Collecting specimens

Sooner or later, every coleopterist will need to consider the moral issues surrounding ‘collecting’. ‘Collecting’ in the sense used by entomologists is almost always a euphemism for ‘killing’. Few specimens are found already dead and although one might think you could ‘collect’ a specimen, study it in life and then release it again, this is not common practice.

It is possible for a beginner to make serious progress in the study of beetles without collecting, but this is a difficult route which few have tried. The literature frequently assumes beginners will collect, and often fails to mention that it is ever possible to identify beetles without collecting them and keying them out. Those who do choose to collect specimens will doubtless wish to limit the number they take and to minimise any negative impacts they may have on populations.



Good reasons for ‘collecting’ beetles

  • To generate reliably-identified records – these can be fed into the national recording schemes, or to Local Records Centres and be used to the benefit of conservation. Records based on specimens can also be re-identified in future if additional species are recognised as British.
  • To develop your own reference collection – this can then be used to develop your identification skills to the point where you can identify more beetles in the field, and begin to study behaviour, habitat use and other aspects of their ecology and natural history.
  • To promote the study of beetles to others, who may have very limited experience of seeing beetles in the field.
  • To study them under a microscope, or to make a dissection.

Bad reasons for collecting beetles

  • To fill a collection the way a stamp collector fills a stamp album.
  • For profit from sale or exchange of specimens.

Collecting as a threat

The collection of specimens is a necessary requirement of many studies on beetles. Whilst the removal of adult specimens from a population is theoretically damaging, its impact is generally likely to be trivial for the species. Many species occur at moderate to high densities within their preferred habitats, and most techniques for finding or trapping beetles are very inefficient. However, in a few cases, detailed below, collecting can be an immediate and significant threat. It is certain that habitat loss and habitat degradation are ultimately very much more serious threats to beetle conservation. And ignorance of the species, their distribution and their habitat requirements is a serious obstacle to their conservation.

Under special circumstances, collecting can be a significant threat: when a species is easily found, occurs in small populations of long-lived adults, has low reproductive rate, occurs in easily damaged habitat, is targeted by beetle ‘twitchers’, or is targeted by greedy or commercial collectors.

The impacts of collecting can be mitigated to some extent by avoiding collecting gravid females (with conspicuously swollen abdomens) and by preferentially collecting males.

The future of collecting

It is interesting to speculate whether the study of beetles will evolve in the way that birdwatching and butterfly-watching have evolved. In those groups, the collecting of specimens has become increasingly outdated and unnecessary as field guides, field identification skills and optical equipment have all improved. This has also opened the way for a much greater number of people to get into those groups. Could certain families of beetles such as the longhorns (Cerambycidae), ladybirds (Coccinellidae) and ground beetles (Carabidae) go the same way?

How to kill a specimen

If you’re interested in this topic, you should definitely read the comments at the bottom of the page too with some excellent contributions from people with much more experience than me of methods 2, 3 and 5.

1. The best method: ethyl acetate.

Place your specimen in a glass tube, with a tight-fitting stopper. Put one or two drops of ethyl acetate (modern name: ethyl ethanoate) onto a piece of tissue paper, put that into the tube with the beetle and seal the lid. The beetle will be ‘put to sleep’ fairly quickly. Although there may be a period of rigor mortis, this will last no more than 4 – 6 hours, after which the specimen will be relaxed, allowing ‘carding’ (setting in position and gluing to a piece of card). Note that the tube must be glass – ethyl acetate dissolves many plastics! Appropriate care needs to be taken in handling this chemical: ethyl acetate liquid and vapour is flammable. Specimens in tubes with a bit of ethyl acetate will be preserved in a suitably relaxed state for several days, possibly weeks, depending on how good a seal the tube has. They will keep longer in a fridge or freezer.

A good tip from Andreas Herrmann which I’d not heard before is to add a drop of water (or spittle!) to the tube along with the ethyl acetate (1 drop water to 3-5 drops ethyl acetate gives best results). The water enables the hydrolysis of ethyl acetate, resulting in acetic acid and alcohol. Most likely the acetic acid protects the beetles from becoming stiff.

2. A useful back-up in the absence of ethyl acetate: boiling

Dropping specimens into boiling water is an effective and quick way to kill them within seconds, and then quickly transfer to cold water. This also helps to clean specimens. Haven’t tried this. It might even be a better method than ethyl acetate.

3. Alternative method: alcohol.

Submerge in 70% ethanol (alcohol) or IMS (Industrial Methylated Spirits). Beetles killed in this way are quite stiff and very difficult to card, seemingly because the alcohol dehydrates the specimen. They will however remain preserved almost indefinitely as long as the alcohol does not dry out. 70% alcohol is flammable, so care needs to be taken.

4. Alternative method: the freezer.

As with alcohol, this doubles as a killing method, and a method for long-term preservation. Specimens killed in this way tend to be quite stiff.

5. The traditional old method: the laurel bottle.

Young laurel leaves, collected in late May or early June, and mashed into the bottom of a big jar provide a cyanide killing bottle to last a whole year, apparently. Old leaves, and leaves which are not chopped finely enough, are much less effective. Laurel bottles apparently work extremely well, if you get it right.


8 Comments

  1. Mick Massie says:

    Killing: I have heard that dropping into boiling water for a second, then transferring quickly to cold water is a ‘popular’ method giving a quick death and a clean specimen and always available. Sounds plausible to me. Any comments ?

  2. markgtelfer says:

    Thanks Mick, Sounds good but haven’t tried it. Page updated.
    Mark

  3. Clive Turner says:

    A note on collecting:
    It makes sense not to kill more than you or others can sensibly utilise but it is important to note this should be the primary reason for limiting collecting.

    I have reams of data on insect mortality induced by natural factors and the activities of collectors pale into insignificance. It is clearly evident, and has been stated directly and indirectly by many leading authorities, that collecting cannot realistically impact on an insect population, and should it ever somehow manage to be a measurable potential threat then the species was certainly doomed in the first place, usually through human induced habitat destruction. The problem is that unlike birds, fungi, bacteria, climate and other reasons for insect death, collectors can be conversed with and easily seen in the act so are therefore the target of well intentioned but misplaced focus.

    The real issue is the science – to be absolutely sure of a record a specimen has to be available. For well-known taxa where the fauna is very well worked this has been substituted for records from contributors acknowledged as competent in that group. However this does not mean that vouchers should not be collected, in fact part of being a competent contributor to data sets also carries with it the responsibility to understand when a voucher is required and when not. For the record: the ultimate test of when a voucher is required would be whether the existing data would satisfy scrutiny by a specialist and on what level it matters (i.e. should a first vice county record be kept as a voucher – yes, should a tenth? – well, that would depend on the timeline of occurrence and is a matter of judgement). Of course, in the field all historical data is not available and so the default position has to be to initially retain vouchers of the potentially more interesting material.

    When visiting a site, particularly outside the UK, you may be the only entomologist to visit for some time and for many years to come. Our current knowledge of species and habitats often results from historical material collected with voracity. Our knowledge of species definitions changes with time and only with a good volume of historical material can judgements be made by researchers. This does not mean you should collect everything encountered but what it does indicate is that by not retaining adequate vouchers you would be doing a disservice to the environment you have visited and once you leave the site that snap-shot can never be recaptured. Similarly, contracted surveyors also have an obligation to properly work a site.

    In this rapidly changing world it is becoming increasingly more important to retain a solid record of what can be found where and when, this means a comprehensive voucher system. A simple species list is rarely sufficient when unravelling historical distributions of most species especially after they have been split. In such instances the previous data is void and the data on properly identified specimens substituted.

    Collecting activity is an essential component of entomology and although it may seem like carnage it is a matter of perception. The beetles in a tube may well be dead but I have few vials that could fill the stomach of a fish or bird let alone come close to the ravages of an unseasonal cold spell, eutrophication, pesticide application, bulldozer or viral outbreak.

    So, none of us want to but it is generally necessary to kill insects to identify them but respect them and the environment by preserving them correctly with accurate data, take only those you need and don’t let anyone make you feel guilty about it – there are not enough entomologists in the world to make a sufficient snapshot of this planet’s insect diversity before it disappears.

  4. Clive Turner says:

    Alcohol preservation:
    Preserve diest into 35%-45% alcohol and later that night or up to a few weeks later transfer to 70% and fix as normal. Don’t fill a vial more than 1/3 full of material.

    This method works well for producing relaxed beetles often with genitalia exuded easing dissection, they can also be mounted far more easily as they are generally not stiff. They are, of course, more delicate to handle but not brittle and due to their flexibility more resilient.

  5. Jason Green says:

    Hi Mark,

    Interesting article. I wouldn’t be surprised if butterfly-collecting ceased, in my opinion they are only about wing-colour; a simple net and field guide would suffice. Ladybirds? Well, 26 are easily-recognised, and I doubt would need collecting. The others such as Scymnus would be needed, for personal familiarity once keyed. Cerambycids? Probably the same.

    I tend to ‘sample’ sites; one or two little blue Apionid weevils from one site, one or two from another – then identify. Further ones a few months later I bring home, knock-out and check then release if they’re the same and keep if different, but only two.

    Good post/comment, Clive.

  6. Jon Cooter says:

    Mark:

    I thought I’d submit a note about the “laurel bottle” having used this method on and off over 20 – 30 years. I’m sure there are much better modern methods available, but laurel costs nothing and in my experience keeps beetles ‘relaxed’ for as long as needed. And in an emergency it is nearly always possible to make one up (I’ve even used a half leaf and a 35mm film canister having captured a desirable beetle when out on non-ento matters).

    Beetles are first killed with ethyl acetate vapour then transferred to the laurel bottle.

    As you point out pick the young bright green leaves during spring. I cut them with scissors into an old fashioned tall sweet jar with screw cap. When there’s about 5cm of clippings in the bottom I ram the lot down with the proverbial blunt instrument – a hammer handle is good. I then place about 2cm of cellulose wadding on top of the cut, crushed leaves.

    For years I have had two boxes of cellulose extraction thimbles as used in the chemistry lab (?Soxhlet extractor). The cellulose absorbs moisture keeping the beetles within more or less dry, at least certainly not in contact with droplets of moisture. I put a few beetles in, then some tissue, beetles, tissue, building up a ‘multi-storey’ tube of beetles. Data goes in first and again last and the whole is plugged with more tissue.

    I’ve also simply put the beetles into screws of tissue (sort of like boiled sweets) with their data, especially if away from home on a trip.

    If the jar is lined with blotting paper (is this still made I wonder?) or similar absorbent paper, so much the better. Moisture is a problem with laurel (don’t put the bottle in sun light or near a radiator for example).

    Some colours are affected by the laurel vapour, but in my experience only if kept in the jar for a few months. Small staphs etc seem OK and can easily be dissected, though I’d not recommend leaving small-fry in laurel atmosphere for more than a few months. The beetle should NEVER come into direct contact with laurel leaves. (That said, as a teenager in the early 1960’s I was often sent beetles from Zululand, RSA – the contents being a mix of beetles and cut/crushed laurel leaves all tipped into a plastic bag, put in a cardboard box, addressed and mailed. I dealt with these upon arrival and the material is extant in pristine condition).

    It seems never to be mentioned but another method I find VERY useful is simply to kill the material and leave it to dry, then carefully pack it between layers of tissue. It can then be quite simply ‘relaxed’ by the usual method (mine is to put water-saturated tissue in a small (say postcard-sized) shallow (say no more than 2cm – 3cm) plastic box and leave it overnight in the airing cupboard or in summer on a sunny window sill.

    I used this last method during a trip to China when the local ‘industrial alcohol’ was particularly useless for preservation (but worked well in the pit-fall trap cocktail mix – Coptolabrus liked it anyway!).

    NEVER USE COTTON WOOL.

    Jon

  7. Simon Horsnall says:

    Boiling: I have found hot water to be a most difficult method to get right. I think it is along the lines of a boiled egg. Whilst the specimen is fairly plastic, there tends to be a level of elasticity which I think may be caused by “cooking” the muscles. I’ve had better results with very hot water (hottest the tap can supply, about 60C) but would only use it in an extreme emergency.

  8. markgtelfer says:

    Thanks Simon, good to have your experience – it’s not an emergency I’ve ever faced.

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