Korean cave specimens and the mysteries of old ethanol

Like most museums, there is no shortage of curation projects at the Frost. I’ve described some of the larger ones on our list—rehouse the Beatty Odonata collection, rebuild the teaching collection, renovate the public space—but today was partially dedicated to saving some Korean insects I found in an old box.

Insects collected from cave sites in Korea, housed in substandard conditions. Photo by Andy Deans (CC BY 2.0).

The collecting event labels point to various cave sites in Korea (see PDF of the Korean cave sites at ScholarSphere), where someone collected these insects in the 1960s. Most of them are camel crickets (Orthoptera: Rhaphidophoridae), but many are flies (Diptera) and a few are diplurans (Diplura: Campodeidae and Japygidae). The latter is what grabbed my attention, as not only do we have a relatively small dipluran collection at the Frost, but these specimens were also quite large for Diplura. I think these specimens have been sitting in the same liquid for 50+ years, and judging by its color and the level of preservative it’s time to rehouse them.

Low level of preservative, which also happens to be quite yellow. Not a good environment for this japygid. Also, the stoppers are corroded and seem to have lost their integrity. Photo by Andy Deans (CC BY 2.0).

It’s also an opportunity to dream about digitization protocols for fluid-preserved specimens. This is just a quick attempt at a mock-up, but perhaps we will end up imaging all of our vials like this:

Mock-up of an approach to image fluid preserved insect specimens. Photo by Andy Deans (CC BY 2.0).

It took very little time to set up and was ultra low tech. One can also clearly read the labels and identifier and get the gist of the specimen(s). Others have found ways to industrialize the wet specimen digitization process, of course, or at least produce much higher quality images. See the presentations at iDigBio’s Fluid-preserved Invertebrate Wiki, for starters. We’ll have to find the right balance between efficiency and quality for our material, as well as a workflow that incorporates a storage upgrade. Most of our vials and jars need to be replaced or at least have their caps replaced.

Another issue regarding old preservative was raised by one of my colleagues: Just what’s in that stuff?

Waste ethanol = old preservative. Photo by Andy Deans (CC BY 2.0).

Some of these old jars, with especially rotten specimens in them, smell highly phenolic—repulsive enough to make at least one of us wonder whether extraordinary measures should be taken when handling the material. (Of course we wear gloves and other personal protective equipment when we handle fluid-preserved specimens, and we dispose of all liquids through Penn State’s Environmental Health and Safety system.) I’d love to know what really comprises the variously yellowish, orangish, brownish, and even blackish (especially for old, large millipedes!) liquids we find in these old containers. My quick sifting of the literature yielded many articles about fluid-preserved collections, but all of them limited their chemical analysis to percent ethanol, percent formalin, and/or pH. And the context of each article was specimen health, rather than curator health. Anyway, this question sits on our shelf as a potentially useful and enlightening undergraduate project. Anyone interested?

Back to the Korean caves. It looks like most of these insects came from karst caves in central Korea (ROK), though a few are from “lava caves”. Most of these are closed to the public, and insects collected there in the mid-1960s are definitely worth preserving.

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Friday Night Lice – week 4 answer

Turkish Spiny Mice (Acomys cilicus), a related species of the host for last week’s louse species. Photo by Ettore Balocchi (CC BY 2.0).

Last week we posted images of a spiny louse and asked for guesses about its mammal host. Well, the louse in the photograph, specimen PSUC_FEM 1005427-2, was collected in Mozambique, off of a Southern African Spiny Mouse (Rodentia: Muridae: Acomys spinosissimus). These mice are notorious for their autotomy abilities. That is, they’re capable of shedding body parts if they get attacked by a predator. Acomys spinosissimus can jettison its tail (see photo above, the far left individual), just like many lizards, but other species of Acomys can shed huge areas of their skin (see high profile but paywalled article by Seifert et al. 2012). Ouch! The louse, by the way, is Polyplax acomydis Kim & Emerson, 1970 (Anoplura: Polyplacidae).

Of course many insects and their relatives use this kind of defense—Tipulidae is a famous example—including close relatives of sucking lice, the non-parasitic bark lice (e.g., Psocidae). Many of them are capable of shedding their antennae. What a crazy coincidence!

Mystery louse is a mystery no longer. Meet Polyplax acomydis Kim & Emerson, 1970. Its host is the Southern African Spiny Mouse (Rodentia: Muridae: Acomys spinosissimus. Click to enlarge! Photo by Andy Deans (CC BY 2.0).

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Friday Night Lice – week 4

Mystery louse. What is this female specimen’s host? Click to enlarge! Photo by Andy Deans (CC BY 2.0).

Here’s an unusual louse, with stout setae (hairs) all over its body and a relatively loooooong abdomen. What kind of environment does this louse live in? Well, the host mammal lives in moist savannas and rocky areas, and it’s not native to the New World. We’ll post the answer on Monday, but feel free to post your guess below!

Not especially large grabbers on the ends of its legs. What kind of fur does this louse live in? Photo by Andy Deans (CC BY 2.0).

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Project Snodgrass: annuli and meres

Insect evolutionary morphology is flooded with axioms that every entomologist is familiar with but perhaps cannot fully understand or visualize. Of course, who has not heard about the mouthparts of entognathous insects, that are retracted into the head capsule or about the male genitalia of Zygentoma, that is really “simplified”.

Illustrations from older papers have been cited and redrawn over and over again for contemporary textbooks and reused in insect morphology lectures. Does this mean that old drawings are so perfect that no additional illustration is needed?  Well, they certainly are good enough to deliver simple messages, but are insect morphology axioms really so simple?

During our “Project Snodgrass” we try to visualize anatomical structures using more advanced technologies and then test if they could serve better as educational tools (Andy’s upcoming ENT 432 class will be a good opportunity for that!). We also wonder about the axiomatic nature of these facts—e.g., are entognathous mouthparts really retracted to the head capsule? Is the firebrat male genitalia really so simple?

Lets start with basal hexapods. Most people, perhaps, consider collembolans, proturans and diplurans to be, let’s say, not the most exciting creatures. These soil-dwelling invertebrates have superficially boring morphology: they are small and reduced, and, on the top of this, most of them are white. But these properties actually make them extremely interesting for somebody in possession of a confocal laser scanning microscope (CLSM). Because their cuticle lacks any melanin their white muscle is visible externally, and because they are really small, the laser can penetrate deeply into their bodies. We can easily visualize internal structures.

Non-insect hexapods share some plesiomorphic (ancestral or primitive) morphological traits that can be found in non-hexapod pancrustaceans (“Crustacea”) but never in true insects (Insecta). Perhaps the most well known is the structure of their antenna, which is composed of true appendage segments (see Smartbox below).

Box 1. Appendage segments vs. annuli Insect appendages are encircled and defined by evaginations of the integument (epidermal cell layer and cuticle) that are attached to the body via muscles. The cuticle of most appendages (e.g. antennae and legs) are composed of repetitive, ring-like sclerites (harder plates) that are connected to each other with conjunctiva (more flexible, membrane-like regions). If a ringlike sclerite receives the site of origin of at least one muscle (i.e. musculated), then the appendage portion defined by the sclerite is called appendage segment, if the sclerite is not musculated, the portion is called annulus (pl. annuli) or mere (meres). In true insects, only the first two ring-like sclerites of the antenna, the scape and pedicel, are segments. The rest of the antenna (the flagellum) is composed of annuli that are often referred as flagellomeres. This antenna type is the annulated antenna. Besides Insecta, numerous crustaceans (e.g. Malacostraca) have annulated antennae (the first malacostracan antenna is composed of two or three basal, musculated appendage segments and a distal, sometimes paired flagellum). Segmental properties of appendage portions are, however, not always fully reflected by the morphological terminology. Ring-like sclerites of mouthparts appendages (maxillary and labial palpi) are usually called palpomeres. Although the “mere” suffix refers non-musculated units, most basal palpomeres are musculated (Fig. Box). Should we somehow differentiate them from the non-musculated palpomeres? A possible solution is provided in a recently published paper from our lab group.

CLSM volume rendered image showing the maxilla of Orthogonalys pulchella, posteromedial view, distal to the top-left, doi: 10.6084/m9.figshare.956281

Figure Box. CLSM volume rendered image showing the maxilla of Orthogonalys pulchella, posteromedial view, distal to the top-left, doi: 10.6084/m9.figshare.956281. Image by István Mikó (CC BY 2.0).

Imms (1939) was the first to observe (and publish) that some basal hexapods (diplurans and collembolans) have truly segmented antennae, and he observed the positive correlation between the presence of Johnston’s organ (a mechanosensory organ of the pedicel that detects motion of the flagellum) in annulated antennae and its absence in segmented antennae (Imms 1940).

Imms (1939) illustrated the antenna of numerous arthropod taxa, including Diplura. His illustration has been widely used in the textbooks over the last 70 years (e.g., Chapman 2010). One of the first goals of “Project Snodgrass” was to image and annotate the antenna of a Campodea specimen (Diplura: Campodeidae).

Projections of Composite-2

Figure 1. CLSM volume rendered media file showing the antenna of Campodea sp. (Diplura:  Campodeidae). Image by István Mikó (CC BY 2.0)

As in many other basal hexapods, the cuticle of campodeids is transparent, so we were able to visualize all but one antennal muscle with the CLSM (Strangely, the same muscle, the depressor of the antenna was the only muscle that was not illustrated by Imms).

Figure 2. CLSM volume rendered media file of the proximal part of the antenna of Campodea sp. (Diplura: Campoidea)

Figure 2. CLSM volume rendered media file of the proximal part of the antenna of Campodea sp. (Diplura: Campodeidae). Image by István Mikó (CC BY 2.0).

The campodeid antenna is externally more simplified than most insect antennae. Only trichoid setae are present (their base is connected with ganglionic enlargements that are more advanced in the distal segments with less developed musculature. The first antennal segment (scape) is short and is connected to the head capsule with 4 muscles (internal and extrenal levators, the extensor and the depressor of the antenna (the last one is not illustrated on Fig. 2). It is interesting, that the second antennal segment (pedicel) is also connected to the head capsule (long flexor of segment 3). Segment 3 is moved by only one scapal muscle (flexor of segment 2) because other scapal muscles (basal longitudinal muscle and depressor of segment 3) insert on segment 3. Segment 3, according to Imms (1940) is connected to the labrum by two muscles (short and long extensors of segment 3). Except two diagonal muscles (depressor of segment 4 and 5) the presence of three muscles (dorsal longitudinal and dorsal and ventral extensor) is shared by all segments from segment 3. We did not find the ventral longitudinal muscle, which was described (but not illustrated by Imms 1940).

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Ovipositors: How are they steered?

Hi. My name is Kyle Burks and I’m a new graduate student here at the Frost. I’m working towards my PhD in entomology. Recently I’ve been working closely with István trying to understand how wasps can steer their ovipositor, the stinger-like egg-laying device.

István and I preparing to bleach some specimens

Parasitoid wasps often have incredibly long ovipositors. Instead of using these structures for defense or killing prey, they use them for laying eggs inside of another insect. Their larvae develop in these hosts and eat them alive from the inside before bursting out, just like in the movie Alien. Some of these wasps are capable of bending and steering their ovipositors as they drill through trees or into other insects’ burrows looking for a host to lay their egg inside of. But how do they flex and curl these long needle-like structures since there are no muscles in the shafts of the ovipositor?

Gasteruption floridanum

The “needle” of the ovipositor is not the syringe-like tube I had always imagined. It’s actually a series of three inter-locking shafts. The dorsal shaft is usually the largest and provides support for the ventral shafts, which slide back and forth. The tip usually has barb-like little teeth, and as one shaft slides back, the teeth bite into whatever the wasp is drilling through as the other shaft presses further, and the wasp “drills” through the substrate with the ventral shafts. This motion allows wasps to drill through things as tough as trees because the backwards motion of one shaft braces and supports the forward motion of the other shaft, and the ovipositor drills forward with little net force. This drilling action is accomplished by muscles inside the wasp’s abdomen (remember: there are no muscles in the ovipositor shaft; all of the muscles associated with the ovipositor are inside the abdomen).

Diagram of ovipositor shaft and tip. The back and forth motion of the ventral shafts allows wasps to “drill” through various substrates.

Recently, we made an interesting discovery. I was dissecting two wasps from the family Gasteruptiidae, and noticed that one had died with its ovipositor in a curved position, while the other had a straight ovipositor. When I dissected out the ovipositor and the abdominal muscles that operate it, I noticed a small bulge at the base of the curved ovipositor. It turned out that the bulge was a small hardened plate, called a sclerite, like the ones that make up the exoskeletons of insects.

In Gasteruptiidae, the seventh sternite (S7) sits at the base of the ovipositor shaft.

But what is so interesting about this sclerite at the base of the ovipositor? The underside of all insects’ abdomens are composed of overlapping plates called sternites, and this tiny sclerite is actually a modified sternite! In these particular wasps that I dissected, the seventh sternite (S7) has been shrunk to only a tiny fraction of the size of the other sternites and is located inside the abdomen at the base of the ovipositor. More interesting still, the lower shafts of the ovipositor pass through this tiny sclerite. When the muscles that would normally connect the sternites to each other to allow for movement of the abdomen (see István’s blog recent blog post) pull on the seventh sclerite, the shafts of the ovipositor are also tugged. This causes the ovipositor to bend!

I think it is fascinating the way that evolution can modify common structures in insects. Theses modifications can be to such a degree that the modified structure barely resembles the unmodified structure and can serve a completely new function!

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