard day’s collecting, he noticed that there were more insects in his tent than he had caught with his butterfly net. He immediately saw the potential and decided to make a trap that exploited the design of his tent. Since then entomologists (most notably Henry Townes) have fine-tuned the original design.
The trap is made from fine netting, ribbon loops and guy-ropes – all supported by a 2m wooden pole and some strong tent pegs. The collecting vessel attaches to the tent using a specially made metal bracket, held in place by a metal ring-fastener. In the above photo you can see the black walls topped with a white roof leading up to the white collecting bottle, attached to the top of the 2m pole. The trap packs down very small and the wooden poll can be cut from any surrounding vegetation. The only equipment needed is a hand-saw (for the polls) and a screwdriver (to tighten the ring fasteners).
The net is erected at 90 degrees to a natural insect flight line, like a hedge, woodland ride or fence-line (like the example to the right). You should make sure that the pitch of the roof slopes upwards and that any creases in the fabric point towards the highest point. A good tip is to point the high point towards the brightest part of the sky – in the northern hemisphere this is due south. Though you can get away with any direction if the lower end is up against a shady hedge.
I have found that the guy ropes can be pegged directly into the ground but some people like to cut short sticks to hold the line up high. This keeps the roof as high as possible and opens up the sides of the trap to maximise the trapping area. In this photo I have used a stick to hold up the rear guy-rope.
A flying insect hits a vertical sheet of netting and instinctively flies up towards the light where a tilted, pitched roof guides it towards one end, where a hole in the netting allows it to pass into the collecting bottle.
The collecting bottle can either be left dry or can be filled with a killing and preserving fluid – like alcohol or water & anti-freeze. I prefer the former because I have found that anti-freeze can leave a greasy deposit on small specimens.
I usually leave the trap running for about two to three weeks at a time but I like to change the collecting bottle every 3 days to prevent the alcohol dehydrating the softer-bodied specimens. The trap is left to collect 24-hours a day in rain or shine – but sunny weather is obviously more productive.
The catch is usually taken home and examined under a relatively low-power (10x) binocular microscope. I remove all the specimens I want to mount up using fine-pointed tweezers – starting with the soft-bodied ones. The remaining specimens are left suspended in alcohol for future reference and for distribution to other experts, The alcohol will prevent decay and keep them fresh for many years – but over time it can discolour the soft-bodied insects.
A Malaise trap is used to ascertain the species diversity on a particular site and, being a ‘flight intercept’ trap, it is particularly good at catching species of flying insect. In my experience the main insect orders are caught in the following proportions:
We have added a huge number of species to our list using Malaise trapping and have managed to re-confirm some sightings seen a long time ago.
Removing specimens from alcohol
This next bit is a response to several questions I have been asked about my methods for dealing with ‘wet’ specimens (i.e. things that have been stored in alcohol).
Some groups like Ephemeroptera (Mayflies) and Trichoptera (Caddisflies) are best stored in alcohol to preserve their soft body structures and can be identified while still wet. However, this is not true of the majority of insects groups like Diptera (flies) & Hymenoptera (bees, wasps, ants, sawflies & ichneumons), where body structures are relatively robust and the identification keys often rely on viewing a dry specimen. This is because these groups are often keyed using fine hairs, subtle colouration or fine dusting, none of which are easy to view on a wet specimen.
Anyway, the transition from wet to dry can be fraught with problems – for instance if you take a small specimen out with a pair of tweezers the first problem you get is that the wings will usually collapse under the surface tension of the liquid! When these specimens dry the wings can be deformed and render the specimen useless. To get over this pour the specimens+alcohol into a petri-dish or do as I do and just move them to a dish of alcohol one by one as you go through the Malaise catch. Then tear up small pieces of filter paper and slide one into the dish. Under the microscope gently drag the specimens onto the filter paper and when you have loaded it up gently lift it out with tweezers allowing the surplus alcohol to run off. If you have done it right the specimens will be left on the papers and the specimens will be dry in about 10 minutes at room temperature. [This tip is courtesy of Dr Donald Quicke, Imperial College, Silwood Park, Ascot]
Another problem you may have is that the softer-bodied insects often get very dehydrated when left in alcohol for long periods. When the alcohol dries out the eyes & abdomen can collapse and render the specimen unidentifiable. This is a more tricky problem but you can avoid it by reducing the time the insects are exposed to the alcohol and by reducing the concentration of the solution. IMS (Industrial Methylated Spirit) is 95% alcohol and this preserves things for very long periods but I usually get away with diluting this to 70% by adding distilled water. This slows down the dehydrating effect but to improve my chances even more I try and empty the trap every 3-4 days and I make sure I go through the catch that evening to remove the soft-bodied things.
Things that have been in alcohol for periods of months or years are obviously going to need different treatments but there are apparently ways of re-hydrating specimens or treating them to avoid the shrivelling effects. The obvious one is to move them to very low concentrations of alcohol over a period of time and then hoping the water has permeated the tissues enough to allow them to dry in a natural shape. This can be difficult though because in a non-sterile environment the specimen may start to decay. Another method is to transfer them directly to ethyl acetate for a few hours and then dry them. This is apparently a very good method but it leaves the resulting specimens very brittle so you must take great care with them afterwards.
Availability of chemicals:
None of these chemicals are hazardous if used correctly in a well-ventilated room but always read the labels and documentation that come with them for up to date safety information.