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.).
On May 8 last year, Julia and I visited her family’s land in Hocking County, Ohio (which we’ll be doing again today, as it happens), and for whatever reason, a little clump of dead wingstem (Asteraceae: Verbesina alternifolia) stems from the previous season caught my eye. I’m always amazed by all the things John van der Linden is able to find living inside of plant stems when there is little or no external evidence, and occasionally I’m moved to split a stem open at random and see what there is to see. So I got out my trusty Swedish army knife, started splitting one of these wingstem stems open, and behold! Something had been tunneling in the pith.
A little more splitting turned up what I was hoping to find: puparia of a Melanagromyza species; one of the so-called “leafminer flies” (Agromyzidae) that is a stem borer instead of a leafminer.
But the fly larvae hadn’t been the only things tunneling in there. As Julia and I split open additional stems, we also found several larvae and pupae that I thought might be Mordellidae: “tumbling flower beetles.”
Naturally, we collected the chunks of stem containing these immature insects in vials to see what they would turn into. The first thing to emerge, on May 18, was this gall midge (Cecidomyiidae):
This was not unexpected; the second Melanagromyza puparium shown above was at the edge of a slight swelling in the stem that I had thought might be a midge gall. Ray Gagné has confirmed that this midge is Neolasioptera imprimata, a species he had described just two years earlier. Here’s a better look at the gall:
Just to the right of the center of the above photo, the midge’s pupal skin can be seen poking out of the gall. A closer view:
Beginning two days later, 22 platygastrid wasps emerged from the same gall—and I only collected half of the gall, so there were probably even more midges and wasps in the other half. Jessica Awad tells me these wasps belong to the genus Platygaster, and that a species ID is unlikely to be possible anytime soon given the current state of knowledge of this genus. All platygastrid wasps (as the family is currently circumscribed) are parasitoids of gall midges.
Also on May 20, an adult mordellid beetle appeared, proving my hunch right. As far as I can tell, there is no one who studies these beetles or knows how to identify them, but this one is now in the Canadian National Collection if anyone wants to have a look at it.
On June 13, two more wasps emerged. One was a braconid parasitoid of the mordellid beetles, which Gideon Pisanty identified (based on photos I posted on iNaturalist) as a member of the tribe Brachistini (Brachistinae).
The other was this figitid wasp, which I recently sent to Matt Buffington along with 50 or so others I’ve accumulated over the past few years, and he hasn’t had a chance to look at it yet. It emerged from one of the Melanagromyza puparia.
Something dark developed inside the other Melanagromyza puparium, but nothing ever emerged from it.
According to Spencer & Steyskal (1986), the only stem-boring Melanagromyza known from wingstem (or any other Verbesina species) is M. verbesinae, which Owen Lonsdale synonymized last year with M. vernoniana—a species that, based on the specimens Owen examined, feeds on other Asteraceae including ironweed (Vernonia), sneezeweed (Helenium), sunflower, and Jerusalem artichoke (Helianthus). Spencer & Steyskal described the puparium of M. verbesinae (the holotype of which was collected in Ohio) as “straw colored, posterior spiracular plates heavily chitinized, closely adjoining, only narrowly divided, each with ellipse of about 12 bulbs around strong central horn.” This seems to fit what I found, so I’ll choose to believe that the flies I failed to rear were M. vernoniana and not something new and exciting that I need to try to find again today.
For more on the fascinating world of stem-dwelling insects, about which I know very little, check out this document John van der Linden recently put together summarizing all the agromyzid flies he’s found in this microhabitat. And you can learn about the other flies, moths, beetles, etc. he’s found living in stems by perusing his photos on BugGuide.
Someone asked me the other day how many new species I’ve found, and I realized that in addition to not being able to give a straight answer for the reasons discussed below, I’d lost track of how many species I’ve helped to describe and name, which is a question that at least has a definite answer. So here’s an update of my original “How Many New Species?” post from three years ago. It’s been fun to see people finding some of these species all over the US and Canada as I peruse the leafminer observations on iNaturalist.
I am often asked how many new insect species I have found (or “discovered”). I’m never quite sure how to answer this. I’ve certainly reared dozens of undescribed species of moths, for instance, that are now sitting in my office or in various museums, waiting to be named. The number for parasitoid wasps is probably even higher. But simply having “found” new species doesn’t count for much if they haven’t been properly documented and named. Also, to me the credit for “discovering” a new species mostly belongs to the taxonomist who does the hard work of comparing it with all the similar species in the world to demonstrate that it is really something new. Of all the species I have coauthored, Marmara viburnella is the only one I felt certain was undescribed (because I went to the trouble of reviewing the larval biology and adult morphology of all the previously described species in that genus) before I passed it along to a taxonomist.
With that said, for my final post of the year, I thought it would be fun to put together a list of all the species that have been either described in papers I coauthored or described based (at least in part) on specimens I collected. (This was partly inspired by my realization that I never got around to writing anything here about most of the 30 fly species I recently described with Owen Lonsdale.) With any luck, this list will continue to grow. What limits the number of new species I’m able to help describe is a shortage not of “new” species to name, but of time that my collaborators and I have available to devote to this task. So the take-home message from this post should not be “Wow, look at all the new species Charley has found!”, but rather, “Wow, we have so much left to learn about our natural surroundings, and we need to support more funding for taxonomy!”
For species I’ve written about before, you can click on the name to see the relevant post.
First, the species I did not coauthor (of these, Orchestomerus eisemani and Adelius floridensis are the only species for which one of my specimens was designated as the holotype):
1. Celticecis cornuataGagné, 2013 – A hackberry gall midge I found in Kentucky while traveling with Noah to check out the periodical cicadas in Nashville and Sam Droege’s bees in Maryland.
2. Orchestomerus eisemani Yoshitake & Anderson, 2015 – A leafminer of Virginia creeper I found at work one day in Plymouth County, Massachusetts. This seems to be pretty close to its northeastern range limit; if you check the map on iNaturalist you can see that I ‘ve since found it as far north and west as Concord, but no sign of it yet anywhere in western Massachusetts or more northern states.
3. Brachys howdeni Hespenheide (in Hespenheide & Eiseman, 2016) – I first found this trailing arbutus leafminer while hiking along the ridge just above the house where I now live. I see the mines in just about every sizable patch of the host plant I encounter.
4. Liriomyza limopsis Lonsdale, 2017 – Owen had already given this species a name based on Canadian specimens collected as adults, but no host plant was known until I reared it from white wood aster (Eurybia divaricata) and whorled aster (Oclemena acuminata) in New York and Massachusetts.
5. Liriomyza pilicornis Lonsdale, 2017 – Similar story, except that Graham Griffiths was the first to rear this species, 45 years before Julia and I found it mining leaves of bastard toadflax (Comandra umbellata) in Massachusetts.
6. Liriomyza pistillaLonsdale, 2017 – Ditto, except in this case the host is cow-wheat (Orobanchaceae: Melampyrum lineare) and Griffiths reared it 40 years before me. I find the leaf mines pretty regularly.
7. Adelius floridensis Shimbori & Shaw, 2019 – This braconid wasp species is known only from a few specimens I reared in 2013 from the St. Johnswort leafminer Fomoria hypericella (Nepticulidae) in Florida.
So that’s seven in the first category. I mostly just happened to give specimens to the right taxonomists at the right time: Ray Gagné was finishing up a revision of all the gall midge species on hackberries; Henry Hespenheide was (and is) in the midst of revising the genus Brachys, and Owen Lonsdale was finishing up a revision of the Canadian species of Liriomyza. In the case of Orchestomerus eisemani, Bob Anderson was inspired to revise that genus after having initially identified the weevils I had reared as O. wickhami Dietz, then discovered his error after I had published a note documenting their natural history. Adelius floridensis got to be described because Dave Wagner directed Scott Shaw and Eduardo Shimbori to me when they asked him if he had any specimens they might use in their revision of the New World braconid wasps of the tribe Adeliini. For the 76 species listed below that I have coauthored, I’m extremely grateful to the taxonomists who took time away from whatever other projects they were working on to help me put names to my natural history observations.
1. Scolioneura vaccinii Smith & Eiseman (in Smith et al. 2015) – A sawfly that mines leaves of huckleberries (Vaccinium spp.), which Julia and I found in western Washington on our first cross-country trip in search of leafminers (though we only were able to rear parasitoids, and the type specimen was reared from a larva Noah and his wife Sydne collected the following year).
2. Megaselia nantucketensis Eiseman & Hartop, 2015 – A scuttle fly that emerged from a midge gall on black oak, collected on Nantucket during the gall and leaf mine survey I’ve been conducting there since 2011.
3. Megaselia risoria Hartop, Wong & Eiseman, 2016 – The naming of this species was a byproduct of my having reared specimens of M. globipyga from a dead tussock moth caterpillar I found at work.
4. Platygaster pruni Buhl & Eiseman, 2016 – A platygastrid wasp that emerged from a midge gall on black cherry, which I collected at work one day in western Massachusetts.
5. Platygaster uvulariaeBuhl & Eiseman, 2016 – A platygastrid wasp that emerged from a midge gall on wild oats (Uvularia sessilifolia)—again collected at work in western MA. No one has yet been able to rear the midge that causes this gall.
6. Platygaster vitisiellae Buhl & Eiseman, 2016 – A platygastrid wasp that emerged from a midge gall on wild grape, collected as part of the Nantucket survey. The midge species is probably undescribed (but I was able to rear some adults, which are sitting in the Smithsonian waiting for someone to decide to revise the genus Vitisiella).
7. Zygoneura calthella Eiseman, Heller & Rulik, 2016 – A dark-winged fungus gnat that feeds in leaves and petioles of marsh marigold. Julia and I first found it while surveying for four-toed salamanders in western Massachusetts.
8. Fenusa julia Smith & Eiseman, 2017 – A sawfly that mines leaves of wild rose, which Julia spotted in Colorado on another leafminer-hunting road trip.
9. Marmara viburnella Eiseman & Davis (in Eiseman et al. 2017) – Another product of the Nantucket survey. The larva of this moth begins as a leafminer, then disappears down the petiole and spends most of its life feeding in the stem. Julia and I reared it from arrowwood, but mines have also been found on various other viburnums.
10. Platygaster tephrosiae Buhl & Eiseman, 2017 – Another one from Nantucket; I reared the two known specimens from midge galls that happened to be on some goat’s rue leaves that Kelly Omand collected for me to feed some leaf-tying caterpillars. I failed to rear the caterpillars, and I haven’t been able to rear the midge yet either.
11. Platygaster vacciniiBuhl & Eiseman, 2017 – The single known specimen emerged from a gall on lowbush blueberry that I collected at the 2016 Berkshire BioBlitz on Mt. Greylock—caused by another midge that has never been reared.
12. Macrosaccus coursetiae Eiseman & Davis, 2017 – Another one Julia and I collected on our first cross-country trip; this one from Arizona, mining leaves of a shrub called rosary babybonnets (Coursetia glandulosa).
13. Phytosciara greylockensis Eiseman, Heller & Rulik, 2018 – Another one from the Mt. Greylock BioBlitz; a dark-winged fungus gnat that mines leaves of bluebead lily (Clintonia borealis).
14. Agromyza fission Eiseman & Lonsdale, 2018 – Owen had already named this species based on a specimen collected in the DC area in 1914, but the type specimen is one Julia and I collected at MJ Hatfield’s “Red Oak Prairie” in eastern Iowa on the way home from Colorado. One of the paratypes came from a larva we collected the next day on Marcie and Mike O’Connor’s land in Wisconsin, and Mike Palmer provided two from Oklahoma. The larvae mine leaves of hackberry.
15. Agromyza sokaEiseman & Lonsdale, 2018 – This is another one that Owen had already named based on a 1914 specimen from the DC area, but as with A. fission its host was unknown. It turns out to be responsible for leaf mines on black locust that since 1982 have been attributed to Phytoliriomyza robiniae (Valley), adults of which were repeatedly associated with black locust but never actually reared. Some paratypes came from specimens Julia and I reared from larvae we collected at the 2016 Connecticut BioBlitz, and the rest came from larvae Tracy Feldman found mining both black locust and wisteria in North Carolina.
16. Melanagromyza palmeri Eiseman & Lonsdale, 2018 – The only known specimen is one that Mike Palmer reared from a sunflower stem (or possibly the roots) in Oklahoma. Stem feeding members of this genus are borers rather than miners, meaning that they don’t form any externally visible trails. So rearing them takes special dedication and/or luck.
17. Ophiomyia euthamiaeEiseman & Lonsdale, 2018 – This species mines leaves of grass-leaved goldenrod (Euthamia graminifolia), mostly on the lower surface. I first noticed mines like this on Nantucket, but those were possibly made by O. maura; the whole type series of O. euthamiae came from my yard.
18. Ophiomyia mimuliEiseman & Lonsdale, 2018 – This species mines in stems of monkeyflower. I first found it at a bioblitz on Julia’s family’s land in southern Ohio, and some paratypes came from the swampy woods right behind our house in Massachusetts.
19. Ophiomyia pardaEiseman & Lonsdale, 2018 – Another species whose holotype I collected in my yard. It is a common leafminer of asters (Symphyotrichum spp.) and seems to be responsible for all of the mines previously attributed to O. quinta.
20. Calycomyza artemisivoraEiseman & Lonsdale, 2018 – This species is known only from two specimens I reared from leaf mines on Artemisia ludoviciana that Mike Palmer collected in Oklahoma.
21. Calycomyza aviraEiseman & Lonsdale, 2018 – Another one that Owen had already named before I reared it; there are several specimens at the Smithsonian from Connecticut, New York, and West Virginia, dating back to 1929. The larvae mine leaves of beggar-ticks (Bidens spp.). I reared some from mines I collected at work, and Tracy Feldman provided some from North Carolina.
22. Calycomyza eupatoriphagaEiseman & Lonsdale, 2018 – This belongs to the same species complex as C. artemisivora. It has been reared from three plants in the tribe Eupatorieae: I found it on white snakeroot (Ageratina altissima) in Massachusetts and on blue mistflower (Conoclinium coelestinum) in Tennessee, and Mike Palmer found it on late boneset (Eupatorium serotinum) in Oklahoma. In teasing apart the members of this complex, Owen found a specimen that was collected in Ontario in 1947, which he included as a paratype. The holotype is from the woods right behind our house.
23. Calycomyza vogelmanniEiseman & Lonsdale, 2018 – I reared the only known specimen from a leaf mine on thin-leaved sunflower (Helianthus decapetalus), which I collected near Burlington, Vermont, where I went to grad school. I named the species after Hub Vogelmann, who founded my graduate program (the Field Naturalist program). He had retired long before I attended, but he was very enthusiastic about my first book when it was published, and he was supportive as I got started on my leafminer book project.
24. Cerodontha edithae Eiseman & Lonsdale, 2018 – This species is an iris leafminer, the only known specimen of which Julia and I reared as part of our Nantucket surveys. I named it after Edith Andrews, who died in 2015, a day after her 100th birthday. She lived on Nantucket for most of her life and was an enthusiastic naturalist to the end. Birds were her main passion, but not long after Julia gave her a copy of my book, Julia went to visit her and found her and her daughter Ginger on their hands and knees in her driveway, getting a closer look at some wasp burrows. When I first met her, I was amazed at the memory of this nearly 100-year-old woman as she quoted from various parts of my book.
25. Cerodontha feldmaniEiseman & Lonsdale, 2018 – Another species known from a single specimen; in this case one I reared from a sedge leaf mine that Tracy Feldman collected in North Carolina. Tracy has been intensively collecting leafminers in North Carolina and elsewhere for the past few years and has found numerous new state records, new host records, and new species.
26. Liriomyza ivorcutleri Eiseman & Lonsdale, 2018 – I reared the holotype from a leaf mine on cup plant (Silphium perfoliatum) that Julia and I found in Iowa. When Owen told me this yellow fly was a new species, I couldn’t resist naming it after Ivor Cutler, the Scottish recording artist responsible for “Yellow Fly,” along with other classics like “I Believe in Bugs.”
27. Liriomyza valerianivoraEiseman & Lonsdale, 2018 – I found the leaf mines of this species in a scrappy wetland in north-central Massachusetts where I was conducting botanical fieldwork with Sally Shaw. I was lucky she was with me, because I never would have recognized the basal leaves of garden valerian, which are totally different from those on mature plants.
28. Phytomyza actaeivoraEiseman & Lonsdale, 2018 – I tried for several years to rear adults from leaf mines on red baneberry (which I’ve found in Vermont and Ohio) before finally succeeding with some mines I found on white baneberry in my neighbors’ woods. I also found mines of what is probably the same species on black cohosh (these are all Actaea species) at Jason Dombroskie’s house in Ithaca, NY, but these were all parasitized like the ones on red baneberry.
29. Phytomyza aesculiEiseman & Lonsdale, 2018 – I first became aware of this species because of photos of buckeye leaf mines that several different people posted to BugGuide.net. One spring when Julia was visiting her parents in Ohio (the Buckeye State), she managed to collect a bunch of larvae, from which I reared the type series. This species is active only in spring, with a pupal diapause lasting nearly a year. The author of this article was grateful when I let him know that his mystery “buckeye leafmining fly” now has a name.
30. Phytomyza confusaEiseman & Lonsdale, 2018 – I named this fly “confusa” because everything about it was confusing. I found the leaf mines at the base of a tree in the middle of a lawn in a big park in Iowa. I tentatively identified the plant as Virginia waterleaf (Hydrophyllum virginianum), but it looked a little weird to me (not to mention that Virginia waterleaf is normally a forest species). Iowa botanist John Pearson suggested that it might be a buttercup such as Ranunculus fascicularis. When Owen initially determined the flies as belonging to a new species in the Phyomyza aquilegiae group, this seemed to fit, since all members of this group feed on plants in the buttercup family as far as is known. I showed my photo of the plant to several other botanists, and they all shared my initial impression that it was Virginia waterleaf, but most were also willing to believe it was Ranunculus fascicularis, and one even examined some herbarium specimens of that species that she said matched in every respect. But Owen later determined P. confusa to be closely related to another new species that I reared from two species of waterleaf (see below), and decided both flies probably are better placed in the P. obscura group, all species of which feed on plants in the mint and borage families (waterleafs are in the latter). This species was also confusing because the leaf mines were hard to characterize—some began with a distinct linear portion and some did not, and by the time the adults emerged the leaves were so crumpled and degraded that I couldn’t decide whether the puparia were formed inside or outside the mines. John van der Linden has since repeated the rearing of this species from Virginia waterleaf, so so the identity of its host plant is no longer a point of confusion, but other details of its biology are still unclear.
31. Phytomyza doellingeriaeEiseman & Lonsdale, 2018 – While working in Maine in July 2013, I collected leafminers from flat-topped aster (Doellingeria umbellata) that Owen determined to be a new species near P. solidaginivora Spencer based on the genitalia. Both of the adults I reared were underdeveloped (as shown here), so when I returned in August I collected some more. I reared some good specimens this time, but Owen determined them to be a different new species, which I named P. doellingeriae. Meanwhile, he decided the first flies were close enough to P. solidaginivora to call them that for now. Incidentally, Spencer (1969) reared P. solidaginivora from a plant in Alberta that he thought was goldenrod (Solidago; hence the name), but Graham Griffiths examined his pressed leaf mines and didn’t think they looked like any goldenrod that occurs in Alberta. Spencer’s drawing of the leaf looks exactly like a flat-topped aster leaf, so that fly is probably misnamed.
33. Phytomyza hatfieldaeEiseman & Lonsdale, 2018 – When Julia and I stayed with MJ Hatfield in northeastern Iowa on the way home from Colorado, we spent a little time exploring the woods on the bluff next to her house with MJ and John van der Linden. Leaf mines that we collected there on sweet cicely (Osmorhiza claytonii) yielded the holotype of this new species. The paratypes also included a number of specimens Graham Griffiths had reared from various Osmorhiza species in the 1970s, plus a few that Ed Stansbury reared in Washington just in time to be included in the paper.
34. Phytomyza hydrophyllivoraEiseman & Lonsdale, 2018 – This species is common on broad-leaved waterleaf (Hydrophyllum canadense) in Ohio, and I collected the mines several times from the woods by Julia’s parents’ house before I finally got a few adult flies instead of parasitoid wasps. I later reared one from the same host in Tennessee (during our brief trip to see the solar eclipse in 2017), and one from a mine I found on Virginia waterleaf while conducting a rare plant survey in the Berkshires.
35. Phytomyza palmeriEiseman & Lonsdale, 2018 – This is another species (like Melanagromyza palmeri) known only from Mike Palmer‘s yard in Oklahoma, and although he gave me a number of leaf mines, only he has been able to rear adults. The larvae mine leaves of coralberry (Symphoricarpos orbiculatus).
36. Phytomyza palustrisEiseman & Lonsdale, 2018 – I found this leafminer of swamp saxifrage while conducting botanical fieldwork in the Berkshires. I check this plant for mines every time I see it, but as far as I can tell the range of this species is limited to one square meter in the town of Washington, MA.
37. Phytomyza sempervirentisEiseman & Lonsdale, 2018 – Julia and I first found this species when we visited Cane Creek Canyon in northern Alabama on our way home from Florida in spring 2013. The larvae form mines on coral honeysuckle (Lonicera sempervirens) very similar to those formed by the closely related P. nigrilineata (Griffiths) on limber honeysuckle (L. dioica) in Alberta. I found more (including the holotype) three years later at the Montague Plains in western Massachusetts. Tracy Feldman also provided a bunch of specimens from North Carolina, and Mike Palmer reared a few from orange honeysuckle (L. ciliosa) in Oregon.
38. Phytomyza tarnwoodensisEiseman & Lonsdale, 2018 – I reared the only known specimens of this species from leaf mines on bush honeysuckle (Diervilla lonicera) I collected in my parents’ yard in western MA. “Tarnwood” is the name my parents gave to their property many years ago, and when I was little this sign that my mother painted used to be on a post at the edge of our yard by the road:
39. Phytomyza tigrisEiseman & Lonsdale, 2018 – The larvae of this species mine leaves of foamflower (Tiarella cordifolia). The leaf mines are very common, but it took me many tries (always getting only parasitoid wasps) until I finally managed to rear a few adults—in my neighbors’ woods right near where I finally found unparasitized puparia of P. actaeivora. The name Phytomyza tiarellae was already taken, so I named this one “tigris” after the tiger stripes on its puparium (going with the “big cat” theme Owen had started with Ophiomyia parda).
40. Phytomyza triangularidisEiseman & Lonsdale, 2018 – This is another one Julia and I found on our first cross-country trip, this time in northern Idaho. The larvae mine leaves of arrowleaf ragwort (Senecio triangularis).
41. Phytomyza vancouveriellaEiseman & Lonsdale, 2018 – Although Julia and I found a few leaf mines of this species on the Olympic Peninsula on that same trip, the only known specimens are a few that Mike Palmer reared in Oregon five years later. The host is Vancouveria hexandra, whose common names include “white inside-out flower.”
42. Phytomyza verbenaeEiseman & Lonsdale, 2018 – One last species (for now) from that first road trip; Julia and I found the mines on western vervain (Verbena lasiostachys) in California.
43. Phytomyza ziziae Eiseman & Lonsdale, 2018 – I reared the holotype and some of the paratypes from leaf mines on golden Alexanders (Zizia aurea) I collected while conducting botanical fieldwork in western Massachusetts. Another came from the same Berkshire BioBlitz that produced the type specimens of Phytosciara greylockensis and Platygaster vaccinii. There are also a few specimens that Graham Griffiths reared from heart-leaved golden Alexanders (Zizia aptera) in Alberta in 1973. It’s a bit curious how many agromyzid species are known only from Alberta and Massachusetts…
44. Agromyza arundinariae Eiseman, Lonsdale & Feldman, 2019 – This and the next eight species were described in a paper dedicated to agromyzid flies that Tracy Feldman collected in North Carolina—I helped with rearing and described the leaf mines, and Owen Lonsdale described the adult flies. Agromyza arundinariae is one of three new species Tracy found on the native bamboo Arundinaria tecta.
45. Agromyza indistinctaEiseman, Lonsdale & Feldman, 2019 – I gave this one the name “indistincta” because there was nothing distinctive about it; it’s the fourth species to be reared from seemingly identical mines on grasses in the genus Dichanthelium, and the adult is pretty darn similar to two of the other three.
46. Calycomyza chrysopsidis Eiseman, Lonsdale & Feldman, 2019 – A leafminer of Maryland goldenaster (Asteraceae: Chrysopsis mariana).
47. Cerodontha enigmaEiseman, Lonsdale & Feldman, 2019 – This one is a similar situation to Agromyza distincta; it is a leafminer of Dichanthelium and is an enigma because it is known from a single specimen Tracy collected in his yard. The adult is very similar to Cerodontha angulata, which is the species I’ve reared every time I’ve collected similar mines on Dichanthelium.
48. Cerodontha arundinariellaEiseman, Lonsdale & Feldman, 2019 – The second species reared from leaf mines on Arundinaria tecta.
49. Cerodontha saintandrewsensis Eiseman, Lonsdale & Feldman, 2019 – The third species reared from leaf mines on Arundinaria tecta, named for St. Andrews University, where Tracy works and does much of his collecting (and the only known locality for this species).
50. Liriomyza carphephoriEiseman, Lonsdale & Feldman, 2019 – Tracy first found this species mining leaves of sandywoods chaffhead (Asteraceae: Carphephorus bellidifolius), and I had already decided to name at after this plant when Owen determined that it was the same species Tracy and I had reared from beggarticks (Bidens spp.) in North Carolina, Vermont, and my own front yard in Massachusetts. It’s still a good name for it; several other Liriomyza species mine leaves of Bidens, but this is so far the only leafminer of any kind to be found on Carphephorus.
51. Liriomyza polygalivoraEiseman, Lonsdale & Feldman, 2019 – A leafminer of whorled milkwort (Polygalaceae: Polygala verticillata).
52. Liriomyza triodanidisEiseman, Lonsdale & Feldman, 2019 – A leaf and stem miner of small Venus’ looking-glass (Campanulaceae: Triodanis biflora).
53. Agromyza princei Eiseman & Lonsdale, 2019 – I reared this species from a leaf mine on black raspberry (Rosaceae: Rubus occidentalis) that Julia and I collected in the parking lot of a cemetery in Hartford at the 2016 Connecticut BioBlitz. It is known from a single specimen, which emerged as an adult a year after I collected the larva. When Owen told me he needed a name for this new species, “Raspberry Beret” popped into my head, so I named it after Prince, who had died shortly before we found the leaf mine.
54. Melanagromyza vanderlindeniEiseman & Lonsdale, 2019 – This species is named for John van der Linden, who reared it from dead stems of Joe-Pye weed (Asteraceae: Eutrochium) he collected in Iowa. John has an incredible knack for finding stem-feeding insects that leave little or no external evidence. He has written about some of his natural history discoveries on his blog, and many more can be found on his BugGuide page.
55. Ophiomyia antennariaeEiseman & Lonsdale, 2019 – Julia and I found this leafminer of plantain-leaved pussytoes (Asteraceae: Antennaria plantaginifolia) in beautiful Cane Creek Canyon in northern Alabama in the spring of 2013, shortly after getting in a car wreck in Mobile. The mines are much like those from which Mike Palmer and I have reared O. coniceps in Oklahoma and New England, and Owen and I almost described this new species in our first (2018) paper, but I temporarily convinced him that we should consider the Alabama specimens to be O. coniceps. However, before that paper went to press, Owen reasserted his original position—though he considers one female from Cane Creek Canyon to be a better match for O. coniceps—so we removed the remaining specimens from that paper and dealt with them in the 2019 paper. As it happens, Julia and I had separated out what we thought might be two different mine types on plantain-leaved pussytoes, and that female was the only one that emerged from mines of the type that O. coniceps makes in New England. But Mike has reared O. coniceps from both mine types in Oklahoma, so evidently the difference isn’t entirely consistent.
56. Ophiomyia osmorhizaeEiseman & Lonsdale, 2019 – Another John van der Linden discovery from Iowa; this one is a stem miner of sweet cicely (Apiaceae: Osmorhiza).
57. Calycomyza smallanthiEiseman & Lonsdale, 2019 – In August 2017, Julia and I met up with Noah’s family in Nashville to see the full solar eclipse, and we found this species mining leaves of hairy leafcup (Asteraceae: Smallanthus uvedalius) just around the corner from Noah’s mother’s house.
58. Liriomyza euphorbiellaEiseman & Lonsdale, 2019 – Mike Palmer reared this one from fire-on-the-mountain (Euphorbiaceae: Euphorbia cyathophora) in Oklahoma.
59. Liriomyza garryaeEiseman & Lonsdale, 2019 – Julia and I found this species mining leaves of silktassel (Garryaceae: Garrya spp.) in Arizona and Texas on the way home from checking out the “super bloom” in southern California in 2017.
60. Liriomyza phloxiphagaEiseman & Lonsdale, 2019 – I reared the only known specimen of this species in 2017 from a leaf mine on phlox (Polemoniaceae: Phlox paniculata) in my mother’s garden in Massachusetts.
61. Phytomyza nemophilaeEiseman & Lonsdale, 2019 – Mike Palmer reared this species from leaf mines on Nemophila parviflora (Hydrophyllaceae) in Oregon.
62. Phytomyza salviarumEiseman & Lonsdale, 2019 – Julia and I found this species mining leaves of several different sage species (Lamiaceae: Salvia) on Ann and Bruce Hendrickson’s ranch in Texas in 2017, the same day we collected the holotype of Liriomyza garryae.
63. Grapholita thermopsidisEiseman & Austin (in Eiseman et al. 2020) – A leafminer of goldenbanner (Thermopsis) reared from leaf mines Julia and I collected in Colorado in the yard of our friends Sally Waterhouse and Denny Radabaugh in 2015.
64. Melanagromyza arnoglossi Eiseman & Lonsdale, 2021 – This species, and the twelve that follow, were described in a paper that included John van der Linden, Tracy Feldman, and Mike Palmer as coauthors. Most of the species were found and reared by John, Tracy, or Mike, so I only have my own photos of a few of them. When available, I’ve made the species names link to relevant BugGuide posts. Melanagromyza arnoglossi is a stem borer John reared from Indian plaintain (Arnoglossum) in Iowa.
65. Melanagromyza gentianivora Eiseman & Lonsdale, 2021 – A stem borer John reared from bottle gentian (Gentiana andrewsii) in Iowa; the type series also includes specimens reared by Andrew Williams in Wisconsin.
66. Melanagromyza hieraciiEiseman & Lonsdale, 2021 – A stem borer John reared from rough hawkweed (Hieracium scabrum) in Iowa.
67. Melanagromyza rudbeckiae Eiseman & Lonsdale, 2021 – A stem borer John reared from cutleaf coneflower (Rudbeckia laciniata) in Iowa; the type series also includes specimens reared by Andrew Williams in Wisconsin.
68. Melanagromyza urticae Eiseman & Lonsdale, 2021 – A stem borer John reared from stinging nettle (Urtica dioica) in Iowa.
69. Melanagromyza verbenivora Eiseman & Lonsdale, 2021 – A stem and rachis borer John reared from hoary vervain (Verbena stricta) in Iowa.
70. Ophiomyia nabali Eiseman & Lonsdale, 2021 – A stem miner and petiole borer that John reared from white rattlesnake-root (Nabalus albus) in Iowa.
71. Ophiomyia rugula Eiseman & Lonsdale, 2021 – This is one Tracy reared in North Carolina from short, gall-like leaf mines on groundsel bush (Baccharis halimifolia). The mine looks like a little wrinkle along the midrib, hence the name “rugula,” which is Latin for “a small wrinkle.”
72. Haplopeodes loprestii Eiseman & Lonsdale, 2021 – I reared the two known specimens from a sprig of California fagonbush (Fagonia laevis) that Eric LoPresti gave me. As explained in the linked post, Eric collected the plant sample because it had some leaf-mining moth larvae, and we never actually saw what the fly larvae did but they were presumably also leafminers.
73. Liriomyza euphorbivora Eiseman & Lonsdale, 2021 – A leafminer Mike reared from snow-on-the-mountain (Euphorbia marginata) in Oklahoma.
74. Liriomyza hypopolymnia Eiseman & Lonsdale, 2021 – This is a sneaky one that mines on the lower surface of leafcup (Polymnia canadensis) leaves. There is little or no trace of the mine when viewed from above, and it’s pretty inconspicuous from below too. Julia and I first found the mines in Iowa by MJ Hatfield’s house in 2015, and we found some more in Tennessee in 2017, but they were all empty or aborted in both cases. Luckily, John was able to rear some adults in 2017, and MJ reared some in 2018, so we were able to describe the species from these specimens.
75. Phytomyza flavilonicera Eiseman & Lonsdale, 2021 – This is a honeysuckle leafminer that Mike found in Oklahoma. Like P. sempervirentis, it mainly feeds on Lonicera sempervirens, and its leaf mine is pretty much identical, but the adult has a yellow face (and, of course, different genitalia). It also feeds on L. flava, and the Latin “flavus” (yellow) in its name refers to this as well as the distinctive color of its face.
76. Phytomyza triostevena Eiseman & Lonsdale, 2021 – This is another sneaky one, which John reared from horse gentian (Triosteum) in Iowa. It makes an inconspicuous leaf mine that soon disappears into a lateral vein and then the midrib, and the Latin “vena” (vein) in its name refers to this.
That’s the species tally so far… I guess for completeness I should mention that I’ve also coauthored one genus:
Aspilanta van Nieukerken & Eiseman, 2020 – I helped Erik van Nieukerken with the paper that described this new genus for six moths species that were formerly placed in the genus Antispila. They had originally been placed in that genus because of a similarity in wing pattern, but they turn out to be more closely related to the very different-looking genus Coptodisca. Just to keep things a little confusing, we made the new name an almost-anagram of the old one. I’ve found three of the described Aspilanta species in my yard, and a fourth not far away—mining leaves of grape, Virginia creeper, and sweetfern—and I’ve met the other two on wild hydrangea in Ohio and on canyon grape in Utah. Here’s the type species, Aspilanta oinophylla; it’s the most recently described of the six, named Antispila oinophylla by Erik and Dave Wagner in 2012:
In that same paper, Erik and I also moved the Florida species Antispila eugeniella Busck to Heliozela. Now, “How many species have you renamed?” is a question no one has ever asked me, but just to satisfy my own curiosity, here’s a quick rundown:
In 2019, in the paper where Tadeusz Zatwarnicki and I documented the presence of the European duckweed leafminer Hydrellia albilabris in Maine, we transferred Cavatorella jinpingensis Zhang, Yang & Hayashi to Hydrellia.
In 2020, in addition to the seven changes mentioned above that Erik van Nieukerken and I made, Don Davis and I transferredCameraria affinis and C. leucothorax to the genus Phyllonorycter, and also declared affinis a synonym of P. mariaeella. And we transferred Phyllonorycter arizonella and P. cretaceella to the genus Cameraria. All of these had been placed in the now obsolete genus Lithocolletis (along with many other gracillariids) until Don assigned them to new genera in the 1983 “Hodges” list; for whatever reason he messed up on these four, and I noticed that they were misplaced in the process of putting together my leafminer book.
And this year, Owen Lonsdale and I published a paper that reaffirmed the synonymy of Chromatomyia with Phytomyza, shortly after Michael von Tschirnhaus proposed formally rejecting all of the conclusions of the detailed molecular and morphological study by Winkler et al. (2009) (instead of merely ignoring them, as most Europeans studying Agromyzidae have been doing for the past 12 years). This also involved reaffirming that Napomyza and Ptochomyza should be treated as subgenera of Phytomyza rather than as full genera. So, in addition to explicitly renaming three Chromatomyia species that had never before been placed in Phytomyza, we implicitly renamed well over 100 species back to names that had been used for them before.
So I guess I don’t have an exact count, and the answer depends on what you think of the arguments Owen and I made in the Chromatomyia paper—which, as with all of the papers I’ve coauthored, I’m happy to share with anyone who is interested.
Anyway, thanks once again to all my collaborators and readers, and I look forward to sharing more discoveries with you in the coming year!
Nine years ago this month, I published the first in a series of “monthly mystery” posts in which I wrote about nagging natural history mysteries with the hope that someone out there might have some answers, or at least suggest some possibilities I hadn’t thought of. That first mystery was the identity of these 3 mm long banana-shaped things Noah and I found stuck to pine needles at Yosemite National Park on July 26, 2008:
As I wrote before, “In photos, they look like they could be larvae, but they were hard and stationary and in person they gave the impression of eggs. There were often several in an evenly-spaced row, and all had tips that curved away from the substrate as shown [above].” One reader suggested that they looked like fly pupae, and another thought they might be tasty with lemon juice and cocktail sauce. No further progress was made until today, when I was scouring Middlekauff’s (1958)* publication on pamphiliid sawflies for every available detail about the natural history of the conifer-feeding species. In his notes under Acantholyda zappei, he wrote:
The pale yellow eggs are about 4 mm. long, tapered at each end and crescent-shaped. The shape is most unusual for a sawfly egg and in size may well be the largest known. . . The eggs are deposited singly, their long axis parallel to that of the [pine] needle, on new growth late in June and early July. When the young larvae hatch, they emerge from the end of the egg which tapers least and begin to spin a loose web around themselves, fastening the outer threads to the needles. . .
It had never crossed my mind that a sawfly could produce eggs that big—it seemed like they would have to be from something along the lines of a katydid or a walkingstick. But sure enough, a quick Google search turned up a fantastic photo of an Acantholyda larva hatching from a very similar egg in Finland.
Acantholyda zappei is restricted to the northeastern US and adjacent Canada (which means I should be able to find its eggs in my neck of the woods, though somehow I’ve never seen anything like those Yosemite bananas in the intervening 13 years), but there are 33 other Acantholyda species in the US and Canada, 14 of which occur in California. Some of those can be eliminated as possibilities because they feed on conifers other than pines, but several have never been associated with any host, and none of their eggs have been described, so the exact identity of the Yosemite eggs will remain a mystery for some time. In fact, the only other Acantholyda in North America whose eggs have been described is the “pine false webworm,” A. erythrocephala, which is a European species that was found in Pennsylvania in 1925 and has since spread at least to New Jersey and Ontario (and apparently Alberta). Here are some eggs I found in May 2011 on a white pine needle in Burlington, Vermont that are consistent with that species:
My only other encounter with this genus (that I know of!) is this adult of A. erythrocephala I photographed in Colchester, Vermont in May 2005:
* Middlekauff, Woodrow W. 1958. The North American sawflies of the genera Acantholyda, Cephalcia, and Neurotoma (Hymenoptera, Pamphiliidae). University of California Publications in Entomology 14: 51–174.
I’m finally getting back to working on my guide to sawfly larvae, after a very busy field season followed by several weeks of focusing on writing papers—some of which have involved documenting new host records and previously undescribed sawfly larvae, so in that way I’ve already been making some progress this fall. This week I’ve started to tackle the chapter on conifer-feeding sawflies, and this morning while taking a break from that and sorting through some of my photos from this past June, I coincidentally came across some relevant photos that I couldn’t resist sharing here.
On September 20, 2020, I led one of my only public walks of the year, during which the group encountered some red-headed pine sawfly larvae (Diprionidae: Neodiprion lecontei) chomping on pitch pine needles (Pinaceae: Pinus rigida). I didn’t have any photos of this species yet, so I took three home to photograph them:
They exhibited their charming behavior of standing up and blowing bubbles when disturbed:
Now that I had these larvae I figured I might as well rear them to adults, so I kept them in a jar with some soil in the bottom and two days later they shed their skins…
…and now they were prepupae, done feeding and ready to burrow down and spin cocoons. The prepupa of this species looks much like the feeding-stage larva but its head is no longer red.
Between June 27 and July 5 of this year, three adults emerged from those three cocoons, but as in my previous “Sawfly Surprise” post, they were parasitoids rather than the sawflies I was expecting. Last time the wasps were eulophids, which is the #1 family of parasitoids I get when trying to rear leafminers, but this time they were perilampids. I’ve just encountered perilampids once before, when three adults of Perilampus platigaster emerged from cocoons of braconid wasp larvae that had emerged from some caterpillars that remain unidentified because every last one of them was parasitized. These new perilampids, which Jeong Yoo tells me are P. hyalinus, have the same adorable general appearance as P. platigaster but with the added bonus of being shiny blue and green.
I presume the above wasp is the male, and the one below (more blue, with larger abdomen) is the female.
In my previous Perilampus post I described the life cycle of perilampids that are obligate hyperparasitoids, entering caterpillars with the hope of finding primary parasitoids to parasitize. It turns out that P. hyalinus is sometimes a hyperparasitoid, e.g. emerging from puparia of tachinid flies or cocoons of ichneumon wasps that are formed within the cocoons of Neodiprion sawflies, but normally when it attacks Neodiprion sawflies it feeds as a primary parasitoid. Tripp (1962)* studied this relationship in great detail and reported that P. hyalinus females lay eggs at the bases of pine needles four to six inches away from groups of second-instar Neodiprion larvae. Why? Because it takes the eggs eight to ten days to hatch, and that’s how long it takes the young sawfly larvae to move about four inches along the stem—and there are nine days between molts, so the sawfly larvae will be in their third instar when they arrive near the eggs. Neodiprion larvae that are fourth instar or older cover much more ground and would just blow past the eggs before they hatched. Why not just lay eggs directly on the sawfly larvae? Well, before the eggs hatched the larvae would shed their skins, so that wouldn’t work out. Maybe by now they could have evolved eggs that hatch more quickly, but with the current arrangement they get to avoid that flailing, bubble-blowing behavior shown above. The fluid that comes out of the larva’s mouth is sticky and could cause major problems for a tiny parasitoid that came in contact with it.
So, the P. hyalinus larvae hatch shortly before the arrival of their slow-moving hosts, and they stand erect on the pine needles until the sawfly larvae blunder into them. Each P. hyalinus larva clings to the surface of its host for less than an hour, and then burrows inside and basically does nothing until the following spring. When the weather warms up and the Neodiprion larva starts to prepare to pupate, the P. hyalinus larva pops back out of it and feeds on it externally (but inside the cocoon), quickly paralyzing it and then devouring it over the next two weeks. It then pupates without spinning its own cocoon, and after another two weeks it emerges as an adult and chews its way out of the sawfly’s cocoon.
If I’d had to guess what series of events had led to these little blue-green wasps emerging from the soil into which my sawfly larvae had burrowed nine months earlier, I don’t know what I would have come up with but it certainly wouldn’t have been that! I’m so grateful for everyone who has taken the time to figure these things out and publish them so that curious naturalists, decades or centuries later, are able to find answers in a matter of minutes instead of having to start from scratch.
* Tripp, H. A. 1962. The biology of Perilampus hyalinus Say (Hymenoptera: Perilampidae), a primary parasite of Neodiprion swainei Midd. (Hymenoptera: Diprionidae) in Quebec, with descriptions of the egg and larval stages. The Canadian Entomologist 94: 1250-1270.
Although most of what used to be my lawn is now an untamed meadow interspersed with gardens, fruit trees, and berry bushes, once a month or so I do break out the ol’ battery-powered lawnmower to maintain a network of trails through it all. On June 11, which was one of those rare occasions, I paused my mowing and flipped over the mower to scrape out the wads of vegetation that were starting to choke up the blades. I was surprised to find a fly puparium, which I at first interpreted to have been something I had run over with the mower that had miraculously made it past the spinning blades intact.
When I found a second one just like it, I realized that these were puparia from larvae that had developed in the decomposing vegetation that I had failed to scrape off the mower after I had finished my previous mowing session. They looked similar to Anthomyiidae puparia—some anthomyiids are leafminers, but others feed on a variety of other things including decomposing vegetation—but they could have belonged to some other related family.
As I continued to dig through that crud, I found this bristly thing, which I believe to be the puparium of a lesser house fly (Fanniidae: Fannia canicularis)…
…as well as a bunch of larvae like this one:
The fact that this larva has a distinct little head capsule tells us that it’s not one of the so-called “higher” flies, like anthomyiids and fanniids, that form puparia (i.e., they pupate inside the hardened skin of the larva, which the adult ultimately escapes by inflating a big balloon from its face). It’s something more along the lines of a midge or crane fly that forms a naked, exposed pupa.
Naturally, I stuffed a chunk of this rotting plant matter, including these larvae and puparia, into a vial to see what they would all turn into. When I checked the vial on June 23 I found about a zillion of these inside:
This is Coboldia fuscipes, a member of the family Scatopsidae, which are known as the “black scavenger flies.” It belongs to the same group of “lower” flies (Bibionomorpha) as the gall midges, fungus gnats, March flies, and wood gnats, and it is the adult that goes with the larva in the previous photo. Nothing ever emerged from the puparia in the first two photos, but it turned out there was a third species of puparium-forming fly in the mix. Two females of Drosophila busckii (Drosophilidae) emerged, also on June 23, and I was able to find one of their empty puparia.
So, not quite the 30+ species of arthropods I found in a cubic foot of my lawn, but I think four species of flies in a handful of decomposing lawn isn’t too shabby either!
Thanks to Brad Sinclair for identifying the adult flies.
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