Antifreeze and Rancid Snails

I should probably issue an advisory with this entry. I thought about including images, but they would be too graphic even for mature audiences. Suffice it to say that science is often (maybe even usually) a dirty, toxic business. I learned that in a convincing fashion over the last two weeks or so while I was processing raw pitfall trap samples.

You may recall that when I first began work here at the University of Massachusetts I was sorting pitfall trap samples from the 2008 field season. This year’s traps were even greater in number since the students sampled two watersheds instead of one. Last year’s samples had already been placed in alcohol, and were about as “clean” as one could expect. My task over the past couple weeks was to render this year’s specimens workable.

Scientists using pitfall traps face several challenges. First, you need to be able to kill the trapped animals quickly, before they can maul each other. Second, you need a fluid that will not readily evaporate over the trapping period. The trapping interval in this case was one week. The solution, in a literal sense, was propylene glycol.

Propylene glycol is the major ingredient in antifreeze. One of the problems with it is that it is a sweet substance that is attractive to vertebrates like raccoons and other wildlife. While antifreeze was once dyed green, it is now often dyed pink so as to help discourage its consumption. Another problem with this compound is that it has a really corrosive effect on some invertebrates. Beetles trapped in the pitfalls often begin to disarticulate. Snails and slugs quite literally melt. You can only imagine the putrification factor. Actually, you can’t imagine that.

My supervisor is a graduate student who is simply amazing. She has a great deal of responsibility, but never asks anyone to do something she wouldn’t do. So, she cleaned the samples from one watershed, and I cleaned the samples from the other. Neither of us ever ended up passed out, or throwing up all over everything. Quite a feat, especially considering that ventilation in the lab leaves something to be desired, and the odd salamander or frog that was accidentally trapped only amplified the odor. The worst was a long-tailed (or “masked”) shrew, Sorex cinereus, that just about brought me to tears and seriously threatened to initiate my gag reflex.

Still, the slug slime was probably the most aggravating, not as much for the smell, but for the fact that it blocked the screen of the sieves we were using to filter the specimens from the antifreeze solution. Pouring out a sample with slugs meant you had debris everywhere, not concentrated in the center of the sieve where you wanted it.

The protocol was to check to make sure the label inside the container (urine specimen containers, how appropriate) matched the label on the outside. Then you wash the label over the sieve and set it aside. You pour the contents of the container into the sieve, and then rinse it gently with water from a squeeze bottle. Next, you invert the sieve, give it a sharp tap to dislodge as many specimens as possible into a bowl, and rinse the rest off with 70% ethanol (ethyl alcohol). Lastly, you pour the contents of the bowl back into the container (after rinsing the container). Did I mention that small quantities of detergent are also used to break the surface tension of the liquid agent in pitfalls? Some samples contained enough suds to do a load of laundry.

Ah, but that is all behind us now, and I can hardly wait to get a better look at the actual specimens. The pitfall traps were done a week later than last year’s traps, and the difference was quite dramatic. All the spiders in this year’s samples were much, much larger than those in last year’s! More beetles, too, it would seem. Now, if I can just get all my food to stop smelling like rancid snails….

Another day in Entomology Land

Once I finished sorting the pitfall trap samples by order (beetles separate from flies, wasps, spiders, etc), then I set about trying to identify the Hymenoptera (ants, wasps, bees) more specifically. Mind you, I am used to dealing with insects over five millimeters long. In the world of pitfall traps, anything that big is absolutely gigantic. Most of what I’m coming across are minute parasitic wasps and a few ants. The following is a typical sort from order to family level or below in terms of classification.

I start by dumping the vial into a watchglass. The specimens usually stick to the vial, so I have to wash them out with alcohol. Some are very stubborn indeed. Then I stick the watchglass under the microscope and start sorting. A sample may contain anywhere from one to twenty or so specimens, hopefully all of them belonging to the same order (Hymenoptera in this case).

I segregate them by the most specific classification I am able. This often requires the use of scientific documents called “dichotomous keys.” A key is a series of couplets, each couplet describing one or more characters of external anatomy of the insect before you. You find the character(s) that match your specimen, then proceed to the next couplet and so on, until you arrive at a family, genus, or species name. If I am keying out an insect from Massachusetts, and I arrive at a family of insects found only in Sumatra, well, guess who made a boo-boo? This can be no fault of your own, though. I once keyed out a wasp to a genus found only in Japan because my specimen was missing one tarsal (foot) spine that had broken off. Sure enough, the other leg had the full complement of spines. Yes, it is enough to drive you crazy.

The microscope I am using is a binocular stereo “zoom” model that, near as I can tell, takes me up to fifty power (fifty times the size of the insect you are viewing). Even this is not always enough. I had to laugh when I came across one couplet in a key that was illustrated with an SEM! Sure, I’ll jut bop on over to my neighborhood scanning electron microscope, no problem. The University of Massachusetts does have one, but you can’t just barge in with your bug. It is a major exercise to render images of anything under one of those machines, including coating the specimen in a thin layer of gold.

Fortunately, my friend Jeff Boettner was able to rustle-up another key that is much more user-friendly. Got to credit Agriculture Canada for producing such fine works, eh?

Now, if it were only still in print….

It is very gratifying to find that your specimen matches the illustrations in the key, like the sculpturing on the propodeum (hindmost part of the thorax) of this cynipid gall wasp (figure "aa" on the page of the book shown here). Yes, the specimen is standing on its head in this imge.

I am truly learning as much doing this work as I am producing for the university, but then, isn’t that what life should be about? Soon I will share more images of some of the spectacular little insects I’m finding in these samples.

A Day in the life of an Entomologist

I thought I would share just how exciting it is to be in a laboratory doing critically important work as an entomologist, processing pitfall trap samples. I’m kidding, of course, when it comes to the “exciting” part, but it truly is important.

The project I am involved with is striving to develop a protocol for assessing the ecosystem health of forested watersheds, a common yet complex habitat in the northeast U.S. The University of Massachusetts has partnered with the Massachusetts State Department of Environmental Protection and the United States Environmental Protection Agency to carry out this research. Dozens of people have been working in the field and in the lab for nearly ten years already. What started as one person’s project has mushroomed into something much greater. Ok, I know what you’re thinking: How does this affect me, Al Franken?

The habitat under scrutiny here includes many streams that run for most of the year, but run dry in the heat of the short summer. Hence, pitfall traps can be set in mid- to late July in places that are under water the rest of the year. Even then, a freak storm can set the watershed running again and flood the traps, or a beaver can dam the place and do the same thing. The former event happened last year, so I am able to process only a portion of the total pitfall trap samples collected in 2008.

A pitfall trap is a container sunk into the soil (or sand, or other such substrate) such that the lip of the container is flush with the surface of the substrate. Any insects, spiders, mites, and other creatures that come strolling by then fall into the trap. A trap is typically set for a week before being collected and the contents preserved in ethyl alcohol.

The resulting sample I see is in a plastic container with a label detailing the site identification and plot number, and date the trap was set and the date it was collected. Comments indicate the type of location (such as a sphagnum moss mound) and the condition the trap was found in when it was retrieved. I pick out the insects, arachnids, and other invertebrates that I can see with my naked eye, placing each taxon into its own glass shell vial. The vial is labeled both inside and out with the plot number, site ID, and taxon name (Coleoptera – beetles in this photo). The inside label also includes the number of specimens.

I also record the same data on a sheet, counting the number of specimens as I go. Once I’m done with the “big” specimens, I then put the container under the microscope and sort out the smaller things. The effect the alcohol preservative has on the specimens can be dramatic. Spiders fall apart. Even beetles start losing legs and abdominal segments. Springtails, normally pigmented, become literal shadows of their former selves, the alcohol having “cleared” them. It is a strange sight to see a ghostly springtail float across your field of view (cue the spooky music).

The amount of debris in a given sample, and/or the number of invertebrates, can greatly extend the time it takes to process a sample. It can also be very confusing at high magnification to tell a segmented plant part from an insect larva!

Stay tuned for further episodes in the life of an entomologist. Next week, Eric goes insane while trying to identify micro-Hymenoptera to family level….

The Lab

Welcome to Holdsworth Hall on the campus of the University of Massachusetts (Amherst), my home for the next six months. The campus is largely deserted now, graduation having happened about a week ago, and my supervisor, Theresa Portante, herself a grad student, is leading her team in the field for the summer. I will be pretty much alone for the next three months.

There is not much to set my workspace apart from any other scientific technician, but I am grateful for a very nice microscope, illuminator, computer with internet, a small clock radio, and lots of ethyl alcohol. No, the alcohol is not for stress relief! It is the preservative used for the trap samples I will be sorting through.

Right now I am sorting through pitfall trap samples taken on each of last year’s study plots, all of them in wetland habitats that dry out during the heat of summer. Each sample comes in a labeled plastic cup with a locking lid. My job is to segregate the invertebrates contained therein into separate shell vials. Each “order” level of classification gets its own vial. It is not as easy as it sounds. Globular springtails and wingless barklice are easily confused, for example.

Lunch is my chance to get outdoors and breathe some fresh air after sniffing alcohol all morning. My favorite haunt thus far is the ”Campus Pond,” a surprisingly lush, well-landscaped water feature. Aside from being mobbed by ducks accustomed to being fed by everyone that lingers on the shore, I find it is a peaceful spot to enjoy a brown bag meal.

At some point I hope to include an entry in this blog that details what it is like to process a sample. Meanwhile, coming soon….let sleeping wasps lie, giant ichneumons, and other stories from the field.