Beetles from extraction samples
Introduction
This is perhaps the most versatile way of finding beetles because it can produce excellent results at any time of year and it can be used to find specimens from any habitat or location. Extraction can be used to find beetles among samples that are otherwise difficult or tedious to work and it can be save a great deal of effort. Because adult beetles are generally inactive through the colder months this is often the way of finding and dealing with them e.g. from tussock or moss samples, and it affords an excellent way of dealing with opportunistic sampling e.g. samples taken from rot-holes etc. in trees blown over during winter gales. These samples can be processed fairly quickly and, using the correct methods, no specimens will be overlooked. This last point is important because many people have a go at sampling and quickly give up because they find either very little or nothing at all, but with the correct processing every last specimen can be recovered very easily and fairly quickly. Admittedly many samples are disappointing, especially when they contain apparently good material, but very good results will quickly occur, these will provide new species and will more than make up for the odd dud sample. It is probably fair to say that the method will be used much less frequently during the spring and summer, obviously because there are so many other things going on in the beetle world, but more especially because results tend to become disappointing in the spring when everything wakes up and disperses away from their overwintering sites. On the other hand some samples such as recently vacated bird and mammal nests cannot be passed over. A great advantage of using extraction techniques is that they can be very lazy; samples can be found very quickly during the winter without travelling very far from transport, they need not be very large and so within a short time a few likely-looking samples can be taken and brought home to be dealt with in comfort. Collecting samples is the first part of the exercise and in many ways the most critical because until an awareness is developed there will be a temptation to collect just about anything and throw it through the extractor. As stated above, the best time to collect samples is during the winter because beetles tend to be active at other times and difficult to find and deal with. We usually restrict ourselves to whatever can fit into a polygrip bag about 20 cm wide, there are easily large enough to take tussocks or decent samples of moss or litter etc., and several can be carried with ease.
Burlese funnels can be used to extract beetles from various different samples
Samples of dead fungus, dry dung or compost can be very rewarding at any time of the year, and recently dead bark along with all the detritus that occurs around it can be very good during the summer. Moss growing on logs can be scraped off with a trowel but it is better to take a sample still attached to the wood and extract the lot, the same applies to arboreal fungus. Tussocks are better taken with a trowel, along with a small sample of soil, rather than cutting them off with a saw. Experience will soon show what the best samples are and how to collect them.
Method
Having taken samples they will need to be dealt with and most people will only be able to deal with one at a time. Each will take about 24 hours to process, only very wet samples taking longer, and so the others will need to be kept sealed for a few days, I use a fridge but this might be considered unsociable by some, especially as they often contain (to be kind) decaying organic matter. But this is usually done in colder weather and so they could be left in a shed or even outside in a protective container that is not exposed to the sun. I have often been asked whether specimens will die through suffocation under these conditions but, so long as the temperature is low, the chemical oxygen demand will be very low and the biological demand will be similarly low so long as the sample is kept cold. Obviously if a sample warms up the bags should be opened periodically or dealt with quickly, but in our somewhat vast experience specimens kept in a fridge for a few days soon respond to heat when put into the extractor. But samples should really be dealt with as quickly as possible, not simply because the specimens within need to be kept alive but also because with a little experience and a few good results the extractor will need to available as soon as the next batch of samples is collected. Also to be considered, and better to do this beforehand, is that samples can become rather aromatic during the extraction process, which is to say, from the non-coleopterists point of view, really foul or offensive. The majority will be ok but beware samples of fungi or dry dung can be a bit strong, from the non-coleopterists point of view that is, and doing the actual extractions in a shed can be a real bonus here. So far, so good; getting hold of and keeping good samples ready for extraction is easy and gets easier and better with experience, but before the next procedure is explained, a brief look at extractors might be useful.
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​Purpose made extractors are readily available, these are usually called Berlese funnels or Tullgren funnels, they are safe to use and delightfully easy to work with but they tend to be expensive. The basic principle is that a sample is slowly and gradually heated so that the occupants become active and move away from the heat and desiccation. Heat is supplied by a low-wattage light bulb suspended above and close to the sample, if the bulb produces too much heat the sample will warm too quickly and the specimens can become ‘cooked’, especially in very wet samples. The ideal set-up uses 15 or 25 Watt bulbs suspended an inch or two above the sample. It is a good idea to obtain a supply of bulbs because they tend to have a short life in enclosed extractors, in our experience they often die after as few as ten extractions. Any specimens upon or near the surface of the sample will move down away from the light then continue to move down as the sample heats up. Ideally the sample will rest on a circular piece of mesh made from thin wire placed near the base of metal or plastic funnel, the spout of which has been removed. The specimens eventually fall from the funnel into a collecting vessel placed directly underneath the opening. This is important because if there is a gap between the opening and the collecting vessel many specimens tend to avoid it and fly off. This set-up is housed in a bucket so that there is no gap at the top through which any specimens can escape. In commercial extractors the light is housed in a lid which fits snugly over the bucket and the funnel and the whole thing is sealed, except for a few ventilation holes in the lid above the bulb. Because of the expense many people will devise their own extractors, often with a light suspended above an open sample in a funnel which is placed on the rim of a suitably-sized container. These invariably work well but it should be remembered that the combination of electrical fittings, wet samples and a container of collecting fluid should be treated with great caution. Two aspects of the extractor need further consideration. Rather obviously many of the modern plasma light bulbs are designed to save energy and so produce little heat. Enough said. The collecting vessel should be small, we use a heavy glass tumbler, and it needs to contain liquid to trap the specimens; water is okay but many specimens will swim to the edge, climb out and wander about the bucket. Alcohol is good if samples cannot be dealt with quickly as they can be tubed and stored, but for samples that will be examined as soon as they are removed the best thing is water to which a drop of detergent is added. If water alone is being used it is probably best to add a few small twigs to which the specimens can cling. In dilute detergent most of the specimens will drown but a surprising number will survive and remain within the container. There are no doubt many ways of dealing with samples but the following works well, at least for us. Samples vary greatly in appearance; some include clay or silt and can be cloudy while others are clear with only a few bits of floating and sunken debris, and samples from heart wood, rot holes or bark debris can include lots of particulates that completely obscure any detail. This is the stage that puts many people off extractions because it is often, or usually, difficult to see anything among the debris; a few years ago I was asked to help somebody with sampling because the results were so consistently poor and it soon became obvious why; the sample was worked over a bucket of water and specimens were taken from the surface with a small brush and put into individual tubes, and once no more specimens were obvious the samples were being discarded. The following will avoid such nonsense.
Remove the tumbler from the extractor and then carefully examine the bucket for any escapees, staphylinids in particular can be very adept at escaping the sampling vessel and may readily take flight when the extractor lid is removed so be careful here. Take a length of very absorbent tissue and fold it a few times so that it ends up about 100 mm square, then place it near to where the sample is to be dealt with. Pour about half the sample through a nylon tea-strainer, this is ideal because it is designed to remove very small particulates and to drain quickly, then swirl the remainder of the sample around a few times so that none will remain stuck to the tumbler, and then strain the reminder. If the sample was cloudy or included lots of silty stuff it can now be rinsed very gently to remove these tiny particulates, then tap the rim of the strainer a few times to remove most of the remaining liquid, experience will show that it’s best to remove as much liquid as possible at this stage. Then tap the contents out onto the folded tissue, a solid tap should remove everything but anything still on the mesh will be obvious, and this is why a plastic strainer
is much better than a metal one. Now take the tissue and place it under a stereo microscope at X10, here it’s best to use an inexpensive model kept for fieldwork and samples as it will get filthy, it just will. It’s best to do this straight away rather than leaving the sample while you tidy up as live specimens are invariably present. Loose samples that include little debris can be examined directly and any specimens removed with a fine brush but BE CAREFUL, many tiny specimens (for me Euplectus are notorious for this) are easily overlooked, especially when they are clinging to debris, so look several times and be very thorough. Some samples will include lots of debris but there is an easy way of dealing with these; when these samples fall from the tumbler they land in a circular pattern which is preserved as the liquid drains into the tissue, using a pair of fine forceps work from one edge moving methodically through the sample to the opposite side, when finished work again from top to bottom and chances are, having worked through the entire sample twice, everything specimen will be found. Don’t take it for granted that every specimen will be found after one pass, it will not, and this will soon become obvious from the number of specimens found during the second pass. Use fine forceps to work the sample, brushes tend to get clogged really quickly and pieces of debris are not easily manipulated for examination using seekers. A fine brush should be used to remove specimens once they have been found because waterlogged specimens are often very delicate, especially when they are ‘bloated’. The best way to deal with specimens is to place them onto a small piece of damp tissue inserted into the plastic lid from a 3x1 inch glass tube, the lid can be placed on the tissue beside the sample and, with a little experience, specimens can be transferred without taking your eyes from the microscope, and because the tissue is damp they tend to transfer readily from the brush. The tube should be prepared beforehand, either with alcohol and a tiny drop of glycerine if the sample is to be stored, or with dry tissue if the if the specimens are to be set straight away, here a few drops of Ethyl acetate (Ethyl ethanoate) can be added just prior to when the lid containing the sample is placed onto the tube. If such specimens cannot be dealt with straight away the tubes can be kept in a fridge for a few days as moisture from the tissue on which the specimens were placed will keep them supple and ready to set or dissect for several days. For the more enthusiastic, the specimens can be set as soon as the sample is dealt with, and this is a good time to do so as the necessary equipment can be at hand for when the sample has been worked through and the specimens are still under the microscope. This is also a good time to dissect as specimens from extraction samples tend to be very relaxed and often a little bloated, and again the necessary equipment can be made available before the sample is worked.
What to expect
Beetles, of course, but be prepared for plenty of other stuff such as spiders, centipedes, millipedes, pseudoscorpions, springtails, hoverfly larvae, earthworms, slugs, snails, woodlice and fleas, among plenty of others. Small velvety cantharid larvae are a regular feature during the winter and, as they tend to remain alive in the collecting fluid, they can be released or reared. Many other larvae tend to survive immersion, even in dilute detergent, and probably the most frequent among these are those of hover flies and mosquitoes. Adults of most things tend to die off fairly quickly. In our experience the most rewarding samples are from decaying trees and marginal vegetation, but samples from any situation can include a diversity of beetles. Decaying trees are good at any time of year but during the warmer months many beetles are active at night and this will probably satisfy most coleopterists. Conversely many beetles spend much of their time among decayed heartwood and so this is always worth sampling, as is the debris that accumulates in rot-holes but use a trowel and slice down for a good sample. Scraping moss and detritus from logs can produce good samples, and, if a dust pan and brush happens to be handy, sweeping the detritus from denuded areas of wood on logs etc at night can produce very good results although such samples tend to fall through the supporting sieve of the extractor and so it will need to be supported on a thin layer of straw or grass clippings during the extraction process. Many species e.g. Plegaderus dissectus or Abraeus perpusillus will appear regularly and these are otherwise easily observed in the field, but extraction will also produce the other tiny histerids that are easily overlooked. The same applies to certain saproxylic laemophloeids, bothriderids and corylophids, and our tiny alexid might also be expected among wood samples. Most samples from wood will include at least some, and often a good many, ptilids, and when sorted from a sample under the microscope the different genera are easily picked-out (even if their identity is unknown.) Aleocharine staphs are also usually common from wood samples, especially where the sample includes mycelia or spores, and again there is usually a good diversity.
Otherwise-elusive groups such as Ptiliidae are often super-abundant in extraction samples
Small twigs covered with resupinate fungi may also be collected and extracted as they can produce good results, even in the winter, in fact no part of the tree should be ignored, and this includes the wood debris and decaying litter which tends to accumulate around the base of the trunk. Other groups such as ciids, erotylids, and tetratomids also occur among wood samples but these are more generally associated with arboreal fungi, which we hope to cover in another article. In the broadest sense members of numerous different families should regularly occur in samples from decaying trees. But a wide variety of tree species, both broadleaf and conifer, should be sampled to obtain the greatest diversity, and the occasional surprising species might occur e.g. in season we have found the malachid Sphinginus lobatus (Olivier, 1790) among debris under beech bark and during the winter numerous specimens of the leiodid Nemadus colonoides (Kraatz, 1851) among debris taken from deep fissures in the bark of an ornamental redwood tree, both in our local park. Things pick up in the spring and samples tend to become less diverse but there are many beetles that disperse at this time and so may appear among samples e.g. various species of Scirtidae, Clambidae and Curculionidae, especially Scolytinae and Cossoninae. There is so much to learn about sampling from trees but much of this comes down to experience, at first any detritus or moss might be taken and this will produce mixed results, but a bad result is as valuable as a good one because it will provide good experience, and after a while the samples get, on average, better, and with extensive experience it is easily possible to walk around a new park or area of woodland and take consistently good samples. A glance around will tell an experienced sampler what trees to head for and what material to sample, but unfortunately this requires a few seasons to get right. But it’s worth the wait. Of course the species of tree being sampled should always be noted, this is not always possible with decayed trunks or logs but experience helps out here as well as wood types tend to become familiar, and this and other data can be written onto each bag with a permanent marker so that there is no confusion between samples.
Many species move away from water to overwinter among marginal litter or under debris and so this is an excellent habitat from which to take samples. Decaying leaf- or reed-litter can be very productive indeed, especially as many species overwinter in groups, and so should be sampled whenever possible. Many of these can also be found by pitfall-trapping in the spring but this is a method we detest as it can be very destructive, especially where a rare species is active in numbers, and so this is our preferred method. In our experience there are three very productive marginal sampling sites; among reed litter, from partly-submerged vegetation, and grass and sedge tussocks. Samples of reed litter should be taken close to the water margin where they are damp but not water-logged, and the best sites are among dense reeds where the litter is deep. Scrape away the loose surface layers of litter and take samples from near the soil surface, the best way is to scrape surface litter and soil into a bag and seal it as soon as possible, and several samples taken from a small area tend to produce a range of species. This method is very good for small staphs, especially Carpelimus, Euplectus and small Stenus but a wide range of species is likely e.g. numbers of the tiny Aleocharine Deinopsis erosa (Stephens, 1832) often occur in this situation in late winter and early spring. Tussocks can be taken from any marginal situation and will often include the usual stuff such as overwintering chrysomelids and coccinellids, but even when taken from as much as 15m from the water they often include wetland species. Choose small tussocks with a densely compact base growing among damp litter, and take the whole thing; they can be cut close to the soil surface but better results usually result from uprooting the whole thing and including some soil and litter, thus various kateretids and nitidulids may appear in numbers. The best way to find hydrochids, hydrophilids and hydraenids etc. is to take samples of litter and vegetation from beside the water, these samples will be wet and we find that much of the water can be drained by using sample bags that have been perforated numerous times with a pin, the water will soon drain but the specimens, which tend to remain attached to the sample anyway, will remain in the bag. This method will invariably produce the usual common species e.g. Anacaena are almost always present but by using this method we discovered A. bipustulata (Marsham, 1802), which we had not found before, to be common throughout our area a few years ago, it remained so for two years and then vanished again. Other genera very likely to be sampled in this way include Laccobius, Enochrus, Helochares and Cercyon, and because they will be examined under the microscope any unusual specimens which might be overlooked in the field will be obvious. The usual idea of wetland margins are banks of ponds and rivers but other habitats such as peat bogs, marshes, tidal pools and estuaries should be sampled, and in the spring excellent results can be had from puddles and flooded tyre ruts in wooded areas.
Samples of fungus often provide various different species from numerous families
Many species move away from water to overwinter among marginal litter or under debris and so this is an excellent habitat from which to take samples. Decaying leaf- or reed-litter can be very productive indeed, especially as many species overwinter in groups, and so should be sampled whenever possible. Many of these can also be found by pitfall-trapping in the spring but this is a method we detest as it can be very destructive, especially where a rare species is active in numbers, and so this is our preferred method. In our experience there are three very productive marginal sampling sites; among reed litter, from partly-submerged vegetation, and grass and sedge tussocks. Samples of reed litter should be taken close to the water margin where they are damp but not water-logged, and the best sites are among dense reeds where the litter is deep. Scrape away the loose surface layers of litter and take samples from near the soil surface, the best way is to scrape surface litter and soil into a bag and seal it as soon as possible, and several samples taken from a small area tend to produce a range of species. This method is very good for small staphs, especially Carpelimus, Euplectus and small Stenus but a wide range of species is likely e.g. numbers of the tiny Aleocharine Deinopsis erosa (Stephens, 1832) often occur in this situation in late winter and early spring. Tussocks can be taken from any marginal situation and will often include the usual stuff such as overwintering chrysomelids and coccinellids, but even when taken from as much as 15m from the water they often include wetland species. Choose small tussocks with a densely compact base growing among damp litter, and take the whole thing; they can be cut close to the soil surface but better results usually result from uprooting the whole thing and including some soil and litter, thus various kateretids and nitidulids may appear in numbers. The best way to find hydrochids, hydrophilids and hydraenids etc. is to take samples of litter and vegetation from beside the water, these samples will be wet and we find that much of the water can be drained by using sample bags that have been perforated numerous times with a pin, the water will soon drain but the specimens, which tend to remain attached to the sample anyway, will remain in the bag. This method will invariably produce the usual common species e.g. Anacaena are almost always present but by using this method we discovered A. bipustulata (Marsham, 1802), which we had not found before, to be common throughout our area a few years ago, it remained so for two years and then vanished again. Other genera very likely to be sampled in this way include Laccobius, Enochrus, Helochares and Cercyon, and because they will be examined under the microscope any unusual specimens which might be overlooked in the field will be obvious. The usual idea of wetland margins are banks of ponds and rivers but other habitats such as peat bogs, marshes, tidal pools and estuaries should be sampled, and in the spring excellent results can be had from puddles and flooded tyre ruts in wooded areas.
Perhaps the best advantage of using sampling is that you will not need to re-visit a site to empty traps or spend time in the field sieving or working through samples, this is lazy but it is a great advantage when time is limited or when a promising site appears but cannot be worked, all that is needed is a few bags and a marker pen. The very local laemophloeid, Cryptolestes spartii (Curtis, 1834) occurs on dead twigs of broom where the bark is peeling, it is a tiny species that is very difficult to find; it can be beaten but it remains still on the sheet and is very difficult to spot. It took us hours to find this species on local wasteland, we beat many plants over a sheet and spent ages looking among the debris in hot sunshine, fortunately my son was able to see the stationary specimens among the debris but I’m sure we missed more than we found. At this time we were very busy with beetles in general and so had to be very selective about what we devoted time to, but with hindsight it would have been easier to beat a few bushes and take all the debris home for extraction, and when travelling to remote sites far from home this method would be excellent for finding the species without too much effort. While searching for Cryptolestes we also found the very local ptinid, Dryophilus anobioides Chevrolat, 1832, this remains our only record of this species but I think that beating Broom and sampling the detritus would provide more. Extraction sampling also allows a very thorough investigation of a local habitat e.g. taking samples from a domestic garden through the year will produce beetles that do not appear in pitfall or flight-interception traps. This can also become (in so far as biology can in general) quantitative if samples are taken on a weekly basis from particular habitats e.g. shrub-bed litter, decaying lawn clippings, moss from lawns or pathways and tussocks that have been allowed to develop for the purpose. Even a small garden will provide enough material through the year for this sort of surveying, and it is very satisfying to collect samples on Christmas day or at midnight on New-Year’s Eve e.g. a few years ago our first specimen of the year, found among a sample of debris taken from beneath a piece of wood placed on bare soil under leaf-litter in a domestic shrub bed, was the tiny blind bothriderid, Anommatus duodecimstriatus (Müller, P.W.J., 1821), an apparently very local species that is no doubt more common and widespread than records suggest. Thus an extractor is well worth the cost of a commercial model or the time involved in making one. There are other extraction methods that involve suspending samples in bags or keeping them in warm and dry containers, and while these can be just as productive, they take longer to process and usually involve more work, but they can process larger samples and they are for the most part passive and involve no wiring or electrical bits. Berlese extraction is quick and the results usually justify the effort, it is also exciting when good result have been obtained and more samples are waiting to be processed from the same batch