This is a little article I wrote to help novice entomologists understand the basic principles behind making an insect collection.
Why do we collect insects?
It’s a fair question – especially when there are such amazing cameras these days and forums all over the web where experts will seemingly put a name to just about anything. So here is my round up of why taking specimens is still essential to help us understand the biodiversity around us:
- To make a positive identification: Photographs of living insects very often do not show the parts that confirm the species, like the genitalia or minute bristles and surface sculpturing. It is possible to identify some insects from photos but even a lot of common insects might only be identifiable to genus or the expert might say “probably X” or “possibly Y” – so they have no way to be really sure.
- It would be impossible to target conservation work or finances to any project where workers weren’t actually sure that the target species was present. Many rare species look almost identical to common species so being able to make a confident identification is still very important to some people.
- To confirm an identification after the fact: Having specimens also allows us to re-examine the work of our peers to make sure that they were correct OR to reindentify something in light of new knowledge. Science isn’t static and we are constantly learning how better to understand the world around us so we often discover cryptic (very similar & closely related) species hidden in what we thought was one species. Without specimens we wouldn’t be able to retrospectively reassess old observations and we wouldn’t be able to do the detailed DNA or morphological comparisons to find them.
- It was once thought that a colony of Linnaemya rossica had existed in Kent since it was first discovered in the 1940s, however when the genitalia of a modern specimen were examined we discovered that it was a different species, Linnaemya picta. The 1940s specimens exist in the NHM so we examined them and were able to confirm that they too were picta and so the Kent colony had always been picta. We did a similar analysis with records of Bithia modesta, which we now know is actually Bithia demotica.
- To allow analysis in the future: Having specimens allows people in the future to use analytical methods that were inconceivable to the collector. The classic example is all the DNA work that is being done on specimens collected before the discovery of DNA. This research is helping us to clarify species boundaries and also to analyse genetic diversity.
- The NHM have been analysing their old specimens in novel ways – such as rehydrating pressed flowers to examine the pollination success in past centuries. This feeds data into the discussions around pollinator decline.
- We need more good data: Having large collections allows researchers to use larger datasets. One record is interesting but when you combine it with hundreds of others then you can start to plot changes in distribution, phenology and in biodiversity.
- We are eternally grateful to the Victorian and Edwardian collectors who left us large samples of specimens that our researchers are now using to feed more data into climate change and biodiversity research. http://www.nhm.ac.uk/our-science/our-work/digital-museum/digital-collections-programme/digitising-butterfly-moth-collections.html
But do we need to know exactly what something is? Why can’t we leave wildlife to get on with it? Well, investigating the world around us is how science has brought civilisation to the level that we have now and the basis of all sciences is to be able to identify what you are looking at. Imagine how much the science of chemistry would have advanced if chemists couldn’t tell gold from lead!
If we can identify species and understand how their populations are being changed by human interaction (e.g. global warming) then we have the best chance to help them in the long term. We can monitor their populations and distributions and better target conservation efforts to the exact causes of decline. Certainly, we can be certain that more ignorance definitely won’t turn back the tide of climate change or habitat destruction. What we need is to provide the evidence to halt damage.
Can’t we just just leave it to the professionals? Surely there’s no reason for amateurs to collect insects any more? This argument stems from a few basic misunderstandings, that “amateur” somehow means “not very good” and that professional entomologists do most of the important biological recording and taxonomy work. Actually in the UK and much of Europe we have a long and distinguished history of amateur entomology leading the work on taxonomy and providing the vast bulk of data on which important decisions are made.
Funding for professional entomologists has been slashed time and time again and the remaining few have to focus on commercial work or very specific tasks. We actually need a lot more specialist entomologists to be able to survey for rare species and priority habitats and it’s clear that this will have to come from the amateur community. Citizen science is the new buzz-phrase and many experts are spending a lot of time and energy training people to identify difficult groups, such as bees, wasps & flies, which together form the majority of our declining pollinators.
Can insect populations survive collection? What if we kill the last one? Collectors these days very rarely take more than a few individuals of each species because they are just sampling a particular area or taking reference material. The days of the Victorian-style collector, taking long series of rare species to fill gaps in his collection, are long gone.
The taking of a few specimens in any location is not going to make any difference to a population of invertebrates because they exist in such large numbers. Insects reach maturity quickly and lay lots of eggs and so their capacity to recover even large population disasters (like a fire or flood) is far greater than with vertebrates, which breed far slower and exist in smaller populations. Predators like birds and other insects will eat many hundreds (even thousands) of insects in a day and this is all considered normal so the effects of a few entomologists really doesn’t amount to anything.
Stuart Ball & Roger Morris once did a highly scientific “mark & recapture” experiment to ascertain population density and movements within a particular habitat. Their results showed that no matter how hard they caught their flies (hundreds were captured) and marked them, the fact that they re-caught so few marked ones showed that they were only affecting a mere 16% of the population. This shows how tiny would be the effect of removing a few individual specimens.
Isn’t killing things immoral? This is a difficult and highly subjective question to which you can find many answers. I have never met an entomologists who enjoys the act of killing but we are scientists who understand that the effect of our science is likely to be far more beneficial in the long term because the knowledge gained will go to help the species we are collecting and the habitats they live in. When entomologists euthanize specimens they do so in the most humane ways and always prior to pinning so that there is no actual cruelty involved.
Furthermore, if one is so worried about the avoidable death of invertebrates then consider the effect that we all have on the world around us. We kill hundreds of insects in our cars and I need not even mention the use of insecticides in the manufacture of the food we all eat. Indirectly we also contribute to the death of millions with global warming, intensive agriculture and mining. There are few ways that we don’t affect invertebrates but if we are careful and we try to mitigate this wholesale destruction then insects will thrive.
Basic reasons that you might want to collect
Entomologists frequently have to catch and kill a few insects to identify them because it isn’t possible to key some taxa without being able to examine them in minute detail under a microscope. Accurate taxonomy & identifications / records are a vital cornerstone of entomological, environmental & conservation sciences because they provide vital distribution and biodiversity information. Taking regular specimens over time will show you variations in the population and without the work of the old collectors we wouldn’t have been able to detect the changes that had happened to species like the Peppered Moth.
Once a specimen has been identified it is almost always kept for future reference in a personal reference collection – they are not thrown away. This is so that the record can be backed up by physical evidence that can be examined later by other workers and it can also be very useful to have reference material that you can compare to any newly identified specimens. This is especially true when you are starting to learn a new group and are not familiar with the range of the fauna. Having specimens also allows us to revisit the work that we did years previously when we were young and inexperienced. We all make mistakes and being able to show the specimens to someone else (or your better self) allows corrections to be made.
Things to think about
When collecting for science (as opposed to ‘pretty displays to hang on the wall’) there are only usually 2 things to think about:
- How can I protect the specimen in the long term?
- How can I present the specimen so that it can be examined and identified as easily as possible?
With each group of insects you will find different answers to these questions, depending on how hard/soft the specimen is and which features are used in the identification keys. For instance, some keys assume that specimens are preserved in alcohol (e.g.. spiders / molluscs) and others assume they will be dry (e.g.. most insects).
In the following article I will assume that we are talking about Diptera & Hymenoptera (my specialities). Also, please remember that a lot of this is my personal preference – other people might/will disagree!!
You can start with dead insects you find at home or in the countryside but they will usually be too dry & brittle to pin immediately. Place them in a plastic box on wet tissue paper for 24 hours to make the soft enough.
If the specimens come from wet traps – traps that collect into liquid (e.g. malaise or pitfall traps) then the insects will usually to be supple enough to pin immediately.
If you collect live insects then the easiest and most humane way to bump them off is to put them in a domestic freezer (-18c and below) for a few hours. Then when you take them out let them defrost for an hour before pinning.
I side-pin and stage-mount most small to medium-sized specimens because it has many clear benefits for Diptera & even some Hymenoptera. The micro-pin goes into the side of the fly’s thorax at a slight angle so that it doesn’t damage the same feature on both sides – all manipulated by fine-pointed forceps. The fly is then pinned to a block of foam and the legs, head & wings are all arranged to show off the essential features. If the specimen is male you can very easily hook-out the genitalia and use more micro pins to hold the bits out while the specimen dries – see below.
When the specimen is dry I make a ‘stage’ using a thin strip of NuPoly/plastazote (high density foam) and push a 38mm, continental-sized, entomological pin (size 3 or above) through one end so that the stage sticks out at 90-degrees. The micro pin with the specimen is then pushed firmly into the foam stage. The stage is positioned half way up the stage-pin, to allow room underneath for labels, but not too close to the top of the pin where chubby fingers are likely to brush against the specimen. (see right).
Staging combined with side-pinning protects small/medium specimens very well while displaying as many features as possible. The stage-pin is strong and easy to hold/manipulate when moving the specimen and the stage absorbs most vibrations.
Alternatively you can direct-pin (see right), either laterally or dorsally (‘top-down’) but I only do this with very large specimens (e.g.. Tachina spp. or bigger) where you can use a relatively thick pin (such as a #1 or #2 thickness). Never use the finest thickness (#0 and #00) 38mm pins because they are almost impossible to push into older cork boxes and also bend and twist when you push them into foam, which can cause the specimen to crack or twang and loose legs and antennae.
Another popular technique (especially with hymenopterists) is gluing the insect to strong card (carding or pointing) but I feel that very fine micro pins are:
- much easier to use – glue is very messy
- hold the specimen in place more reliably – if the glue isn’t perfectly tacky then insects can drop off later
- allow for moving later – glue can be dissolved but it isn’t easy and bear in mind that you might not want a white background for photography
- need not destroy many/any surface features – glue can cover up more than a single pin hole
Also, I like to have a single mounting method for most of my specimens and glue doesn’t work well for groups like Diptera, which have fragile bristles.
Knock-on benefits of staging
Coincidentally, micro-pinning lends itself very well to bulk-collecting of groups that take a while to identify.
I usually go out for a day in the field and catch maybe 50 insects of various orders and families. I also work with Malaise traps, which can generate vast numbers of specimens on a single day of sorting. This is just too many to work with in a single day so I reduce the work I have to do by creating batches of insects.
I simply pin all the specimens with micro pins and put them in a flat, clear, plastic box with a sheet of foam at the bottom (see right) with a data label containing all the collection data for that group of insects. The specimens stay here to dry and allow me to continue working on other batches in the meantime.
When I have time to identify them I pin a specimen to a new stage, identify it and then put the specimen’s labels (copy of the data & a separate label for the name) on to the big stage-pin. The specimen is then ready to be transferred to the collection-proper. The other specimens may stay in the plastic box for years waiting for me to have time to work on them but while they are in this state they take up very little room and they are very well protected from dust and damage.
My standard kit (see right):
- L-shaped foam block for holding pinned specimens under the microscope
- Drilled aluminium pinning block for positioning stages & labels at uniform heights on the stage pin. If you standardise the height of your specimens & labels it makes it much easier to compare dozens of specimens in trays
- Very fine waterproof ink pen for writing labels (0.1 or 0.05 thickness). The photo is a bit old and I actually use black Pigma Micron 005 or Zig Millennium 005 pens these days because they use archival quality inks. You can buy them quite cheaply from sites like Cult Pens [EDIT: I’m currently also trialling a Copic Multiliner SP 0.03, which has a slightly finer point]
- Box of continental-length, 38mm pins & pre-made stages
- Box of spare foam NuPoly strips for stages and some glass tubes containing micro pins and more continental pins.
- Small pair of scissors, for cutting up labels
- Curved and straight pointed forceps for manipulating specimens and micro pins. I don’t like the grooved ‘entomological’ forceps because they ‘twang’ pins too often.
This all fits neatly into a plastic Tupperware box for storage and transportation.
The best forceps are made by Swiss companies like Dumont and Ideal Tek and it is always worth buying good quality forceps because they will be finer with harder steels and the points will meet perfectly. I tend to use types:
- #3c (watch-makers, pointed): These are my standard forceps – they are not as fine as the #5 or #7 but the extra width behind the point makes them much less likely to cross/twang when holding a micro-pin.
- #4 or #5 (biological, straight, fine pointed): Very fine for holding specimens or tiny micro-pins.
- #7 (biological, curved, fine pointed): As above but the curve actually makes the points less likely to cross/twang and, when combined with a #5, make a very good combination for micro-pinning specimens or manipulating genitalia.
The basic tweezer steel is Inox or stainless steel – it is quite magnetic and can tarnish a bit if you get acid on it but it is a good all-round steel. The next up would be the anti-magnetic and anti-acid steels like Dumoxel or Dumostar. You don’t really need these but Dumoxel is quite nice because it can be a pain if your pins become magnetised and stick to the forceps.
You can sometimes pick up bargains on eBay and my favourite forceps are a pair of Dumont 3C forceps in Dumoxel steel, which cost me all of £3.50 – these would normally retail for about £30. Expect to pay between £20 and £40 depending on whether you purchase from an online supplier or specialist shop.
Always remember to replace the plastic point protectors when you are not using forceps because they have a knack of being dropped/dragged onto the floor and will usually land points down – wrecking the alignment and bending the tips. You can resharpen forceps to try to fix bent tips but they will never be as good.
I use a variety of different pins for different jobs:
- Continental length (38mm), stainless steel, nylon-headed pins for use as stage-pins (in size 3 – 5 – approximately .55mm diameter) or sometimes for direct-pinning very large specimens (in size 1 or 2). If the specimen is small enough to need a thinner pin then you should be staging it. If you use size 0 or below then the pins will twang and you’ll damage specimens. You can get away with cheaper “Asta” steel-headed pins for staging with foam but I would use a higher quality pin for direct pinning specimens and for piercing card mounts.
- Micro pins for direct-pinning most of my specimens:
- A1 size (0.0056″ x 10mm) for anything tiny
- B2 size for medium specimens
- D3 size (0.01″ x 10mm) for larger/standard specimens
Microscope & light source
A good binocular microscope with lighting system is an essential part of any entomologist’s equipment and is probably the most expensive thing you will buy. Ideally you want to invest in something that has very good optics and a good, bright white light source. You don’t need to have very strong magnification – about 5x-40x is perfectly adequate for most flies & wasps. When you are ready to buy a microscope the main thing is to try out plenty of different models and test them with a few of your own specimens.
I currently use 2 setups:
- Leica S8APO with Chinese 144-LED ring-light – a lovely microscope but a bit expensive for most beginners to buy (~£4000 new from a Leica dealer). Fitted with 10x wide-field eyepieces the zoom range is 10x to 80x and the camera port on the top coupled with the fine-focus rack-stand allows you to take stacked photos of whatever you are looking at. The optical quality is absolutely superb.
- My first microscope was a MEIJI EMZ zoom binocular microscope with a zoom range of 0.7x – 4.5x and 10x eyepieces giving me 7x – 45x total magnification. The light for this scope is a small Mini-Fluor fluorescent light attached to the microscope itself. I find it very good and easy to work with and the most important thing – no eye-strain after working with it for hours on end!
The 144-LED ring-lights can be used on virtually any microscope and can be bought on Amazon or eBay for around £60 – just shop around.
I like to have one of the eyepieces fitted with a graticule – a grid or ruled line that helps you measure relative lengths of objects on the specimen. They fit inside one of the eyepieces and are fiddly to put in so the standard technique is to either have it in a spare eyepiece or just leave it in one of your main eyepieces all the time.
Secondary lighting can also be very useful when peering into dark corners of a small specimen. An easy and cheap solution to this is to get a small, household angle-poise desk lamp fitted with an energy-saving fluorescent or LED bulb. These can be pointed to shine directly at the specimen from any angle and the tube is never usually hot so you don’t risk burning the back of your hand – or melting the specimen’s wings!!
Working with genitalia
Some identifications rely on being able to examine the male genitalia and you can usually use the genitalia as a good confirmatory character. For this reason I always recommend that if you have a male you should always try to hook out the genital capsule and aedeagus when the specimen is fresh. You never know when they will be important but it’s much easier to do while they are soft and pliant, than try to relax an old specimen – a little bit of work early-on will save you a great deal of hassle later.
All you need to do is to use a fine micro-pin and gently tease open the genital capsule. You should find that it hinges on its dorsum and you can hold it open with a few crossed micro-pins until the specimen has dried and is set (see right). Sarcophagids are an extreme example because they have such huge genital capsules but they are also a good example because the work is incredibly hard to do unless you open it while the specimen is still very flexible.
For some groups (e.g. sarcophagids) it helps if you can also locate and tease out the aedeagus, a small chitinous filament that’s analagous the penis in higher animals. This is pretty small and of course it is the most internal & anterior part of the external genitalia so it is difficult to get to – but it isn’t impossible and it makes the genitalia preparation complete. It usually pops out if you extend the cerci fully – about 90-degrees to the body.
Sometimes you will have to remove the genitalia completely and in these cases they can be clarified in a weak solution of potasium hydroxide (KOH) but it isn’t usually necessary. Be very careful with this chemical – it is strongly caustic and care should be taken to keep it away from the skin and eyes!
I normally store separated genitalia in little plastic capsules that can be pinned beneath the specimen, just above the data label (see right). The genitalia are suspended in a solution of glycerine. This system protects them but also allows them to be examined later – though it can be tricky to see much through plastic and glycerine so you have be prepared to remove them.
Alternatively, some people choose to store the genitalia in a water-soluble, transparent resin DMHF. When dry DMHF becomes a hard, long-lasting and transparent droplet that both protects and holds the genitalia, yet allows it to be examined and can be dissolved later if necessary.
All specimens should have at least 2 labels:
- the data label (first under the stage) containing:
- Place of capture (region, place name & map reference)
- Date of capture (often with the month in roman numerals to prevent confusion between American and European date formats)
- Name of collector
- (optional) method of collection
- the determination (or ‘det’) label (usually the lowest label) containing:
- Species name
- Name of determiner
- Year of determination
If the specimen was reared from a host or collected as a larvae you might also want to give it a rearing label (under the data label) containing:
- host name
- date of capture
- any other notes relating to the rearing
These labels are made from card or thick 160gsm paper so that they don’t drop or rotate on the pin. Never use thinner paper – it won’t grip the pin properly and it will start to spin round and need replacing very quickly.
I normally print as much of the static information as I can, using laser or inkjet printers fitted with indelible inks. At the beginning of each year I print:
- A sheet of part-completed data labels for each of my favourite collecting sites containing all the information except the day & month of collection.
- Sheets of completed det labels for the groups I study most frequently. I collect a lot of tachinids and I am also asked to det tachinids throughout the year so it makes sense for me to carry around sheets of pre-printed det labels to save me having it write them each time. Obviously, I print more labels for the common species and less of the rarities.
- A sheet of ‘blank’ det labels (with just my name and the current year) which I will use for all non-tachinids.
Resist the urge to write additional information on the back of the labels – nobody will see it or check it and the specimen will have to be picked up to read it (increasing the potential for handling damage). If you have additional information then print a bigger label and print it on thicker card – the larger labels will actually protect your specimen better!
These days specimen data should always be digitised and kept on computer so that the records can be distributed to recording schemes and other taxonomists. I keep my specimen collection data in a spreadsheet which I store and maintain on Google Docs, for ease of access. My standard layout is:
- binomial name
- country [CAPS]
- map reference [in any format]
- voucher [only used if there is a voucher code associated with the specimen]
- dataset [used to group large donations]
- collection [used to mark which collection it can be found in – e.g. CMTRPAL, GIVEAWAYS, or the name of the person I gave the specimen to]
With this kind of database you can generate mail-merged data labels and determination labels fairly easily. You can also quickly format it up to be sent to recording schemes or to iRecord and the NBN.
As long as insect specimens are kept dry and pest free they will last for centuries so the storage requirements can be very simple indeed. You could just start with sturdy, air-tight Tupperware/food storage boxes and glue some 6mm foam onto the bottom to hold your pins. You could arrange them so that each box holds 1 genus or just mix specimens together – whatever you can cope with without loosing specimens in a muddle!
Most amateurs use purpose-built, wooden store boxes (see right). Second-hand boxes can be bought from most natural history museums (from around £10 – contact Max Barclay at the Natural History Museum in London) and you can just line the bottom with a 6mm or 9mm sheet of plastazote foam. You can also buy new boxes from entomological dealers but they can be a little expensive.
You can also use snap-lock Tupperware food storage boxes with 9mm foam glued to the base of the box or inside the lid. This is a very cost-effective system and has the benefits that they are air-tight (so very pest-resistant) and you can see the specimens without opening the box. However, be very careful not to store damp specimens in these boxes because they will soon be covered in mould if you are not careful. Many people pin sachets of silica-gel into these boxes to prevent humidity building up.
I used store boxes for many years but as my reference collection started to reach 300 species with 2000 specimens it became too difficult to manage. Inserting new species often meant that I had to move lots of specimens around and that brought damage and took a long time. Also, store boxes are not very air-tight and so pests can get in and if not noticed for a while can cause immense damage.
Drawers & unit-trays
Once your collection spans 5 or more store boxes it starts to become very unwieldy to curate and refer to. Each time you insert a new species you have to shuffle the rest up or down in the box to make space and each time you do this you have to handle each specimen, risking more accidental damage.
Most museums use a system of air-tight, steel cabinets and wooden, glass-topped drawers. Into these drawers you can drop small, standard-sized, card boxes lined with foam called unit-trays. Each tray is sized to be a multiple of the smallest unit so that they can be swapped around and organized into neat rows. Each unit-tray then contains 1 species.
You can easily move the boxes around and slot in larger or smaller units as your collection grows without actually handling any pins. Also you reduce the amount of handling each specimen gets in its lifetime because routine checking of a species can be done by just picking out the whole unit tray and holding it under the microscope.
- it’s easier to quickly see into the glass-topped drawer
- reduced individual specimen handling
- much easier to add new species
- much easier to rearrange your collection
- broken body-parts tend to stay in the appropriate tray
- the unit-trays give added protection while the specimens are out of the drawer
- takes up more space
- more difficult to carry around and take to workshops etc.
- it is much more expensive
As I mentioned earlier, pests are a worry for all collectors – if museum/carpet beetle (Anthrinus sp.) get into a collection they can reduce your prized specimens to dust in a matter of weeks. But if you are worried about pests you can always freeze your specimens in a domestic freezer (2-4 weeks at -20C will do the trick). Museums do this as a matter of course because many insecticide chemicals (naphthalene etc.) are now frowned-upon or banned. Just place your box in a plastic bag and expel as much air as possible before sealing it up so that air and moisture can’t get in during the freezing process. When you remove them you should allow the bag+box to reach room temperature before opening it and letting air in – otherwise condensation will form on your specimens!
Sometimes it might be necessary to send pinned specimens to another expert for identification. This need not be a problem as long as you have good, sturdy boxes with a deep (9mm) layer of dense foam attached firmly to the bottom. Into this pin your specimens and if you think the stages might spin you can hold them in place with more 38mm stage-pins.
It is also possible to send specimens from Malaise trapping in glass tubes of alcohol protected by plenty of cotton wool and a sturdy cardboard box. Just remember to use a plug of tissue to push the flies to the end of the bottle/tube so that they cannot slosh around in transit. Bubbles and simply the movement of the specimens rubbing against each other can cause a surprising amount of damage.
If you are sending them internationally customs will generally not be interested as long as you label them clearly – “Dead entomological specimens for scientific study, no commercial value”. If in any doubt always check with experts in the destination country, in case they have any special requirements or prohibitions.
I don’t advocate any one supplier over another and I don’t get any commission for recommending people on this list. But here are the suppliers that I use and have found reliable:
- Watkins & Doncaster (http://www.watdon.co.uk/): The most famous entomological supplier in the UK – very reliable with a good range of pins and other equipment (nets & malaise traps etc). They are also the only supplier I know for the NuPoly staging strips.
- David Henshaw (http://www.ebay.co.uk/usr/djanddhenshaw): A good range of cheap pins, chemicals and useful things like genitalia storage tubing and flat plastic boxes for pinning specimens into. His eBay shop currently only stocks the pins so contact him directly for a price list that covers his full range.
- B&S Entomological Services (http://www.entomology.org.uk/): A good range of nets and malaise trap parts.
- EntoSphinx (http://www.entosphinx.cz/en/): A good range of boxes, pins and equipment.
- Paradox (http://www.paradox.co.pl/): A good range of boxes, pins and equipment.
Whatever you do, start by thinking hard about the groups you are studying and adapt what I have said for your own circumstances. There are no hard and fast rules other than the 2 points I made in the opening paragraphs – protect your specimen and present it so that it can be identified easily. Also, remember that you need to have a system that suits your circumstances and your lifestyle – if it is too much hassle you won’t enjoy your hobby and you won’t have the time to do the really enjoyable and exciting parts of the work. Lastly, if you have any comments you’d like to make or any suggestions for improvement then feel free to leave a comment somewhere and get in touch! 🙂