When Julia was in high school, she built this little cabin in the woods behind her family’s house in central Ohio:
One chilly morning last April, when we stopped there on our way to spending a week exploring the Ozarks, we took the door off the hinges so that its rotten bottom could be repaired (the result can be seen in the above photo). We carried it back to her parents’ driveway and laid it on the ground, with the inside facing up, and I was surprised to see a greater bee fly (Bombyliidae: Bombylius major) clinging to it.
I’ve rarely even attempted to photograph these flies because they are normally in constant motion as they buzz from one spring wildflower to the next like tiny hummingbirds. I guess they have to sleep sometime though, and this one hadn’t yet thawed out from the previous night’s freezing temperatures, so I was able to get a close look.
As the above photo makes clear, it was resting on the webbing associated with a cottony-textured spider egg sac. Fluffy ones like this are usually the work of either orbweavers (Araneidae) or cobweb spiders (Theridiidae). Right next to it was a smooth, disc-shaped egg sac like this one:
Egg sacs of this form are generally the work of hunting spiders (e.g. crab spiders) as opposed to those that spin webs to catch their prey. These particular eggs sacs had distinctive radiating “spokes” of silk around their margins. That may be indicative of a particular species, but I have no idea which one!
Also note that in the crack right next to the above egg sac, there is a web of a tube web spider (Segestriidae: Ariadna bicolor). These spiders wait in their tubes and dart out to grab passing insects that bump into the fine strands of silk that radiate from their entrances. Here are two more tube webs:
While I was taking pictures of all these things, some tiny, previously unnoticed bugs started to warm up enough to wander around on the door. One was this ~3.5 mm long moth:
This is Phyllonorycter celtifoliella (Gracillariidae), which as a larva forms underside tentiform mines in hackberry leaves. Most Phyllonorycter species seem to overwinter as pupae in their mines, but the one time I reared this species, the adult emerged in October. This one found under loose bark in Iowa in January confirms that this species overwinters as an adult. …Okay, it’s dead, but I’m pretty sure the photographer killed it to take the photo. Here is a live one that was among a group of 60 or so found in Nebraska in November, which “were tucked behind the bark of a dead oak tree and were hiding amongst the silk cocoon, larval and pupal exuviae, left over from a Giant Leopard Moth.” So I may have been witnessing this moth waking up in the spring for the first time, after having spent the winter behind the door of Julia’s cabin. (This was not the case for the bee fly, which overwintered as a pupa in the burrow of a ground-nesting bee.)
And wandering about among all of these things were numerous garden springtails (Bourletiellidae: Bourletiella hortensis), at most 1 mm long, which came in a variety of colors:
Beyond what lives behind the door to Julia’s old cabin, the woods immediately surrounding it are the type locality for Phytomyza aesculi and P. hydrophyllivora (Agromyzidae), and are also where I found this lovely moth that bores in Ohio buckeye petioles, and where I photographed a bunch of different bugs visiting narrow-leaved spring beauty flowers a few years ago.
Ever since I made a place on BugGuide to collect photos of them over a decade ago, I’ve been wanting to see (in person) one of those weird wingless gall wasps that can be found throughout the winter. Two years ago I put them on the “winter bug bingo” card I made for the online “Bugs in Winter” course I put together, and I think one participant managed to find one, but I wasn’t so lucky.
Last fall at the end of a game of tennis, I noticed a white oak leaf at the edge of the court with a couple of “oak pea galls” on it, like this one:
This gall is caused by Acraspis pezomachoides (Cynipidae), and knowing that Acraspis is one of the genera with wingless adults, I decided it was time to be a little more proactive in my quest to see one, so I took the leaf home and stuck it in a jar. But alas, I just got a couple of lousy parasitoids.
Last month, while leading a workshop on identifying invertebrate tracks and sign, I picked up a white oak leaf with an “oak hedgehog gall” on it—made by the related species Acraspis erinacei. It was already brown like the one pictured above, but here’s what a fresh one looks like:
I decided to try again, so after everyone had gotten a good look at the gall, I had Julia put it in her backpack.
A week or so ago, I remembered the gall and had Julia fish it out of her backpack. I put it in a vial on a shelf in my office, and every couple of days I’ve remembered to check the vial (most of my other bugs having been put away for the winter at this point).
Last night I checked, and there she was!
Here’s what the gall looked like after she emerged (the exit hole is at the bottom, near where it’s attached to the leaf).
The adults that emerge from round, faceted galls like this—provided that they are not parasitoids or inquilines—are all wingless females. Like most cynipids, Acraspis species have a two-part life cycle. These females climb up to white oak leaf and flower buds and lay eggs in them. Tiny, inconspicuous galls form in the buds (see examples at gallformers.org), and winged males and females emerge from these in the spring. After mating, females lay eggs in midribs of white oak leaves, and the galls that will produce the next set of wingless females develop over the summer.
Virtually all of the bugs I raise are unknowns that have to be killed and preserved to be identified, so it was nice, for a change, to raise one that already has a name and whose whole life cycle has already been worked out. This morning I took her out to a white oak sapling in the woods behind my house and set her on it, thinking maybe I’d get to see her lay some eggs.
No such luck; she didn’t seem to like this sapling, and started making her way down its stem. So I moved her to the trunk of the white oak tree that was right next to it. There, she decided to just sit and preen for a long, long time.
It was below freezing out and my fingers were getting cold, so I decided to leave her to it.
When I stopped to feed the chickens on my way in, I found another wingless weirdo waiting for me by the door of the shed: a female fall cankerworm moth (Geometridae: Alsophila pometaria). ‘Tis the season, I guess! Her eggs will overwinter, hatching in the spring into inchworms that will nibble the leaves of the apple or cherry tree overhead, from whence she presumably came.
More wingless wonders await—I’m always excited to find a snow scorpionfly or snow fly trudging across the snow in winter.
Oh, and tonight I checked the vial with the gall again and discovered that another Acraspis erinacei female has emerged. As noted on gallformers.org (which I highly recommend for identifying galls and reading up on what’s known about them), these galls may contain between two and eight central cells with individuals of A. erinacei (or their parasitoids) developing in them, as well as additional cells around the periphery with inquilines developing in them. Inquilines are (in this case) cynipid wasps that develop inside the galls induced by other cynipid wasps, without (necessarily) killing the original inhabitants. Twelve winters ago, before I knew anything about the life cycle of Acraspis species, I collected some A. pezomachoides galls from under a white oak tree, assuming that what emerged would be Acraspis adults, but instead I got nothing but inquilines, which emerged in early June—thereby skipping the alternate, bud gall-inducing generation of their host species, and appearing just in time to find some developing “pea” galls to lay eggs in. This one’s a male:
The Leafminers of North America project I created on iNaturalist a few years ago has been an excellent way for me to collect new host plant and geographic distribution records for known leafminer species, as well as to identify new mysteries in need of investigation (there are now nearly 1000 rows in my spreadsheet of mystery leaf mines). However, although the vast majority of observations added to the project actually show leaf mines, there are regularly also photos showing things like fungal or viral diseases, or even pine pitch that has landed on the surface of a leaf. Many photos do in fact show evidence of insect feeding, but with “window feeding“—where one leaf epidermis is consumed and the other is left intact—mistaken for leafmining, where by definition both epidermises are left intact and the insect larva feeds between them. Some people simply are not aware of what defines a leaf mine, and post photos of insect feeding where both epidermises are consumed. It was because of this mistake, incidentally, that I happened to see this observation of feeding sign on an elm leaf in Quebec, which I suggested was the first evidence of the Asian “elm zigzag sawfly” (Argidae: Aproceros leucopoda) in North America, which led to this paper confirming its presence here; this discovery was cited in another paper, published a few days ago, as an example of “The benefits of contributing to the citizen science platform iNaturalist as an identifier.”
A distinction that is a lot less clear is where the line is between leaf mines and galls, since many gall-inducing insects do live and feed between the epidermal layers of leaves. In particular, the galls caused by some midges (Cecidomyiidae) are so flat that they don’t seem to have deformed the leaf at all. One gall midge, Monarthropalpus flavus, has even been given the common name “boxwood leafminer,” even though it does cause distinct swellings on leaves, which shouldn’t be mistaken for mines.
Back in July 2013, when I was in the early stages of putting together my leafminer book, I asked gall midge specialist Ray Gagné whether any cecidomyiids could be considered leafminers:
In considering what to include in this book on leafminers I’m working on, I’m wondering if any gall midges qualify. I note that Monarthropalpus flavus is given the name “boxwood leafminer,” but I’m unfamiliar with its gall and don’t know whether it deserves the title any more than any number of other midges that make flattish leaf galls. When I’ve seen occupied spot galls such as those on Smilax, Uvularia, or the Cornus ones I sent you today, they seem very similar to the “blotch” mines produced by sap-feeding gracillariid moth larvae. I suppose what I’m asking, essentially, is whether the size and shape of such galls is determined by the movements of the larva, as in a leaf mine, or whether they are formed in some other way. I haven’t been able to watch them for any considerable length of time, but seeing the larvae positioned non-centrally in these galls suggests to me that they do, in fact, move around, enlarging the spot as they feed.
As to whether they mine the leaf, I have never myself observed their behavior closely or that of the other leaf spot gallmakers. I have read that the boxwood pest, an extremely active larva, feeds in a circular fashion, the reason the gall features an empty circular space. One has to take into account that cecidomyiid larvae have piercing-sucking mouthparts, unlike agromyzids and other miners, so I suppose the cecidomyiids both suck the juice from surrounding cells and possibly cause more cells farther along the periphery to produce additional food. That is analagous to what is going on in the Macrodiplosis leaf swelling on oaks, except there you wouldn’t call the cecidomyiid larvae leaf miners. For that reason, I wouldn’t list the Parallelodiplosis or the Mon. flavus as leaf miners, although I certainly can understand why you might do so for completeness in a leafminer book.
The Cornus (dogwood) leaf spots I mentioned and the Parallelodiplosis Ray mentioned were both referring to Parallelodiplosis subtruncata, which I wrote about in January 2014. In that post I was focusing on the “green island” phenomenon I’d seen in connection with galls of this species I found in Idaho in September 2012, and I made no mention of the observations I’d made of the same species in my own yard just a few months before I wrote the post.
On September 26, 2013, I noticed these two galls on a leaf of alternate dogwood (Cornus alternifolia) at the edge of my yard.
Flipping the leaf over, I saw that each gall still had a larva inside.
So I decided to monitor their progress to see if the galls got any bigger. Here they are on September 27:
September 30 (with some condensation inside after it rained the previous day):
October 7, after the larvae dropped to the ground.
If you compare each photo to the previous one, you’ll see that the size and shape of the little blisters containing the larvae doesn’t change at all. Each larva is just squirming around within the cavity that the plant has created in response to its presence, drinking the juices. So although these galls are superficially similar to leaf mines, the larvae are not doing any actual mining.
I should mention here that some gall inducers do have chewing mouthparts and actively excavate the tissue within the galls. Those that come to mind that feed in leaf galls include the moth Heliozela aesella (Heliozelidae) on grape, and “Pontania” sawflies (Tenthredinidae, now placed in the huge genus Euura) on willow. But their galls are obvious swellings/deformities in the leaf tissue, and no one would confuse them with mines.
I should also mention, maybe, that not all leaf mines are flat. “Underside tentiform mines” are sometimes mistaken for galls because of the way they distort the leaf. However, they start out as normal, flat mines excavated by the larvae, and the distortion is the result of the older larva spinning silk within the mine; there is no manipulation of the growth of the plant as there would be in a gall. The silk contracts as it dries, causing the edges of the mine to draw together, which results in a wrinkled lower epidermis and the upper epidermis buckling to form a roomy “tent.” This is a useful adaptation for avoiding parasitoids and predators, since the larva now has three dimensions in which to try to escape, instead of just two.
Incidentally, here’s what the Parallelodiplosis galls looked like on the upper surface on October 7, when I found them empty:
By October 11, there was a bit of a “green island” effect, but obviously not of any consequence to the well-being of these particular larvae.
Things have been quiet around here on BugTracks lately, due to another busy field season (and due to spending a good chunk of my summer computer time updating the 300+ page Asterales chapter in Leafminers of North America, which is now finished). Any day now I’ll start going through my photos from this spring and summer and will be posting the highlights here, as soon as I finish up a few papers I’ve been working on about—you guessed it—leafminers and sawfly larvae.
In the meantime, a box of 2023 Leafminers of North America calendars has just arrived, and as with last year, I will send a copy to anyone who makes a donation of at least $30 (the amount WordPress charges me each year to keep this blog free of annoying ads) before the end of November, which you can do here (select “Send,” and then include your mailing address in the notes). As with last year’s calendar, each month shows a whole leafminer life cycle instead of a single full-page photo per month.
Also, last month when the “January” issue of Proceedings of the Entomological Society of Washington was finally published, it included four papers of mine, three of which reported on discoveries made during my 2020 inventory of leafminers and sawfly larvae in my yard (and the fourth included something I found in my neighbor’s beaver meadow). Here’s a quick summary of those discoveries, with links back to my original blog posts about them.
Eiseman, Charles S. and Owen Lonsdale. 2022. First North American record of Phytomyza origani Hering (Diptera: Agromyzidae), a leafminer of cultivated herbs in the mint family (Lamiaceae). Proceedings of the Entomological Society of Washington 124(1): 177–183.
As I had assumed, this leaf mine I found on the oregano by my front door on June 20 proved to be the first North American record of Phytomyza origani. Since this species was thought to be a strict specialist on oregano, it was a little surprising when the leaf mines on apple mint (which I first noticed in Julia’s water glass) turned out to be the work of the same fly. Coincidentally, within a month of my rearing P. origani from apple mint in my yard, Yuliia Guglya reared it from other Mentha mints in Ukraine, as I learned about when I was given her paper to review last spring (which happened to be the same week that Owen Lonsdale examined and identified my flies).
Eiseman, Charles S., David R. Smith, Bill Sheehan, and Tracy S. Feldman. 2022. Macrophya Dahlbom spp. (Hymenoptera: Tenthredinidae) feeding on Asteraceae. Proceedings of the Entomological Society of Washington 124(1): 39–45.
This little cutie that I found on the underside of a Canada goldenrod leaf by the driveway on June 13,
as well as this snazzy older larva I found on late goldenrod by the chicken run on June 21,
both turned out to be Macrophya senacca, a species that Gary Gibson had described when he revised the genus Macrophya in 1980, but nothing had been known about its immature stages or host plants until now. Both larvae burrowed into jars of soil and emerged as adults the following spring.
Eiseman, Charles S. and David R. Smith. 2022. A review of the Nearctic fern-feeding sawflies (Hymenoptera: Tenthredinidoidea), with new host records and larval descriptions. Proceedings of the Entomological Society of Washington 124(1): 18–38.
On June 6, while gazing down at a garden from the back deck, I spotted this larva on a clump of lady fern,
and once I’d brought it inside I discovered this tiny larva on the same frond,
It was a bit tricky to keep track of who was who, when more eggs and larvae kept coming in every time I collected more bits of lady fern, but I was able to rear all three species to adults. The first larva emerged as an adult the following spring, and turned out to be—as I’d guessed it might be—Strongylogaster macula, which had previously been reported only from Europe and Canada.
The second larva bored into a sumac twig I offered it and emerged as an adult the following spring, revealing itself to be Thrinax albidopicta, whose larva hadn’t been described before (and previous host records attributed to this species were based on misidentifications). Other larvae of that species that I collected after the first one emerged as adults without any diapause, the first one on June 22. As they get older, larvae of T. albidopicta develop adorable little black hats.
And the third larva turned out to be Aneugmenus flavipes, a species previously known to feed only on bracken fern; the first adult I reared from lady fern emerged on June 29.
Eiseman, Charles S. 2022. New rearing records reveal Phytosciara greylockensis Eiseman, Heller, and Rulik (Diptera: Sciaridae) is a polyphagous leafminer of herbaceous plants. Proceedings of the Entomological Society of Washington 124(1): 174–176.
The title of this one pretty much says it all; when Julia and I first discovered Phytosciara greylockensis at the 2016 Berkshire BioBlitz on Mt. Greylock, the larvae were feeding on bluebead lily, but when we were conducting a survey for rare dragonflies along the shore of a small river in 2020, I found them on buttercup, violet, water pennywort, and sensitive fern all within an area of about a square meter, and then a couple of weeks later I found one on a monkeyflower leaf in my neighbor’s beaver meadow.
And for completeness, here are two more papers that were published within a day after the above four came out. These two aren’t about my yard and I wasn’t the lead author, but unlike the first four they are open access so you can follow the links below to read them online if you want (if you’d like a PDF of any of the others, let me know).
Xuan, Jing-Li, Sonja J. Scheffer, Owen Lonsdale, Brian K. Cassel, Matthew L. Lewis, Charles S. Eiseman, Wan-Xue Liu, and Brian M. Wiegmann. 2022. A genome-wide phylogeny and the diversification of genus Liriomyza (Diptera: Agromyzidae) inferred from anchored phylogenomics. Systematic Entomology 2022: 1–20. (full article)
Chen, Taibin, Xiaohua Dai, and Charles Eiseman. 2022. A checklist of gymnosperm-feeding leafminers (Arthopoda, Insecta) in North America and Europe. Biodiversity Data Journal 10: e91313. (full article)
As I go through my Leafminers of North America e-book and update each chapter for the (now nearly complete) second edition, I’ve been putting together a spreadsheet of mystery leaf mines that need further investigation. There are now over 700 rows in that spreadsheet, and new mysteries continue to be added faster than old ones are solved.
Today Julia and I drove 4.5 hours (round trip) to explore a site in western Vermont with the hope of investigating one of these mysteries—a nepticulid moth that so far is known only from a few empty mines scattered across eastern North America. I first became aware of this species several years ago when Erik van Nieukerken told me that, while perusing the pressed leaf mines that are preserved in the Canadian National Collection, he had seen mines of an unknown nepticulid on dogwood leaves from Ontario. Later, while working either on the dogwood key in my book or on my first paper on agromyzid flies with Owen Lonsdale, I discovered that I had photographed one of these mines in western Iowa in September 2012 and had filed it under Phytomyza agromyzina. P. agromyzina is a leafminer of dogwoods that is common across North America as well as in Europe and Asia; here is a typical example on a leaf of alternate dogwood (Cornus alternifolia):
Here is the mystery moth mine from Iowa, found on roughleaf dogwood (C. drummondii):
Both are simple linear mines, but whereas the frass trail of the agromyzid fly alternates from side to side (this is most evident toward the end of the mine), the nepticulid moth deposits its frass in a central line. This is because the fly larva lies on its side while it feeds, periodically rolling over onto the other side, while the moth larva lies on its back or on its belly.
The central frass line is more clearly visible in this fresh (but apparently aborted) mine I photographed on flowering dogwood (C. florida) in southeastern Ohio in early August of 2016:
The third time I found one of these mines was a few months later, in October 2016, at the site in western Vermont we visited today:
This example was on gray/panicled dogwood (C. racemosa). Although the frass pretty well fills the width of the mine and it’s not easy to see that it forms a central line, on close inspection it is made up of closely spaced zigzagging arcs, which are characteristic of Nepticulidae.
And I have never seen another of these mines since, including today. (The trip was still worthwhile, since we got to explore a beautiful place as well as add two species to the list of moths known from Vermont and collect a couple of mines of another mystery moth.) Nor has anyone else, as far as I know. Just to be sure of this, this evening I reviewed the ~100 iNaturalist observations of Phytomyza agromyzina that I hadn’t already looked at. As a result of this exercise (which involved weeding out a number of observations that didn’t show P. agromyina, or even leaf mines or dogwood in some cases), there are now 550 verified observations of P. agromyzina on iNaturalist, and I have one more mystery leaf mine on dogwood to wonder about, thanks to this leaf that Jeff Clark photographed in Virginia last October:
This mine has a central frass line as in the nepticulid, but the silvery-whitish color indicates that it was formed entirely in the leaf’s epidermal cells, as opposed to in the mesophyll. Also, the frass is in a continuous line rather than composed of tiny particles, indicating that the larva was consuming only the liquid contents of the cells. This type of mine is characteristic of moths in the family Gracillariidae, and this is the first I’ve heard of a gracillariid making a long, linear mine in dogwood leaves (there is an unknown Marmara species that makes linear mines in dogwood stems, but that’s another story). This looks very much like a Phyllocnistis mine, but without seeing that it ends in a silken pupal chamber, I can’t be sure that that’s what it is*. Here’s a link to Jeff’s original observation, which includes the full-resolution photo as well as a second photo showing another leaf with the same type of mine.
So if there are any dogwoods near you, I’d greatly appreciate it if you could keep an eye out for either of these mystery mines, and please collect any you find! Even mines with dead larvae inside would be tremendously valuable, since it may be possible to match them to adult specimens through DNA barcoding. And if you find any occupied agromyzid mines, those would be worth collecting for rearing too: I recently reared some adults from these mines that Owen Lonsdale identified as Phytomyza notopleuralis, which probably should be synonymized with P. agromyzina, but more specimens are needed to demonstrate that there is no clear line between the two species. For further details about that, see:
Eiseman, Charles S., Owen Lonsdale, John van der Linden, Tracy S. Feldman, and Michael W. Palmer. 2021. Thirteen new species of Agromyzidae (Diptera) from the United States, with new host and distribution records for 32 additional species. Zootaxa 4931(1): 1–68.
* Edit, 8/21/2022: Natalia Kirichenko just reminded me about this paper, published four years ago, which discusses four species of dogwood-mining Phyllocnistis species in Northeast Asia, describing three of them as new. Could the Virginia species be one of them? All four have publicly available DNA barcodes, so that question would be easy to answer if someone can collect a mine with a larva or pupa inside…
Today I break my four-month silence to bring you this:
Yesterday morning while we were eating breakfast on the back deck, Julia exclaimed something like “The poop beetles are eating the groundcherry!” This wasn’t news to me; a week or so ago I had noticed the tiny larvae, with poop piled on their backs, on a leaf of one of the potted groundcherry plants we had overwintered indoors with the hope of actually getting some fruit out of them this year. But when I looked over at the plant now, I saw the reason for her alarm: the top of the plant had been reduced to a “Y” of two blunt, naked branches, and when I went over to inspect, I saw that each fork of the Y was topped with a “flower” of larvae that were working together to munch the branch down to nothing. I thought their symmetrical arrangement produced an image that, while somewhat stomach-churning—especially in the middle of breakfast—was also oddly compelling. So of course I ran to get my camera. And then I gathered up all the larvae and threw them to the chickens, even though I knew they would react exactly as they did: come running up excitedly to see the latest offering, then stop suddenly a foot or two away, cock their heads quizzically, and walk away.
If you’re not familiar with these larvae, here’s a side view of the same scene to give you a better sense of what we’re looking at:
They are larvae of the three-lined potato beetle (Chrysomelidae: Lema daturaphila, or another similar Lema species). And being the good botanists that they are, they know that groundcherries (Solanaceae: Physalis) have nothing to do with cherries (Rosaceae: Prunus), but belong to the nightshade family, along with potato, tomato, eggplant, and goji.
Here’s an adult found on our deck railing last June—when I don’t think we had any nightshades there to speak of:
And another on one of our goji bushes seven years ago, being inspected by a group of Lasius ants.
Carrot (Apiaceae: Daucus carota) is native to Europe but widely cultivated and has become a ubiquitous weed in North America (also known as Queen Anne’s lace), so you’d think we’d have a pretty good handle on what bugs eat it by now. You’d be wrong.
Black swallowtail caterpillars (Papilionidae: Papilio polyxenes) are well known to feed on a variety of native and nonnative plants in the parsley family, and I often see them munching away on wild carrot leaves in my yard…
…but I’ve also come across a surprising number of other carrot-feeding insects in my yard that don’t seem to have been documented before. In 2017 I reared two adults of the micro-moth Epermenia albapunctella (Epermeniidae) from larvae that initially made tiny mines in the leaves, later feeding externally from little webs. This species was not previously known to feed on carrot, or to mine leaves. When I did my intensive year-long cataloguing of leaf and stem miners in my yard in 2020, it was #40:
And then #179 was a stem-mining fly; the relevant section of that post is repeated here:
Leaf (stem) miner #179: Ophiomyia sp. (Agromyzidae), on wild carrot / Queen Anne’s lace (Apiaceae: Daucus carota). I was excited to find this mine on the evening of September 5:
No Ophiomyia is known to feed on wild carrot, but Julia and I found a bunch of similar mines on an isolated clump at Black Rock Forest in New York late last August while conducting our survey for leaf-mining moths there. The puparia in those mines were all black, and only eulophid wasps emerged from them. The puparium in the above mine (visible as a bulge along the upper margin of the stem) was whitish; unfortunately it turned out to be empty already.
In this close-up, the pair of little black anterior spiracles of the puparium are visible poking through the stem epidermis at far left, and there is a longitudinal opening associated with those—along with a more conspicuous transverse slit to the right of them—indicating that the fly has already emerged. I spent a good chunk of yesterday pulling up wild carrot stems around the yard, and I found six stems with intact puparia (plus one more empty one, and one or two that seemed to still have larvae in them). To give a sense of how sparsely distributed these mines are, this is how many stems I had to inspect to find a half dozen of them (note Brenda in the background; she followed me around for most of the time that I was pulling them up, and was often literally underfoot):
All of the mines were confined between two nodes in the stem as in the example shown above. John van der Linden has observed similarly constrained stem mines (both agromyzid and Marmara) on Ageratina altissima, Polymnia canadensis, and Veronicastrum virginicum in Iowa.
That’s where I left the story, so I’ll pick it up from there. At the beginning of October 2020, this braconid wasp (subfamily Opiinae) emerged from one of the six puparia I’d collected on September 6.
When I looked at the puparia under magnification to figure out which one it had come from, I discovered that two of them had exit holes (one of them had evidently only looked intact to the naked eye), so now I just had four left. I assume this braconid came from the puparium with the more conspicuous hole:
On October 16 I put all of my rearing projects into the fridge for the winter; I took them back out on March 1 of last year. On March 29, another parasitoid emerged: this time, a pteromalid in the genus Herbertia.
Nothing ever emerged from the remaining three puparia. So naturally I was watching closely for the first mines to appear last summer, and I spotted the first one on August 4. This prompted me to spend the next couple of hours pulling up every wild carrot stem in the section of my front yard bounded by the driveway, upper vegetable garden, and shed, yielding a total of four mines: three in stems, each of which already contained a greenish-white puparium like the one toward the right side of this photo…
…and one mine in a leaf stalk, in which a larva was still feeding (at right):
I suspect that in this species a black puparium is an indication that a parasitoid will emerge, and a whitish puparium means there is some hope of a fly emerging.
When I went to pull up the last stem before quitting for the day, I was shocked to discover that not only did it have two mines in it, but they were Marmara (Gracillariidae) instead of Ophiomyia.
There are no previous records of Marmara from wild carrot, or from anything else in the parsley family for that matter. But the continuous central frass line in these mines told me at once they were Marmara; in Ophiomyia stem mines the frass is much less conspicuous, and it is deposited either in widely spaced grains or in little strips that alternate from side to side. One of the Marmara larvae is visible to the right in the above photo, but it’s a little hard to make out. Here’s a close-up of the other larva, with its head pointing toward the upper left corner:
Needless to say, I stuck this stem in a ziplock bag to see if I could rear the larvae to adult moths.
On August 6, I pulled up all the wild carrot stems from another section of the yard, found a few more puparia, and three days later an adult Ophiomyia emerged from one of them! For some reason it was already dead when I found it, even though I’d been checking the rearing vial regularly.
It’s a female, which means that when Owen Lonsdale gets around to examining it, all he’ll be able to tell me is that it’s some kind of Ophiomyia. Without male genitalia, I’m no closer to getting a name on this fly than I was when I just had parasitoid wasps.
On August 14, another fly emerged! …Another already-dead female.
On August 13, a braconid emerged—this time belonging to the subfamily Alysiinae.
Alysiine braconids have weird, outward-facing mandibles that they use to pry open the host fly’s puparium along an existing line of weakness at the anterior end, so that the emerging wasp leaves an opening similar to the one an emerging fly would leave, as opposed to the ragged-edged hole an opiine braconid chews.
On August 12 I stripped yet another section of my yard of its wild carrot stems, found a few more puparia, and two days later a fly emerged from one of them! …Another sorry-looking female.
Another alysiine braconid from the August 6 collection emerged that day. On the 15th, the fly that I had collected as a petiole-mining larva on August 4 emerged as an adult… another lousy female.
It’s still a mystery to me how one fly after another managed to make itself so dead in such a short amount of time. On August 17, another adult emerged—from another carrot-pulling session on August 10 that I guess I neglected to mention—and this one was alive!
…But it was just another female. How many females do I have to rear before we decide that this must be a parthenogenetic species, and there never will be any males? I don’t know, but more than five.
On August 21, a pteromalid emerged from one of the August 6 puparia; this time a miscogastrine rather than a herbertiine.
On August 22, another alysiine braconid.
On August 25, one of the Marmara adults emerged! It had rubbed some of its wing scales off in the bag, but that was okay; as with the flies, distinguishing them is all about the male genitalia.
Two days later, the other Marmara! It was a little drunk for some reason and kept flipping on its back, so its wings were even more rubbed than the first one’s, but no matter; they both had abdomens, and that’s what counts.
I pretty thoroughly inspected the carrot stem from which they had emerged and couldn’t find either moth’s cocoon. Some Marmara species exit their mines to spin cocoons, and others cut out a little flap in the stem epidermis at the end of the mine and spin their cocoon under that. I was too busy with fieldwork to keep looking right then, but I wanted to know what this species does, so I kept the stem in the bag to examine again when I had more time.
On September 1, another miscogastrine pteromalid emerged from one of the Ophiomyia puparia.
On September 9, this little dark-winged fungus gnat (Sciaridae) appeared in the bag with the Marmara-mined stem.
I had continued to pull up wild carrot stems from my yard throughout August. An Ophiomyia puparium I collected during the August 21 session produced this eulophid wasp (subfamily Entedoninae) on September 6:
On October 11, I looked over at that bag with the Marmara-mined stem in it, and there were several more of those dark-winged fungus gnats in it. There was also a weevil:
I had noticed some powdery frass coming out of a hole in that stem a while back, so I knew there was some kind of larva boring inside it, and it didn’t come as a complete surprise to see the weevil in the bag. With a quick internet search, I learned that there is a species that looks sort of like this called the carrot weevil (Curculionidae: Listronotus oregonensis); the sources I found mentioned it feeding on the root, but I figured the focus was on that because that’s the part people care about, and it seemed plausible that the same species could also bore in carrot stems.
I surmised that the fungus gnats must have been developing inside the weevil’s tunnel, feeding as larvae on its frass and the damaged/dying plant tissue. By November 5, a total of 60 of them had emerged. Sixty!
I sent the weevil to Bob Anderson at the Canadian Museum of Nature, along with some others I’d accumulated over the past few years, and he told me, “The mystery weevil from Daucus is a Listronotus but I don’t think it’s oregonesis as it’s a bit small for that species.” No comment on what he did think it was.
I sent the fungus gnats to Kai Heller in Germany, and he reported: “All individuals belong without doubt to the same species, namely Bradysia impatiens. This is the common greenhouse midge, which has a worldwide distribution. . . Unfortunately this is not a very interesting record.”
I sent the flies to Owen Lonsdale, who hasn’t had a chance to look at them yet, but we already know what he’ll say, since no males ever emerged.
Probably no one will ever look at the wasps.
As for the Marmara specimens, they came along at an inopportune time, when Julia and I were both impossibly busy, and they were part of a batch of moths that were left on spreading boards for several weeks in a box that had no mothballs left in it. Some time in the fall, we discovered that booklice had eaten most of the abdomens in the box, and the Marmaras were not spared. In fact, one of them is now nothing more than a thorax on a pin.
I’m reminded of this tragedy every day, because back in June, a booklouse appeared in my camera’s viewfinder:
It hung out in the upper left corner there for a few days, then it disappeared for a week or so, but then it reappeared, changing position a few times, until it finally died right near the middle of the field of view, where it still is to this day.
On the plus side, just a few more months until more wild carrot stems start popping up all over my yard; maybe it will all go better this time around!
Well, the mystery presented in my previous post was solved within an hour of my posting it, but before I get to that, let me back up and chronicle the adventures Julia and I have had in moth dissection so far. As I mentioned previously, we started out by practicing on some specimens I collected for a class over 15 years ago; thanks to the professor’s eagerness to be done with his position at UVM and move on to his new one at UC Berkeley, all of the collection data were thrown away and all we know is that I collected these moths somewhere in New England (mostly in Burlington, VT, but I know I got some in Massachusetts and Maine). In this post I’ll just show the males; females are an entirely different matter, and we’ll deal with them some other day.
On February 4, Julia dissected the first batch, beginning with this noctuid—which, because the wings haven’t been spread properly, I’m thinking I actually found dead on a windowsill in our house at some point and just stuck it on a pin and put it in the box of unimportant specimens from grad school in case I had a use for it at some point. [Edit: It’s a “bicolored sallow,” Sunira bicolorago. I did in fact find it dead on a windowsill in the living room last January but had neglected to label it.]
That day Julia took photos of her dissections by holding her phone up to one of the microscope eyepieces. For this one and the next, she used a stain called “Orange G” instead of Eosin Y, and no Chlorazol Black. Just experimenting. Here are the genitalia of the moth before the coverslip was added…
…and after. As you can see, the angle at which you’re viewing the genitalia, and the way in which they happen to be smooshed when slide mounted, has a big effect on their appearance. For this reason, these days permanently slide-mounting genitalia is frowned upon, and the alternative is to remove them from the slide when done examining them and put them in a tiny vial in some viscous fluid (a lot of people use glycerin; Terry Harrison uses lactic acid because it helps to neutralize the potassium hydroxide, which otherwise can continue to slowly dissolve the genitalia for years and may ultimately destroy the specimen), which is then placed on the pin together with the rest of the moth.
Next up was this substantially smaller moth, a crambid (subfamily Crambinae, and I think tribe Crambini), which despite the unspread wings I definitely did collect for that class. [Edit: It’s Agriphila ruricolellus, the “lesser vagabond sod webworm.”]
Julia’s notes for this one say “very hard to spread,” referring in this case to the valvae of the genitalia, which are normally clamped together and have to be spread apart to examine and photograph. I guess because it didn’t turn out too well, she didn’t take a photo, so here’s a quick one I took just now using our new DSLR-to-microscope adapter. The lack of any bubbles in this one may be due to the fact that she used Hoyer’s mounting medium instead of lactic acid, or maybe she was just lucky. We haven’t found any correlation between how we put on the cover slip and how many bubbles there are, other than that any attempt to improve the situation always makes it worse.
…with completely different genitalia. (Back to Eosin Y, lactic acid, and phone camera for this one.)
On February 6, Julia dissected three more males. The first was this moth that I had labeled as a tortricid when I took the class, but I’m pretty sure it’s actually a gelechioid of some sort. [Edit: Yes, it’s Depressariidae: Machimia tentoriferella, the “gold-striped leaftier.”]
The genitalia are just in the tip of that wee abdomen. In this next photo I’ve included the abdomen along with the removed genitalia (with phallus detached) so you can see the relative size.
And here’s the same photo cropped down to just the genitalia.
The second moth of the day was this one, which I’d labeled as a noctuid… moths with a wingspan of more than 1 cm or so really aren’t my thing, so please chime in if you know better. [Edit: it’s a notcuoid, but the family is Erebidae: Palthis asopialis, the “faint-spotted palthis.”]
The phallus is too far away on the slide to include in the same photo with the rest of the genitalia, so here are two separate shots. (This was another “Orange G” one).
And Julia’s last male dissection to date was this pyralid, which I did a terrible job of pinning. I’m still no good at it, which is why Julia gets to pin all of the moths around here.
It’s the Indianmeal Moth (Plodia interpunctella), a common pest of dry stored foods, and I must have grabbed it from my kitchen. These moths were abundant in the house where I lived in Burlington, due to one of my housemates essentially moving in with his girlfriend and leaving a cupboard full of neglected packaged foods.
Not Julia’s best work, but the genitalia do match what’s shown here if you squint your eyes just right.
This is the other thing that was mounted on the slide… I wasn’t sure if it included the phallus, but I guess the object at the bottom of the photo pretty much matches the shape of the phallus shown at the above link.
On February 12, it was my turn to give dissection a shot. I started with this big ol’ hemlock looper (Geometridae: Lambdina fiscellaria). Yes, it’s pinned upside down.
Not having a phone, I tried taking photos with the Olympus TG-4 held up to one of the eyepieces.
Based on the example shown here, the asymmetrical claspers (?) are typical of this species.
After dissecting a female of a different geometrid species, I tried this gelechioid, which is the same species as Julia’s #4 (they were collected together and have the same wing pattern and genitalia). [Edit: Machimia tentoriferella again.]
For another example of how the appearance of the genitalia can change depending on how you look at them, notice how before I flattened this with the coverslip there is a pair of “lips” directed straight at the camera near the top of the structure…
…and after adding the coverslip, those “lips” are smooshed downward.
Since the valve on the right was folded, I took off the coverslip and tried again. My second try was a big improvement, except for all the darn bubbles.
I tried one more time and miraculously ended up with fewer bubbles, as well as more widely spread valvae (not necessarily an improvement, but just to point out that the orientation of the valvae is of no significance when it comes to trying to identify species).
On February 17, the DSLR-to-microscope adapter had arrived in the mail, and since I hadn’t ruined any (male) specimens so far, I decided to try out a couple of unknowns and see if I could actually use genitalia to identify them. First up (being the biggest) was the willow leaf-tying crambid I wrote about last time.
To summarize how my ID attempt went, I tried Munroe’s (1976) key to the genera of Pyraustinae and didn’t reach a satisfying conclusion, other than that my moth clearly was not one of the species illustrated in that publication. I posted a link to my blog post on Twitter and a couple of moth-y Facebook groups, along with a cry for help, and about 45 minutes later Chris Grinter replied “Looks like a great match to Framinghamia helvalis“, with a link to this image in the Moth Photographers Group North American Lepidoptera Genitalia Library. He was clearly right, and early the next morning Steven Whitebread and Aaron Hunt independently told me the same thing.
The main problem was that this moth is in the subfamily Spilomelinae instead of Pyraustinae! It had occurred to me to check before I got started, since Munroe distinguishes the two groups right upfront.
Pyraustinae: “Praecinctorium weakly bilobed, the lobes parallel and longitudinal, diverging at an angle ventrolaterad from tip of praecinctorium proper; forewing of male with straplike frenulum-hook arising from costa near base, in addition to the retinaculum (a group of stiff scales arising farther back on the wing and also helping hold the frenulum in place) (Forbes, 1926: 331-332); valve of male genitalia almost always with basally directed clasper, one or more of its basally or dorsally directed lobes usually with conspicuous setae or erect scales; bursa of female genitalia almost always with rhomboidal or mouth-shaped, spinulose, transversely keeled signum.”
Spilomelinae: “Praecinctorium strongly bilobed, the lobes transverse and often projecting visibly beyond each side of base of abdomen; male with retinaculum but no frenulum-hook; valve of male genitalia with its clasper, when present, directed distad, rarely with any obvious basally directed lobe, and without conspicuous erect setae or scales; signum of female various, but not rhomboidal.”
Isn’t it nice how the only figure reference is to something in another publication from 50 years earlier? My eyes sort of glazed over as I read the the first two characters (the praecinctorium turns out to be a structure on the abdomen that is specific to Crambidae), and I thought, “well, the tip of what I’m calling the clasper is pointed downward, and those guys who told me it’s a pyraustine probably know what they’re talking about…”
The other problem was that Munroe (1976) did not cover the subfamily Spilomelinae, and to this day there still is not a monograph covering the North American species of this group. Munroe did, however, illustrate the genitalia of Framinghamia helvalis in a 1951 publication. The genus got its name from the fact that the type specimen was collected in Framingham, Massachusetts. F. helvalis is the only known species.
Steven Whitebread pointed out that I could have identified this moth by searching the Mass Moths website for species of Crambidae known to feed on willow (and known to occur in Massachusetts, like all species on the site). There turn out to be just two, and the other looks nothing like F. helvalis (and there is just one record of it from Massachusetts). F. helvalis has not been documented on Nantucket before, but has been reared from willow on Martha’s Vineyard.
Of course, wanting to put a name on the moth was only part of why I dissected it. I also wanted experience using genitalia for identification, and to that end, here is an edited version of the diagram I included in my last post, with corrections in red (many thanks to Chris Grinter, Jim Hayden, and JoAnne Russo for their feedback).
It turns out that the genus Framinghamia is characterized by not having an uncus. And apparently “distal” refers to the end of the aedoeagus/aedeagus/phallus that is distal with respect to the moth’s head, not distal with respect to the rest of the genitalia (if you look back at my previous post, you’ll see that the phallus was pointing the other way when in situ). Jim Hayden informed me that the pointy thing that appears in Tony Thomas’ image right behind where the uncus would be (and also shown in Munroe’s 1951 drawing) is the anal tube, which is supported by the subscaphium (the sclerotized ventral wall). It is often removed from dissections because it is easily torn, not informative, and full of poop.
Having successfully dissected this moth, and having not yet gotten bogged down in trying to identify it, I moved on to my first leafminer, which was a Phyllonorycter (Gracillariidae) that Mike Palmer had reared from black cottonwood (Salicaceae: Populus trichocarpa) in Montana in 2017. The leaf mines are shown here.
I chose this moth because I have several specimens from the same rearing (so it was okay if I wrecked one), because the Salicaceae-feeding Phyllonorycter species are among the only North American gracillariids for which genitalia illustrations have been published (Davis & Deschka 2001), and because the key to species is based entirely on male genitalia.
To sex gracillariids I don’t go looking for tiny bristles on the leading edges of the undersides of the hindwings; I just look at the tip of the abdomen. The first specimen I looked at was a female, but the second was a male:
As with the Framinghamia, I soaked this moth’s abdomen in Chlorazol Black for an hour or two instead of just using the red stain. Here is the result (maybe more black than needed, but better too much stain than too little if I want to be able to actually see anything; the Eosin Y hadn’t had much of an effect):
Did I mention that this moth had a wingspan of 8 mm and the abdomen was only 2 mm long? So this thing I was now left with was less than 1 mm long, and I needed to spread those narrow valvae open somehow. Here’s a side view of the same thing, with the tiny tiny detached phallus floating off to the right:
Looking at this second view, I was confused, because there seemed to be three layers of things to deal with. I interpreted those two skinny things to the left and front as the valvae, and the long, central structure as the tegumen, which left me scratching my head about that last thing to the right. In Davis & Deschka’s illustrations, in addition to the detached phallus there is another part that is illustrated separately: the eighth abdominal sternite. I thought this sternite would be part of the soft “pelt” from which I had removed the genitalia, but the illustrated sternite for the species I seemed to have was very similar in shape to this object, so I removed it and here’s what I was left with:
With this view, it is clear that what I had just removed was in fact the tegumen (oops), and what I had thought was the tegumen was actually the pair of valvae (which I never did succeed in spreading apart, but it no longer seems necessary). The good news is, it is possible to identify the species from the above photos. Using Davis & Deschka’s key (which, unlike Munroe’s key, refers to helpful figures!), all I have to do is observe that the valvae are symmetrical, that they are longer than the aedoeagus (and gradually taper to a “slender, simple, often sinuate apex”), and that the vinculum is Y-shaped, abruptly tapering to an attenuate apex, and I know I’ve got Phyllonorycter nipigon. The genitalia of my moth match the illustration for that species well. I’m still not clear what those outer two wispy things are, but that horseshoe-shaped structure (the transtilla?) is supposed to be oriented the other way, and it seems like those wispy things may be attached to it in such a way that they would be pointing the other direction if I managed to flip the horseshoe thing to match the illustration, so maybe they are some extraneous structure that was supposed to stay with the rest of the abdomen when I did the dissection. Once again, I welcome input from anyone who knows the answers!
[Edit: Thanks to Aaron Hunt and JoAnne Russo for all the IDs!]
I’ve managed to study insects intensively for over a decade, writing two books and publishing over 50 scientific papers that included the descriptions of 76 new species and one new genus, without ever learning to dissect anything. I have relied on various collaborators to do the dirty work of examining genitalia and other minutia necessary for many identifications and all species descriptions, leaving me free to focus on rearing and documenting natural history. But as boxes of undescribed and undetermined moth species have continued to pile up in my office with no progress toward getting names put on them, I’ve become resigned to the fact that it’s up to me to ensure that all of these wee mothies have not died in vain.
A few years ago Jason Dombroskie was kind enough to give me and Julia a lesson in dissecting micro-moths, but the laundry list of chemicals, other supplies, and equipment required to practice it at home created an inertia that has been hard to overcome, especially when our time is already more than full of things that we don’t need to acquire new skills or materials to do. Last month we finally bit the bullet and got set up to do it, which included studying this tutorial prepared by Sangmi Lee and Richard Brown, and getting additional advice from Terry Harrison and Tony Thomas.
Beginning two weeks ago, we practiced on a few specimens I had collected 15+ years ago for the one entomology class I ever took (a general undergraduate course I took while in grad school at the University of Vermont; the specimens were rendered useless when the professor told the TA to throw out all the students’ notebooks, which contained the collection data, before we had a chance to pick them up at the end of the semester, so nothing was at stake if Julia and I botched the dissections), and that went well enough that two days ago I decided to try out dissecting some undetermined specimens I had reared, and see if I could use the genitalia to identify them.
I started with a relatively large one, because while the method is basically the same for all moths, it gets more difficult the smaller you go, and I don’t want to risk ruining any precious tiny leafminer specimens, which is what I’ve mostly got. Back in late July of 2017, during one of our visits to Nantucket to search for leafminers and other understudied herbivorous insects, Kelly Omand led me and Julia to a little patch of dwarf prairie willow (Salicaceae: Salix occidentalis) to see if there were any unusual bugs on this uncommon plant. We didn’t see any galls or leaf mines, but some kind of moth larvae had tied some of the leaves together in little clumps.
We collected a few of these clumps, and in October this moth emerged:
With a wingspan of around 2 cm, this was many times larger than the moths I know anything about, but I knew it was something in the superfamily Pyraloidea and figured it was probably in the family Crambidae. I posted this photo on good old BugGuide.net, where Aaron Hunt and Kyhl Austin confirmed my suspicion and placed it in the subfamily Pyraustinae, and it has been sitting there ever since. With a moth this nondescript, photos just aren’t going to cut it if you’re looking for a species- or even genus-level ID.
Fortunately, I saved the specimen—which involved relatively humanely gassing it to death in a jar with ammonium carbonate, after which Julia pinned it, spread its wings, and mounted it; and then keeping it safe from marauding booklice, dermestid beetles, and the like for the next several years by periodically refreshing the mothballs in its airtight box. To get a look at the genitalia, I had to carefully remove the abdomen with forceps (some specimens would much rather break in the middle of the thorax, so that the hindwings come off along with the abdomen), then place it in a vial of ~20% potassium hydroxide overnight (Terry advised that first dunking it in 90% alcohol helps it sink in the KOH instead of floating up at the top) to dissolve some of the extraneous tissue. In the morning I moved the abdomen into a tiny puddle of 30% ethanol where I used fine-tipped paintbrushes to remove scales from the surface, then moved it into a tiny puddle of a red stain called “Eosin Y” where I let it sit for a few hours. Next I rinsed it in another tiny puddle of alcohol and then moved it into a tiny puddle of another stain called “Chlorazol Black,” where I left in for another hour or so.
Now (after rinsing the abdomen in another tiny puddle of ethanol) it was time to actually do the dissection, which involves a pair of super fine-tipped foreceps in each hand. Normally with males you’re supposed to keep the “pelt” of the abdomen intact and just remove the genitalia from the tip, whereas with females you carefully tear (or snip, if you have a pair of $300 tiny scissors) the abdomen open along one side, because females have more complicated internal parts to deal with. This specimen had appeared to be a male, because the frenulum consisted of a single bristle, but in my previous practice session I had dissected several pyraloid moths with that same feature and all had turned out to be females. So I didn’t trust that determination and I opened up the “pelt” just to be sure there wasn’t a corpus bursa, and so forth, in there.
There wasn’t, but the tip of the abdomen also didn’t look very much like that of any male moth I had seen previously, so I puzzled over this for a bit until I finally decided to pull off a thin membrane that seemed to be enclosing it, and voila! The valvae magically fell open and it was most definitely a male. I don’t yet have a good setup for taking photos through a microscope (which will require, among other things, a better microscope), but here’s what it looked like:
The thing poking out at lower right is what most lepidopterists call the “aedeagus,” but Jason and Kyhl say that’s technically wrong (I forget why) and use “phallus” instead, which certainly makes it clearer to non-entomologists what we’re talking about. The phallus sort of gets in the way of things and you’re generally supposed to remove it, so I did, and then I moved both parts into a drop of lactic acid I’d placed on a microscope slide, and then flattened it all out with a coverslip, which always results in a few bubbles, but trying to fix those only creates lots more bubbles, so I left them as they were:
So now what? Now it was time to consult the compendium of North American pyraustine moths and figure out who this moth was. Fortunately, such a compendium exists (it does not exist for Gracillariidae, the largest family of leaf-mining moths, but we’ll come to that problem later), and the PDF can be downloaded for free here if you want to play along: Munroe (1976), the one labeled “Fascicle 13.2A: Pyralidae, Pyraustinae (part 1).”
Turn to page 8 and we find that what is now Crambidae: subfamily Pyraustinae was treated as Pyralidae: Pyraustinae: tribe Pyraustini in 1976, and on the next page we find that the key to genera of Pyraustini is based entirely on genitalia. After a few couplets it is based entirely on male genitalia, so it’s a good thing that’s what we’ve got! Soon flustered by all the unfamiliar terminology, and irritated by the complete lack of references to any figures, we take to flipping through all the photos of genitalia in the back of the book, and we find that this clearly isn’t any of the species illustrated. So then we return to the key and start trying to figure out what all the terms mean, one by one.
I labeled the photo with my interpretations of all of the terms that came up in the key (based on various online sources, including the glossary at Pacific Northwest Moths and several highly pixilated thumbnail images from https://britishlepidoptera.weebly.com/male-genitalia.html that came up in a Google image search—I can’t see the full versions because my malware-blocking software won’t let me visit that website), and then added five more (lower right) for completeness. I welcome corrections if any lepidopterists are reading this!
The genus I landed on was Loxostege, and I didn’t try the species key because a quick look at photos of Loxostege adults suggested that this isn’t the right genus. And that’s where I’ll leave this story for now, because I don’t know where to go from here without some input from someone who knows more about this than I do.
Hey, this blog now has over 1000 subscribers! Thanks everyone for your continued interest in my esoteric natural history investigations.
I’m still slowly working my way through the photos I took last summer, during which one of my several jobs involved exploring ridges and summits in the southwestern corner of Massachusetts. On August 9 I visited Jug End in Egremont, where the Appalachian Trail passes through a very nice example of a ridgetop pitch pine – scrub oak woodland, which is a rare thing in Massachusetts.
One of the nice things about scrub oak (Fagaceae: Quercus ilicifolia) is that all the acorns are down low, providing opportunities to see “plum galls” of Amphibolips quercusjuglans (Cynipidae) while they’re still attached. I normally only see these galls on the forest floor, after they’ve dropped from canopy red or black oaks, and I’d been noticing them for several years before I learned that they grow out of the sides of acorn caps. Here are some of the ones I saw at Jug End that day:
The acorn to the left has a single gall, the one at lower right has two, and the one in the background has three full-sized ones plus a fourth underdeveloped one—quite a load for one little acorn to carry. But back to the lower right, notice that there are a couple of bugs sitting on one of the galls. These are an ambush bug (Reduviidae: Phymata) eating a crabronid wasp that I’m told is in the genus Crossocerus. When I saw this I wished I’d lugged a better camera with me, but here’s the best I could do with the little point-and-shoot that fits in my pocket:
If you cut one of these “plum galls” open, you’ll find that it consists mostly of apple-like flesh, with a hard, spherical cell in the center where a single wasp larva is developing.
According to Weld (1959), “Tranformation takes place in the fall; emergence in the spring Feb. 17 – May 14. Mo. and is distributed over more than one season.” I think he meant by this that the larva pupates and becomes an adult in the fall, but overwinters before chewing its way out of the gall—and the “Mo.” means that those were the emergence dates recorded in Missouri. MJ Hatfield reported here that she collected galls in Iowa in September 2008; nothing had emerged by June 1, 2010, so she cut one open and found a live larva inside, prompting her to save the remaining galls, and an adult emerged from one of them in April 2011.
I collected some oak plum galls from the forest floor near Amethyst Brook in Amherst, MA in August 2011, and nothing emerged the following spring, but some time in the fall of 2012 (while Julia and I were traveling around the western US for two and a half months in search of leafminers), two cynipid wasps emerged and died. They were not Amphibolips quercusjuglans, though; they were inquilines (developing in galls induced by A. quercusjuglans), and Matt Buffington identified them as belonging to the genus Ceroptres. One was a female…
…and one a male.
The container with the galls also had a tiny, shriveled moth larva in it:
I think it must have been the same larva that I photographed on September 1, 2011, when it had just emerged from one of the galls:
I’m not sure what it would have needed to complete its development, but I guess I just left it in the container with the galls and hoped for the best. That dried larva is now at the Smithsonian along with the two wasps, so theoretically it could still be identified using DNA barcoding.
On September 21, 2019, I was leading a walk at the Fitzgerald Lake Conservation Area in Northampton, MA, when a little girl presented me with an acorn plum gall she had just found and demanded that I cut it open. I did as I was told, and somehow managed to cut right through the central cell without killing the larva inside. The larva, I was surprised to see, was not that of a gall wasp—it was another caterpillar! I have no photos of it, because I put the gall back together and kept it that way to let the larva finish feeding without being disturbed further. However, I just found this photo on BugGuide, taken by Tom Murray in Concord, MA on October 23, 2011, that shows the same thing:
It appears to be the same type of larva that I’d had emerge from a gall collected in Amherst just a few weeks earlier.
As for the more recent gall from Northampton, nothing emerged until the following May—and instead of the adult moth I was hoping for, it was an ichneumon wasp, which had developed as a larva feeding on the moth larva inside the wasp gall.
So the question remains: Who is this caterpillar that feeds inside of acorn plum galls, and does it do so exclusively? The fact that it lives inside the cell where the gall wasp is supposed to be suggests more than a casual association (and also suggests that it may eat the wasp larva). It would also be interesting to know whether this ichneumonid exclusively parasitizes this moth species that feeds inside of cynipid wasp galls, but based on my understanding of ichneumonids I think it more likely parasitizes a variety of moth larvae that feed in concealed situations (fruit and stem borers, leaftiers, etc.).
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