Beetle Collecting 101: How to pin a beetle

It has been a long time since I initiated my Beetle Collecting 101 series (more than two years!), and to date the first issue—Beetle Collecting 101: Dress for Success—remains the one and only lesson that I’ve posted. I really had intended to follow that up with nittier-grittier posts on the actual mechanics of collecting beetles and processing the specimens for long-term preservation, but I didn’t and don’t know why other than to say, well… life happens. It’s never too late to fix something, however, so as a long overdue follow up I thought I would give a short video lesson on how to pin a beetle—specifically a cerambycid (longhorned) beetle. Featured in this short (4:31) video is the lovely Megacyllene decora (amorpha borer), which I found back in early September at a site in Missouri’s southeastern lowlands. Click the image to be directed to the video.

Copyright © Ted C. MacRae 2012

16 thoughts on “Beetle Collecting 101: How to pin a beetle

  1. Neat video. It reminds me that I still need to follow up on my Tiger Beetle Mojo: Comments on Curation post.

    Sometimes when using that coarse opened-celled foam the tarsal claw(s) can stick when being removed and possibly break off. I attach a sheet of bond/copy paper over the foam surface before pinning.

    Just my .02 ¢

    • I’ve tried that but don’t like the way the paper moves around – it just bothers me… When I’m ready to remove the specimen I just pull it up slightly and then use the forceps to free any hanging claws.

  2. Cool, Ted. I’ve never done that, but it’s interesting to see the process. So after what’s shown in the video, how long do you have to leave it in on the Styrofoam? And when you go to remove it, do you sometimes have breakage because things get stuck to the Styrofoam?

    • It depends on the size of the specimen, humidity, etc. Generally a week is long enough for all but the most gigantic of beetles, but sometimes a specimen can seem dry but over time the legs droop if it wasn’t completely dried before removing. I tend to do things in large batches and, thus, don’t get around to removing all the beetles I’ve pinned on a block until several weeks (or months) later, so I don’t really have any problems with removing specimens too soon.

      There are two possible ways things can get stuck to the foam – tarsal claws is one (see reply to Crooked Beak), and the other is with larger beetles that can leak fluids from the pin hole on the ventor that then dries between the body of the beetle and the foam on which the body is resting. When I remove a specimen, if it doesn’t come up right away with finger force I use pinning forceps to remove it – pulling straight up is sufficient to break any bond with little risk of damage to the specimen. Some beetle groups seem especially prone to this (e.g. Silphidae), and in that case I will lay down tissue paper over the foam before pinning to soak up most of the fluids.

  3. It would be nice to see a follow-up video where you remove the pins so viewers could see the fruits of your labors. In doing so, you could address the questions Tony asked above. I’d also love to see how you deal with tiny specimens that spread their elytra because they were collected directly into ethanol – something like Euderces or if you want a to show something super-challenging, maybe Lampropterus or one of the small, short-winged genera. I recently had to deal with specimens in these genera and I would like to see how someone else deals with them.

    • Sounds like a good follow up. For specimens with the elytra spread out, I use the forceps to tuck the wings back into a fold and then hold the right elytron down while inserting the pin. Once the beetle is on the foam block I use brace pins to fold the left wing and then hold the left elytron down over it.

      I tend to use smaller gauge pins compared to most—I only use #2 for relatively large beetles (like that Megacyllene and above). Chrysobothris femorata-sized beetles get a #1 pin, while the smaller Chrysobothris-sized beetles get a #0 (the pin size I use most commonly). I will pin some of the larger Agrilus with a #00, but once the size gets smaller than that then I resort to pointing – this includes things like Euderces, Sternidius, Lampropterus, etc. Small gauge pins were bad in the past because the material used for pinning bottoms in drawers was often very hard, but modern plastozoate foams have made even the thinnest of pins no problem to use. The hardness of the body also influences which pin size I use, with harder-bodied beetles getting smaller gauge pins (to avoid risk of breakage trying to insert the pin). Except Zopherus – I drill a starter hole and then use a hammer and a 16d common nail (just kidding!).

      • Interesting how you handle smaller specimens. I think everyone with a decent amount of experience with small specimens has their own subtly-unique approach. I generally don’t use pins smaller than #2 – if I think it is too small to get a 2 through, I point it. I agree that the plastozoate makes using smaller-diameter pins easier. However, just last week I broke the abdomen off of a small wasp because the pin (perhaps a 0 or 00) bent when I was trying to push it through a new det. label. The downside of my “no smaller than a 2” rule is that forces me to brace-pin small specimens on the block without a handle (i.e., not on a pin or point), and this makes getting the elytra closed a bit tricky. I usually end up using at least 5 or 6 pins for a 3-4 mm specimen (it is often hard to see the specimen for the pins!). Once the specimen dries, I remove the brace pins and point it. Positioning without pinning or pointing first also makes it tricky to do anything with the legs – you’re pretty much stuck with whatever you get because the focus is getting the elytra closed properly. It also probably takes longer than it would if starting by pinning, so if I had a ton of material to go through, then the practicality of my method breaks down.

        Have you tried a nail gun for Zopherus?

        • Your approach sounds reasonable if #2 pins are your limit, although with large numbers of specimens I can see this might be rather time consuming (I deal with ~1,000–2,000 specimens/year). Since only my very smallest specimens end up on points I don’t worry too much about the legs and antennae—I can generally just tuck them into place after pointing and they stay in fairly good position.

          When adding labels to small gauge pins, I grab the bottom of the pin with the forceps (that are just about always in my left hand) and use my right hand more to stablize the pin head then push down on it. Yep—everybody has their own subtly-unique method.

          Ah, the “#2-only vs. any-sized-pen” debate—almost as famous as the European-American “card vs. pin” debate!

  4. I like to us my fingers to hold the specimen instead of forceps for the first part – either against the styrofoam, or up off the substrate, under a scope to start the pin. Then, I use 2 pins instead of forceps when manipulating appendages, that way I can jab either pin down into the styrofoam to stick the piece into place, and grab another pin to continue.

    I generally use #2 pins for most things, and for all bracing. When too tiny for #0 (my smallest size pin) I point. If you let the glue dry for a minute or two, it is possible to use brace pins on the pointed specimen to straighten appendages, at least to a certain extent.

    Nice article! You need to include an episode on pulling tails on Buprestids…

    • Fingers work fine as long as the specimen is large enough. With small specimens I get better control and leverage with the forceps (which are always in my left hand). A pin in both hands also works fine, and I have used that method in the past. I moved to keeping the forceps in the left hand because I can use them to actually grab an appendage to move it where I want if it isn’t cooperating with the pins, while with the tips closed it still functions essentially like a pin.

      A wanker-pulling video would be fun if I had a camera capable of doing something that closeup. I generally do genitalia work exclusively under the microscope, even with relatively large specimens.

  5. Thanks for this video, I am always excited to see the curating techniques and specimen processing work-flow of other entomologists!

    From this way this video is lit, it is hard to tell whether you using stainless or black “Japanned” pins. It may be worth mentioning which one you use, to forestall a tragic outcome for some young beetle-pinner.

    (When I first began collecting insects in 1996 I was poor and poorly informed, and used the black pins to mount most of my insects. Since then, many things have happened including my move to New Orleans, and many of the old black pins have severely rusted or otherwise decomposed with time and exposure to insect guts. Since I made this unhappy discovery, I use stainless pins for all my specimens, and only use the cheaper black pins as braces. My subtropical habitat may play a role in this, and pin deterioration may not be as big of a problem in drier parts of the world.)

    • Yikes – I’ve not encountered that problem before, and Missouri is not exactly the Sahara Desert. Still, it doesn’t get as humid here as it does in gator country, and for most of the past decade I’ve had a small room dehumidifier running on constant. I do use the black pins, although a high-quality brand (Elefant).

      Let that be a lesson, kids!


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