Collecting insects
This is a little article I wrote to help novice entomologists understand the basic principles behind making an insect collection.
Entomologists frequently have to catch and kill a few insects to identify them because it isn’t possible to key some orders 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.
Once a specimen has been identified it is almost always kept for future reference in a personal reference collection. This is usually so that the record can be backed up by physical evidence that can be examined later by other workers and it can also be useful to have “one I did earlier” that you can compare to any newly identified specimens.
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 short & 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 (eg. spiders / molluscs) and others assume they will be dry (eg. 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!!
Mounting specimens
I pin most specimens on their sides (lateral pinning) with very fine micro pins; slightly diagonally so that I don’t destroy the same feature on both sides.
I then make a ’stage’ using a thin strip of plastazote (high density foam) and push a thick (size 3 or above) ‘continental-sized’ entomological pin (the ’stage-pin’) 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 fingers are likely to interfere with it. (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), ‘top-down’ (dorsally) but I only do this with very large specimens (eg. Tachina spp. or bigger) where you can use a relatively thick pin (such as a #1 or #2 thickness).
Another popular technique (especially with hymenopterists) is gluing the insect to strong card but I feel that very fine micro pins are:
- much easier to use (glue is very messy)
- allow for moving later (glue can be dissolved but it isn’t easy)
- need not destroy much/any surface features
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 put the specimen’s labels (copy of the data & a separate label for the name) on the big stage-pin, under the stage. 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.
Equipment
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
- very fine waterproof ink pen for writing labels (0.1 thickness)
- box of continental-length pins & pre-made stages
- box of spare foam 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 (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.
Pins
I use a variety of different pins for different jobs:
- “Continental” length (38mm) pins for use as stage-pins (in size 3 or 4 – approximately .55mm diameter) or sometimes for direct-pinning very large specimens (in size 1 or 2). I would never use a thinner pin because thin pins flex too much and, although they push into foam relatively easily, you will have to push them into cork at some time and this is much harder! If the specimen is small enough to need a thinner pin then you should be staging it.
- Micro pins for direct-pinning most of my specimens:
- D3 size (0.01″ x 10mm) for larger specimens
- A1 size (0.0056″ x 10mm) for anything small
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.
Mine is a MEIJI zoom binocular microscope with a zoom range of 0.7x – 4.5x and 10x eyepieces giving me 7x – 45x total magnification. The light 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 imrtant thing – no eye-strain after working with it for hours on end!
I do occasionally struggle at the top end when trying to identify something really tiny, like a pteromalid wasp, but I just know my limitations and avoid these as much as I can. In most of these cases (when you get to magnifications over 50x) it would probably be advisable to prepare slide mounts and use a stage microscope instead. That’s a completely different world though and I have no experience of that.
Accessories
You might also like to think about buying a spare 10x eyepiece fitted with a graticule. A graticule projects a grid or measuring line onto the image and allows you to measure relative distances accurately. This is very important when trying to work out the relative width of the frons in tachinids or the ovipositor/tibia ratio in ichneumonid wasps.
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 bulb/tube. 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 either rely on being able to examine the male genitalia or you can use the genitalia as a good confirmatory character. For this reason I always recommend that if you have a male dipteran you should always try to hook out the genital capsule when you first pin the specimen. You might not need to do it but it’s much easier 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 flexible.
For some groups (eg. 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.
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-soluable, 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.
Labels
All specimens should have at least 2 labels:
- the data label (under the stage) containing:
- Place of capture (region, place name & map reference)
- Date of capture (often with the month in roman numerals)
- Name of collector
- (optional) method of collection
- the determination (or ‘det’) label (usually the lowest label) containing:
- Species name
- Sex
- 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 indellible inks. At the begining 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 (potential 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!
Storage
As I mentioned earlier, I use the flat, clear, plastic boxes for unidentified specimens and they keep everything clean and dry and take up very little space. But how you arrange your identified specimens (your reference collection) is largely down to personal preference and budget.
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 large, purpose-built, wooden store boxes (see right). Second-hand boxes can be bought from most natural history museums and you can just line the bottom with a 6mm or 9mm sheet of Alveolit foam. Alternatively you can buy new boxes from entomological dealers.
As I mentioned earlier, pests are a worry for all collectors – if museum/carpet beetle (Anthinus 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 black-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 after 2-4 weeks you should allow the bag+box to reach room temperature before opening the bag or box and letting air in – otherwise condensation will form on your specimens!
Drawers & unit-trays
Once your collection spans 5 or more store boxes it starts to become very unweildy 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.
Pros:
- 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
Cons:
- takes up more space
- more difficult to carry around and take to workshops etc.
- it is much more expensive
Posting 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) later 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. But this might cause some raised eyebrows if you send them internationally because customs never like to see alcohol being brought into the country, whether it has flies in or not!
Conclusion
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 seen 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! 