2018 Arizona Insect Collecting Trip “iReport”

Hot on the heels of the previous installment in this series, I present the sixth “Collecting Trip iReport”; this one covering a trip to Arizona during July/August 2018 with Art Evans and—like the previous installments in this series—illustrated exclusively with iPhone photographs (see previous installments for 2013 Oklahoma2013 Great Basin2014 Great Plains, 2015 Texas, and 2018 New Mexico/Texas).

This trip was a reunion of sorts—not only had it been 20 years since I’d collected in Arizona, it had also been 20 years since I’d spent time in the field with Art Evans—which just happened to be in southeast Arizona! For years I looked forward to our next opportunity, and when he told me of his plans for an extended trip to take photographs of his forthcoming Beetles of the Western United States, I couldn’t pass up the chance. Art had already been out west for five weeks by the time I landed in Phoenix on July 28th, and together we drove to Cave Creek Canyon in the Chiricahua Mountains and spent the night before beginning a 7-day adventure in and around the “Sky Islands” of southeastern Arizona.

As with the recent New Mexico/Texas post, the material collected still has not been completely processed and curated, so I don’t have final numbers of taxa collected, but there were a number of species—some highly desirable—that I managed to find and collect for the first time, e.g., the buprestids Acmaeodera yuccavoraAgrilus restrictus, Agr. arizonicusChrysobothris chiricauhuaMastogenius puncticollis, and Lampetis webbii and the cerambycids Tetraopes discoideus and Stenaspis verticalis. Who knows what as-yet-unrecognized goodies await my discovery in the still unprocessed material?!


Day 1 – Chiricahua Mountains, Cave Creek Canyon
After arriving at Cave Creek Ranch late last night, we awoke to some stunning views right outside our room!

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View of Cave Creek Canyon at Cave Creek Ranch, Chiricahua Mountains.

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Cave Creek Ranch, Cave Creek Canyon, Chiricahua Mountains.

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Cave Creek Ranch, Cave Creek Canyon, Chiricahua Mountains.

The first buprestid of the trip was a series of Pachyschelus secedens on Desmodium near Stewart Campground. We beat the oaks and acacia along the way to Sunny Flat Campground but didn’t find much. Once we got near Sunny Flat I did some sweeping in an area with new growth of Helianthus sp. and got a series of Agrilus huachucae, a few lycids, and one Leptinotarsa rubiginosa. I beat one Acmaeodera cazieri from Acacia greggii and found another on flower of prickly poppy (Argemone sp.). On the roadside at Sunny Flat I found several Acmaeodera spp. on a yellow-flowered composite – one A. rubronotata, one A. solitaria(?), and three A. cazieri. Also collected one A. cazieri on a rain gauge, Mecas rotundicollis and one as yet undetermined acanthocinine cerambycid on miscellaneous foliage, one tiger beetle (Cicindela sedecimpunctata?) on the roadside, and two orange lycids in flight.

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Majestic peaks loom over the canyon.

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Blue pleasing fungus beetle (Gibbifer californicus) – family Erotylidae.

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Me with Margarethe Brummermann.

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Reddish potato beetle (Leptinotarsa rubiginosa) is an uncommon relative of the much more well known (and despised) Colorado potato beetle (L. decemlineata).

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Margarethe Brummermann searches for beetles in Sunny Flat Campground.

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Bordered patch (Chlosyne lacinia) – family Nymphalidae.

Desert flats east of Portal, Arizona
We came to this spot to look for Sphaerobothris ulkei on joint-fir (Ephedra trifurca), but after not finding any for awhile I got distracted by some big buprestids flying around. Caught several Hippomelas sphenicus, one Gyascutus caelatus, and two Acmaeodera gibbula on Acacia rigida, and the first and third were also on Prosopis glandulosa along with Plionoma suturalis. We finally found S. ulkei – searched the area for almost three hours, and Art and I each caught two and Margarethe caught one – also one each of P. suturalis and A. gibbula. I also got a mating pair of A. gibbula on Acacia greggii. After dinner, we went back and placed an ultraviolet light – checked it a couple hours later and got a nice series of Cylindera lemniscata and a few meloids (for Jeff).

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Desert flats below Portal, Arizona – dominant woody vegetation is mesquite (Prosopis glandulosa), sweet acacia (Acacia rigida), and three-pronged joint-fir (Ephedra trifurca).

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Art Evans photographing Hippomelas planicauda in the ‘studio’ afterwards.

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Sphaerobothris ulkei, collected on Ephedra trifurca.

Day 1 of the trip ended in typical monsoon fashion – heavy, thunderous rainstorms moved into the area during late afternoon, dimming prospects for blacklighting. Still, we set them up anyway at several spots and checked them later in the evening (flood waters preventing us from going to all the spots we wanted to). Not surprisingly, the one trap that yielded interesting specimens was in the lowest (warmest) area and received the least amount of rain. For me it was a nice series of Cylindera lemniscata.

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Day 2 – Southwestern Research Station, Chiricahua Mountains, Arizona
There is a large stand of a narrow-leaved milkweed (Asclepias sp.) at the station, so we stopped by in our way up the mountain to check it for beetles. Got a nice little series of Tetraopes discoideus (tiny little guys!) on the stems as well as a few Rhopalophora meeskei, two Lycus spp., and one Pelonides humeralis on the flowers.

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Tetraopes discoideus (family Cerambycidae).

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Rhopalophora meeskei and Lycus sp. on Asclepias sp.

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At the Southwestern Research Station with Barbara Roth, Art Evans, and Margarethe Brummermann.

Road from Southwestern Research Station to Ruster Park
After leaving the SWRS on our way up to Rustler Park, we stopped to check a couple of bushes of New Mexico raspberry (Rubus neomexicanus). Margarethe thought there might be lepturines on the flowers, but instead we found a few Acmaeodera spp. and some Rhopalophora meeskei.

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New Mexico raspberry (Rubus neomexicanus).

Further up the road we made another quick stop to check roadside flowers – just a single A. rubronotata on a yellow-flowered composite, but spectacular views of the valley below.

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Looking west from the Chiricahua Mountains, Arizona.

Gayle Nelson once told me about finding Chrysobothris chiricahuae on pine slash at Rustler Park, so I was pleased to see several fresh slash piles when we arrived. I saw a Chrysobothris (presumably this species) on the very first branch in the very first pile that I looked at, but I missed it (damn!) and didn’t see any more in that pile. However, in the next pile I visited I saw two and got them both. I looked at a third pile and didn’t see any, nor did I see any more on the two previous piles that I looked at. Still, two is better than none (assuming this is, indeed, what they are!).

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Rustler Park, Chiricahua Mountains, Arizona.

Chiricahua National Monument
Not a bug collecting stop, but we wanted to drive into the monument and see the incredible rock formations which are best appreciated by driving through Bonita Canyon and then up to Massai Point. The unusual spires, columns, and balancing rocks are a result of erosion through vertical cracks in the compressed volcanic ash which was laid down in layers 25 million years ago and then uplifted. Tilting during uplift caused vertical fractures and slippage, into which water then worked its way to create today’s formations. One of the columns I saw is 143 feet tall and only 3 feet in diameter at one point near the base! Mexican jays were our constant, close companions as we hiked through the pinyon pine/oak/juniper woodland.

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Vicinity Gleeson, Arizona
There is a wash across N Ghosttown Trail with stands of Baccharis sarothroides growing along the sides. Art previously collected a single Cotinis impia on one of the plants, so we came back to check them. We didn’t find any, but we did find two fine males and one female Trachyderes mandibularis on a couple of the plants. I also found a dead Polycesta aruensis.

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Vicinity Tombstone, Arizona
Art saw Gyascutus caelatus here previously, so we came back and found them abundantly in sweet acacia (Acacia rigidula), which was in full bloom. They were extremely flighty and hard to catch, so we each got only four. I also collected one Stenaspis solitaria on the same and a Trachyderes mandibularis female in flight.

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Trachyderes mandibularis female

At another spot nearby, we stopped to look for Lampetus webbii, which Art had seen but not been able to collect when he was here a couple of weeks ago. We did not see any (but read on…), and I saw but did not collect a Trachyderes mandibularis and two Stenaspis solitaria. I also saw and photographed some giant mesquite bugs (Thasus neocalifornicus).

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Giant mesquite bugs (Thasus neocalifornicus).

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Note the heavily armed and thickened hind legs of the male (L) versus the more slender and red/black banded hind legs of the female (R).

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Not sure of the ID (other than ‘DYC’ – damned yellow composite).

The day ended enjoying steaks, Malbec, and Jameson with two of the best hosts ever!

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Day 3 – Box Canyon, Santa Rita Mountains, Arizona
Our first stop of the day was Box Canyon, a gorgeous, rugged canyon on the east side of the range. Mimosa dysocarpa was in bloom, off which I beat two Agrilus aeneocephalus, several Hippomelas planicauda, and one Stenaspis solitaria. Norm gave me an Acmaeodera cazieri that he’d collected on an unidentified yellow-flowered composite, and right afterwards I found some small, low-growing plants with purple flowers and sticky leaves (eventually ID’d as Allionia incarnata, or trailing four o’clock) to which Acmaeodera yuccavora and A. cazieri were flying in numbers. After that I crawled up top and beat the mesquites, getting one Chrysobothris sp., a mating pair of S. solitaria, and a couple of large clytrine leaf beetles.

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Box Canyon from just above the dry falls.

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Prickly poppy (Argemone mexicana) blooming along the roadside.

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Hippomelas planicauda mating pair on Mimosa dysocarpa.

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Allionia incarnata, flower host for Acmaeodera cazieri and Acm. yuccavora.

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Acmaeodera cazieri (left-center).

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Acmaeodera yuccavora.

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Lubber grasshopper (Taenipoda eques). The striking coloration warns potential predators that it is chemically protected.

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Datana sp. caterpillars.

Vicinity Duquesne, Arizona
We came here to look for Tetraopes skillmani (this is the type locality). We found the host plant (Sarcostemma sp.), but there were no beetles to be seen anywhere. Maybe another location nearby…

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Sarcostemma sp. (family Asclepiadaceae).

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Patagonia Pass, Patagonia Mountains, Arizona
We went up higher into the mountains to get into the oak woodland, where I hoped to find some of the harder-to-collect oak-associated Agrilus spp. Right away I beat one Agrilus restrictus off of Emory oak (Quercus emoryi), but no amount of beating produced anything more than a single Enoclerus sp.. I also beat the Arizona oak (Q. arizonica) and got only a single Macrosaigon sp. On Desmodium sp. I collected not only Pachyschelus secedens but a nice series of Agrilus arizonicus. For me it is the first time I’ve collected either A. restrictus and A. arizonicus, the former being quite uncommon as well, so all-in-all not a bad stop.

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Agrilus arizonicus mating pair – the males are brighter green than the females, which are more coppery.

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Unidentified plant.

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Me, Art Evans, and Norm Woodley.

Sycamore Canyon, Santa Cruz Mountains, Arizona
We came here for night lighting, but while we still had light I did some sweeping in the low vegetation and collected a mixed series of Agrilus arizonicus (on Desmodium sp.) and Agrilus pulchellus – the latter a first for me, along with two small cerambyids that could be Anopliomorpha rinconia. Conditions were perfect (warm, humid, and no moon), and we had lots of lights (Art’s five LED units, Steve’s MV/UV combo setup, and my UV setup), but longhorned beetles were scarce – just one Prionus heroicus and one Lepturges sp. for me, and Steve got a few others including a nice Aegomorphus sp. I did also collect a few scarabs – Chrysina gloriosa and Strategus alous – because they’re just so irresistible!

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A beacon in the night!

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Art, Steve, and Norm checking the lights.

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Chrysina gloriosa.

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A male oz beetle (Strategus aloeus).

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Eacles oslari is a western U.S. relative of the imperial moth (E. imperialis).

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Insects whirring around my head!


Day 4 – Prologue
One of the downsides (if you can call it that) of having great collecting is the need to take periodic “breaks” to process all the specimens and make my field containers available for even more specimens. Thanks to Steve and Norm for making their place available to Art and I so we can do this before heading out to our next set of localities.

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Copper Canyon, Huachuca Mountains, Arizona
Copper Canyon is the classic spot for finding the charismatic Agrilus cavatus (see photo), but first we did some sweeping in the low vegetation near the parking area, where Norm got one Agrilus arizonicus and two Agrilus latifrons – and gave them to me! (Thanks, Norm!) I did some beating of the oaks, and after much work I ended up with a single Agrilaxia sp. and pogonocherine cerambycid on Emory oak (Quercus emoryi) and a couple of giant clytrines on the Arizona oak (Q. arizonicus). I then started sweeping the low-growing Acaciella angustissima – right away I got two A. cavatus. They were in the area past the cattle guard on the right where lots of dead stems were sticking up, and although I continued to sweep the plants more broadly in the area I never saw another one. Finally, Norm called me up to a small Mimosa dysocarpa near the car off which he collected three Agrilus elenorae – and gave them to me! (Thanks, Norm!) I gave the tree a tap and got one more, and in my last round of sweeping I came up with a Taphrocerus sp. (must be some sedges growing amongst the grasses).

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Copper Canyon to the northwest.

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Copper Canyon to the north.

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Agrilus cavatus on its host plant, prairie acacia (Acaciella angustissima).

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Robber fly (family Asilidae) with prey (a ladybird beetle).

Bear Canyon Crossing, Huachuca Mountains, Arizona
There was quite a bit of Mimosa dysocarpa in bloom along the roadsides on the west side of the Bear Canyon crossing, which I beat hoping to find some more Agrilus elenorae. I didn’t find any, but I did get several more Hippomelas planicauda, which is a nice consolation prize – and a great photo of the last one! Other than that I did a lot of sweeping and found only a single Acmaeodera cazieri.

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Bear Canyon to the south.

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Bear Canyon to the north.

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Hippomelas planicauda on one of its hosts, velvetpod mimosa (Mimosa dysocarpa).

Appleton-Whittell Research Ranch of the National Audubon Society, Elgin, Arizona
Cool temperatures and a blustery wind discouraged most insects from finding our blacklights. However, our blacklight did find some other interesting local residents. These two individuals could be the stripe-tailed scorpion, Paravaejovis (Hoffmannius) spinigerus, a common species in Arizona and southwestern New Mexico.

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Day 5 – Miller Canyon Recreation Area, Huachuca Mountains, Arizona
There was a lot of Baccharis sarothroides growing in the lower canyon near the parking area, so I checked it all out hoping to find Tragidion annulatum. None were seen, and in fact there was very little insect life in general. I did pick up a couple of Acmaeodera solitaria by sweeping – not anything significant but the 15th species buprestid of the trip and found a dead Cotinis mutabilis, and Art got a nice series of Chalcolepidius click beetles on B. sarothroides and Prosopis glandulosa. Puzzling the lack of insect activity, given how green all the plants were and how fresh the growth looked. I guess we’ll have to look elsewhere.

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Acanthocephala thomasi, a leaf-footed bug (family Coridae).

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I was all lined up for a side shot of the bug when suddenly he took flight.

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Turkey vultures hanging out waiting for me to die!

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Actually they were all hanging out around a dead cat, some of which I scared up as they were feeding on it.

Vicinity Naco, Arizona
We decided to try some desert thorn-scrub habitat so headed east towards Bisbee. Just north of Naco we saw some habitat where it had rained recently – everything was green with the sweet acacia (Acacia rigidula) and creosote (Larrea tridentata) in full bloom. Immediately out of the car I found a Dendrobias mandibularis on Baccharis sarothroides (and when I came back to it later I found a big, major male on it – see photos). On the sweet acacia we found a handful of Gyascutus caelatus (one of which I got a nice photo of), a mating pair of Sphaenothecus bivittatus, and a Cymatodera sp. Finally, out along the roadsides a riot of different yellow composites were in full bloom, including Heliomeris longifolia off which Art got a couple of Acmaeodera solitaria and I got two specimens of a large Acmaeodera sp. (blue-black with numerous small irregular yellow spots).

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Dendrobias mandibularis – major male.

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Them’s some mandibles!

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Gyascutus caelatus on Acacia rigidula.

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A blister beetle (family Meloidae) in the genus Zonitis – either sayi or dunnianus – on Heliomeris longifolia.

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Heliomeris longifolia – host flower for both the Zonitis blister beetle and Acmaeodera sp. jewel beetle.

Vicinity Tombstone, Arizona
We decided to go back to the spot north of Tombstone where Art had earlier seen Lampetis webbii and give that species another shot. We looked at the Rhus sp. tree that he’d seen them on, and then we each followed the wash in opposite directions looking at the Rhus trees along them, which growing above the banks but never further away than about 25 feet. Along the way I collected several more Gyascutus caelatus on sweet acacia (Acacia rigida), which were more abundant this time than last and also easier to catch. After walking about 1/4-mile down the wash I saw something fly from a Rhus tree and land low on the bushes nearby. I quickly netted it, pulled it out, and was elated to see that it was, indeed, Lampetis webbii! I searched the Rhus in the area more carefully but didn’t find any more, then found some Rhus growing up along the road. At one point, I saw a large buprestid fly and land high in the top of another Rhus tree. I couldn’t tell for sure if it was L. webbii, but I extended my net as far as I could, positioned it beneath the beetle, and tapped the branch hoping it would fall in. Unfortunately, it flew away instead of dropping, so I can’t say for sure whether it was L. webbii or just a wayward G. caelatus. At any rate, L. webbii is yet another species that I have not collected before now and the 17th buprestid species of the trip.

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Lampetis webbii, collected on Rhus sp.

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Stenaspis solitaria on Acacia rigidula.

Ramsey Canyon, Huachuca Mountains, Arizona
After returning from Tombstone, we visited Pat & Lisa Sullivan at their home at the end of Ramsey Canyon. Pat is a scarab collector who runs lights at his home nightly, and after a delicious dinner we spent the rest of the evening checking the lights. I was hoping to collect Prionus heroicus, and I got my wish. Also got Prionus californicus and several other non-cerambycid beetles such as Chrysina beyeri, C. gloriosa, Lucanus mazama, and Parabyrsopolis chihuahuae (the latter a first for me). I also placed a prionic acid lure (thanks Steve!) and got three more male P. heroicus. We also hunted around the rocks and roadsides hoping to find Amblycheila baroni but didn’t find any. Art did, however, find a female P. californicus and gave it to me (thanks!).

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Meeting Pat Sullivan!

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Darkling beetles (family Tenebrionidae) such as this one come out at night to feed on decaying vegetation.

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Chrysina beyeri (family Scarabaeidae) is one of three species in the genus occurring in Ramsey Canyon.

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Black-tailed rattlesnake (Crotalus molossus), collected by Pat in Ramsey Canyon.

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Sidewinder (Crotalus cerastes lateropens), collected by Pat in Yuma County.

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“Sometimes the best collecting is inside!”


Day 6 – Vicinity Sonoita, Arizona
Unsuccessful attempt to collect Hippomelas martini, only recently described (Bellamy & Nelson, 1998) and part of the type series taken somewhere near this spot (“20 mi NE Patagonia, Hwy 82”) by “sweeping roadside vegetation”. At other locations it had been recorded on Calliandra sp., and I found patches of the plant here along and on top of the road cuts. This gives me confidence that I found the right spot, but I didn’t encounter this or any other beetles by sweeping the patches or visually inspecting them.

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Box Canyon, Santa Rita Mountains, Arizona
We decided to come back to Box Canyon since we’d had such good luck last time. I started at the spot above the dry falls where I collected so many Acmaeodera cazieri and A. yuccavora on flowers of Allionia incarnata. This time it was hotter, drier, and windier, and the flowers were semi-closed. Still I found a few of each. I then started walking down the road towards the lower canyon crossing where I would meet up with Art. Things were really hopping on the Mimosa dysocarpa, with Hippomelas planicauda abundant (finally collected my fill) and several other Buprestidae also beaten from the plants: Agrilus aeneocepahlus, Acmaeodera scalaris, Acmaeodera cazieri, Chrysobothris sp., and a species of Spectralia! (seven species of Buprestidae at one location I think is the high for the trip.) I checked other plants and flowers along the way down but didn’t find much.

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Halfway down from the “dry falls”.

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The “dry falls” about halfway up the canyon.

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Pseudovates arizonae – the aptly named Arizona unicorn mantis.

Lower Madera Canyon, Santa Rita Mountains, Arizona
Madera Canyon is perhaps the most famous insect collecting locality in Arizona – maybe in the country, and it is hard to make a visit to Arizona without stopping by here. We elected to work the lower canyon first in an area where Chrysobothris chalcophoroides has been taken on Arizona oaks (Quercus arizonicus). Hiking towards the oaks I found some Stenaspis solitaria in a Baccharis sarothroides and marveled at the variety of other insects active on the plants (see photos) – later I would also collect an elaphidiine cerambycid on the plant. Next I started working the oaks, beating every branch I could reach with my net handle. With one whack of the stick a single Paratyndaris sp. and a single Brachys sp. landed on my sheet – those would be the only buprestids I would collect off the oaks! Other than that I collected one Hippomelas planicauda on Mimosa dysocarpa for the record. While I was working the oaks up in the knoll, the weather started turning with blustery winds, and I could see the rain coming in the distance. By the time I got down from the knoll the rain had arrived, and I walked back to the car in a sunny downpour using my beating sheet as an umbrella!

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Madera Canyon in the Santa Rita Mountains.

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Acanthocephala thomasi on Baccharis sarothroides.

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What appears to be a so-called “cricket killer” wasp (Chlorion aerarium) also feeds on sap on Baccharis sarothroides.

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A longhorned beetle, probably in the genus Aneflus, rests on the foliage of Baccharis sarothroides.

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Rain headed my way!

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Rain passing into neighboring Florida Canyon.

Montosa Canyon, Santa Rita Mountains, Arizona
Just to try something different, we went to Montosa Canyon – the next canyon south of Madera Canyon – for tonight’s blacklighting. We set my sheet up just E of the crossing and Arts ground units back to the west along a gravel road on the south side of the crossing. Moths came in numbers, but the beetles were light – I collected only blister beetles (Epicauta sp.) and a Cymatodera sp. checkered beetle at the sheet, a series of tiger beetles and a female Strategus cessus at the second ground unit, and a male Strategus aloeus and two Stenelaphus alienus at the third ground unit.

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A gorgeous sunset to start the evening.

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A deepening dusk brings the promise of insects at the lights. 

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A bee assassin bug, Apiomerus flaviventris.

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An ocotillo, or calleta, silkmoth – Eupackardia calleta.

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One of the western riparian tiger beetles.


Day 7 (last day) – Vicinity Continental, Arizona
There was a photo posted on BugGuide of Stenaspis verticalis taken last week, so we decided to give it a shot and see if we could get lucky and find it ourselves. We checked all the Baccharis sarothroides within ½-mile if the spot but didn’t find it. I did, however, collect four Euphoria leucographa, two Chalcolepidius smaragdula, two Aneflus spp., and singletons of Stenaspis solitaria and Dendrobias mandibularis. I also took a couple of Hippomelas planicauda on Mimosa dysocarpa – just for the record!

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Euphoria leucographa on Baccharis sarothroides.

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Chalcolepidius smaragdinus on Baccharis sarothroides.

Lower Madera Canyon, Santa Rita Mountains, Arizona
We returned to work the lower canyon area. I’d heard that the tiger beetle Cicindelidia obsoleta santaclarae has been taken in the area last week so was hoping to run into it. While Art worked the east side of the road I worked the west, initially following FR-781 into what looked like grassland areas where the tiger beetle might occur. I didn’t see any but took Acmaeodera scalaris on Heterotheca sp. flowers and Acmaeodera solitaria on Argemone mexicana flowers. There was also a fresh wind-thrown mesquite (Prosopis glandulosa) with a bunch of Chrysobothris octocola and one Chrysobothris rossi on it. Still the area looked abused from grazing and was uninteresting, so I looked for another area to explore.

Northwest of the parking lot I spotted another grassy area that was dotted with Baccharis sarothroides, so I decided to give that area a look. After clambering several times through barbed wire fence, I reached the area and began to give it a look. Still no tiger beetles, but every time I passed a B. sarothroides I inspected it closely. I’d looked at several plants when I came upon one with a Stenaspis solitaria sitting in the foliage, and when I looked down on one of the stems and saw a big male Tragidion sp. on the underside of the stem. After securing it, I looked closer at the plant and saw a pair of annulated antennae crawling up another stem – I knew right away it was a mating pair of Stenaspis verticalis! After carefully moving to the other side to confirm, I dared to take a few photos in situ (see below) and then secured the couple. Of course, this gave me newfound motivation to work the entire area to look for more. It was very hot by then, and I was already quite thirsty, but I summoned up all the stamina that I could and worked as many plants as I could, ending up with six Tragidion spp. and three Stenaspis verticalis. The latter was one of my top priority targets for this trips, and the only thing more satisfying than getting it is doing so on my last day on the field.

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View to south edge of Madera Canyon – Elephant Head is at the right.

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Chrysobothris octocola female ovipositing on freshly killed mesquite (Prosopis glandulosa).

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Tragidion sp. mating pair on Baccharis sarothroides.

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Mating pair of Stenaspis verticalis on Baccharis sarothroides.

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Chalcolepidius lenzi at a sap flow on Baccharis sarothroides.

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Lateral view of Chalcolepidius lenzi.

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Barrel cactus in bloom.

Montosa Canyon, Santa Rita Mountains, Arizona
We  returned to Montosa Canyon and stopped at the Astronomy Vista partway up. It was hotter than bejeebuz! There was not an insect to be seen except giant cactus bugs and a single Euphoria leucographa that Art found on a sapping Baccharis sarothroides. Temp was 103°F even at this elevation!

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Stunning vista during the day! 

We needed to escape the heat, and I wanted to see oaks for one more crack at Mastogenius, so we drove up to the 13-km marker and I collected on the way back down to below the 12-km marker. Conditions were much more agreeable (temps in the 80s), and near the top there was a Ceanothus sp. bush in bloom, off which I collected Rhopalophora meeskei and Stenosphenus sp. – both genera represented by individuals with black versus red pronotum. Then I started beating the (Mexican blue, I believe) oaks, and right away I got a Mastogenius sp.! Kinda small, so I’m thinking not M. robusta and, thus, probably M. puncticollis (another species new to my collection). I also beat a largish Agrilus sp. that I don’t recognize, a few clerids, two R. meeskei, one Stenosphenus sp., and a couple of leaf beetles. There was also another type of oak there – Arizona white, I believe, which I beat as well but only got one clerid.

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Spectacular views from 7000 ft!

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A lichen moth on flowers of Ceanothus sp.

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The biggest, fattest, bristliest tachinid fly I have ever seen!

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The spectacular vistas just keep on coming!

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An ancient alligator juniper stares down yet another sunset (perhaps its 50 thousandth!).

We stopped by the Astronomy Vista again on our way back down the canyon, and I found a pair of Moneilema gigas on cholla (Opuntia imbricata).

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Obligatory dusk shot of Moneilema gigas on Opuntia imbricata.

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Another individual on the same plant.

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Sunset over “Las Cuatro Hermanas”.

It was a fantastic seven days in the field with Arthur, and it was a great pleasure to (in some cases, finally) meet Margarethe, Barbara, Steven, Norm, and Pat. I appreciate the warmth, generosity, and hospitality that all of them displayed to me and look forward to our next encounter, hopefully in the near future.

© Ted C. MacRae 2019

Fun with eucraniines!

During my February/March 2015 visit to Argentina, I had the opportunity to travel to west-central provinces of San Juan and San Luis with Federico Ocampo for a weekend of insect collecting. Up to that point most of my collecting in Argentina had been limited to the northeastern provinces (Chaco, Corrientes, and Misiones), so I was excited for the chance to explore a radically different biome. West-central Argentina represents a transition zone from the flat, wet, treeless plains of the Humid Pampas in east-central Argentina (Buenos Aires, Santa Fe, and Córdoba Provinces) to the massive Andes Mountains running along the western edge of South America. This area is home to the Monte, a desert biome characterized by volcanic sediments, piedmont plains, large mountain blocks and dry salt lakes. Conditions in the Monte are generally more hospitable than in the neighboring Atacama and Patagonian Deserts lying north and south of the Monte, respectively. As a result, the flora and fauna in the Monte is relatively rich and characterized by a diversity of shrubs, grasses, and cacti.

Dunas de Encón

Encón Dunes, San Luis Province, Argentina

Of the several sites we visited in the area, the most remarkable was “Las Dunas de Encón” (the Encón Sand Dunes) in San Luis Province. Belonging to a larger system covering some 250,000 hectares—the largest in South America (and, thus, sometimes called the “Argentinian Sahara”)—the dunes are thought to have formed some 100–200K years ago as a result of dry conditions brought on by Quaternary glaciations. I find sand dune systems endlessly fascinating due to their unique and often endemic plants and animals and have visited many systems in North America (Bruneau, Coral PinkGlamisGreat, Medora, St. Anthony, and others), but this was the first sand dune system I’ve had the opportunity to see outside of the U.S. Federico, a scarab specialist, shares that fascination and has, in fact, described a number of species in the scarabaeine tribe Eucraniini—endemic to South America—that utilize these very sand dunes (Ocampo 2005, 2007, 2010). He was hoping one or more of them might be out and about; I was hoping to see anything, really.

Host for Lampetis spp.

Parkinsonia praecox? – adult host plant for Lampetis baeri and L. corinthia.

One of the first plants that caught my attention was a woody, fabaceous shrub that looked very much like what I would have previously called Cercidium, now Parkinsonia, and which after a bit of digging I conclude is likely Parkinsonia praecox. Woody, fabaceous shrubs in desert habitats are a sure bet to host jewel beetles, so I began paying special attention to each shrub as I wandered by. It wasn’t long before I saw a large, brilliant metallic green jewel beetle sitting on an outer branch of one of the shrubs—it was one of the most beautiful jewel beetles I have ever seen out in the field with my own eyes! I managed to catch it, and over the next few hours I collected not only several more of this species but also several individuals of an even larger, more somber-colored species. I was able to identify them as Lampetis baeri (Kerremans, 1910) and L. corinthia (Fairmaire, 1864), respectively, when I compared them to material in the collections at Fundacion Miguel Lillo, Instituto de Entomologia, Tucuman, Argentina [IFML]) during my visit there the following week (see photos below).

Lampetis baeri (Kerremans, 1910)

Lampetis baeri (Kerremans, 1910) [IFML]

Lampetis corinthia (Fairmaire, 1864)

Lampetis corinthia (Fairmaire, 1864) [IFML]

As a jewel beetle enthusiast, you would think that was the highlight of my day. In fact, the fun had only started. For a time after our arrival, Federico pointed out burrows likely made by eucraniine adults, but we didn’t see any evidence of activity at first. It wasn’t long, however, before we found the first adult—a fine Eucranium beleni Ocampo, 2010, the largest of the three species occurring at this site (about the size of our North American Deltochilum). One of the more obvious features of eucraniines is their enormously enlarged forelegs and pronotum to hold the musculature required to carry—that’s right, carry!—provisions to the larval burrow (in contrast with the more commonly seen habit among members of the subfamily of using the hind legs to push provisions to the burrow). This unusual morphology gives these beetles not only an amusing, shuffling gait but also a rather comical method for turning themselves upright (as seen in this video narrated by Federico). There are other dung beetles that pull, rather than push, larval provisions (e.g., Sisyphus spp., which stand on highly elongate hind legs and walk backwards while pulling the dungball), but eucraniines seem to be the only ones that actually lift provisions off the ground to carry them. In the case of E. beleni, this involves carrying pieces of dung with the forelegs held out in front of the head while walking forward on the middle and hind legs (Ocampo 2010). I didn’t get to see that behavior with E. beleni, but I did see it with one of another of the eucraniines we found that day (see below). In the E. beleni photo below, note the brushy middle and hind tarsi—an adaptation for walking on loose sand.

Eucranium belenae

Eucranium belenae Ocampo, 2010 walks on its middle/hind legs while holding its forelegs aloft.

Eucranium belenae burrow

Eucranium belenae burrow plugged with a piece of dung.

The second species in the group that we encountered was Anomiopsoides cavifrons (Burmeister, 1861). This species is much smaller than E. beleni (about the size of a large Onthophagus), and unlike E. beleni—and, in fact, most other dung beetles—the larvae of A. cavifrons develop on plant matter rather than dung. Both males and females provision the larval burrows with pieces of plant debris that they pick up with their front legs and carry back to the burrow while walking on their other four legs. This rather amusing video shows a male bringing a piece of debris back to his burrow, then exiting to find and retrieve another piece of debris to bring back to the burrow. The molar region of their mandibles is heavily sclerotized for masticating the plant fibers in preparation for the larvae. There are a couple of other species in the tribe that opportunistically include plant matter in their diet, but A. cavifons seems to be the only one known to utilize dry plant matter in desert habitats almost exclusively (Ocampo 2005). Anomiopsoides cavifrons was far more abundant in the dunes than E. beleni, and by early to mid-afternoon they were encountered with such regularity that I stopped even looking at them.

Anomiopsoides cavifrons male at burrow

Anomiopsoides cavifrons (Burmeister, 1861) male at burrow entrance.

We also were fortunate to see a few individuals of the third species known from these dunes, Anomiopsoides fedemariai Ocampo, 2007. This species is intermediate in size between the extremes represented by E. beleni and A. cavifrons and utilizes pellets of the plains viscacha (Lagostomus maximus), a species of rodent in the family Chinchillidae, for food (Ocampo 2007).

REFERENCE:

Ocampo, F. C. 2005. Revision of the southern South American endemic genus Anomiopsoides Blackwelder, 1944 (Coleoptera: Scarabaeidae: Scarabaeinae: Eucraniini) and description of its food relocation behavior. Journal of Natural History 39(27):2537–2557 [pdf via DigitalCommons].

Ocampo, F. C. 2007. The Argentinean dung beetle genus Anomiopsoides (Scarabaeidae: Scarabaeinae: Eucraniini): description of a new species, and new synonymies for A. heteroclytaRevista Sociedad Entomología Argentina 66(3–4):159–168 [pdf via SciELO Argentina].

Ocampo, F. C. 2010. A revision of the Argentinean endemic genus Eucranium Brullé (Coleoptera: Scarabaeidae: Scarabaeinae) with description of one new species and new synonymies. Journal of Insect Science 10:205, available online: insectscience.org/10.205 [pdf via DigitalCommons].

© Ted C. MacRae 2016

Beetle Collecting 101: Fermenting bait traps for collecting longhorned beetles

One of the most useful collecting techniques for those interested in longhorned beetles (families Cerambycidae and Disteniidae) is fermenting bait traps. I was first clued into the use of such traps soon after I began collecting these beetles in the early 1980s and encountered a series of rather old publications by A. B. Champlain and S. W. Frost detailing their usefulness and the diversity of species found to be attracted to them. Champlain & Kirk (1926) listed 15 species of Cerambycidae attracted to bait pans containing a mixture of molasses and water. This list was expanded to 37 species by Champlain & Knull (1932), who noted that a mixture of one part molasses to ten parts water in a gallon-pail seemed to give the best results. Frost & Dietrich (1929) listed 20 species captured with a mixture of one part molasses to 20 parts water. Twelve of the species they mentioned were not listed by Champlain & Knull (1932), and the list of Frost (1937) included two additional previously unrecorded species.

I made extensive use of fermenting bait traps during my 1980s survey of longhorned beetles in Missouri (MacRae 1994) using a mixture of one part molasses, one part beer, nine parts tap water, and a sprinkling of dry active yeast to start fermentation. This recipe was based on that of Champlain & Knull (1932) (although I must confess that I do not remember where I got the idea to add beer and yeast). During that study, I collected 13 species of longhorned beetles using this method and found in other collections specimens of three additional species also collected with fermenting baits. Of the species I collected, the most significant was a large, attractive Purpuricenus that closely resembled P. axillaris (which was also collected in the traps) but clearly was not that species. These eventually proved to be undescribed after I was able to examine type material in the Museum of Comparative Zoology at Harvard University, leading to a review of the genus in North America and the description of the new species as P. paraxillaris (MacRae 2000). Since then I’ve employed fermenting bait traps to collect Cerambycidae in other parts of the country (MacRae & Rice 2007), and I now have records of 72 species of U.S. Cerambycidae documented as being attracted  to fermenting baits.

Molasses-beer fermenting bait trap

Molasses-beer fermenting bait trap.

My interest in this technique was renewed some years ago when I finally succeeded in collecting the spectacular Plinthocoelium suaveolens in fermenting bait traps placed on glades in extreme southwestern Missouri. During my Missouri survey, I had done the bulk of my bait trapping along the edges of glades just south of St. Louis in Jefferson County, and while I had a record of this species in those glades I had never collected it there myself. Finally, last year I observed one of the host trees (gum bumelia, Sideroxylon lanuginosum) on these glades with the characteristic P. suaveolens larval frass pile at the base of the trunk, prompting a renewed effort this past season to collect the species there using fermenting bait traps. In early June I placed a series of traps at Valley View Glades Natural Area (~4 miles NW of Hillsboro) and Victoria Glades Natural Area (~2.5 miles S of Hillsboro). At both locations four traps were placed along the upwind interface between dry, post oak woodland and dolomite glades. Traps were spaced about 50–100 yards apart and hung to ensure exposure to sunlight but minimize the chance they would be discovered by vandals. Each trap consisted of a 2-L plastic bucket with a small hole drilled near the rim on each side and a length of wire attached to allow hanging from a nail in the side of a tree. Two baits were used: 1) molasses/beer, and 2) red wine. The molasses/beer recipe was based on Guarnieri (2009)—more concentrated that what I have used previously, and was prepared by combining a 12-oz (355 mL) jar of dark molasses with an approximately equal volume of tap water in a 1-L plastic bottle, agitating thoroughly, and bringing to one liter volume with tap water. At the trap site, about 500 mL of diluted molasses was added to the trap, followed by a 12-oz can/bottle of beer and one-half of a 7-g packet of dry, active yeast. Red wine bait was a cheap jug variety, undiluted, with about 500 mL added to the trap. Molasses/beer and red wine were alternated in the traps at each location and replaced every two weeks or if excessively diluted by rain or evaporated during hot, dry conditions. Traps were checked weekly from early June to mid-September by pouring the trap contents through a kitchen strainer over an empty bucket and transferring beetles with forceps to empty vials. Once back at the vehicle, tap water was added to each vial and the vial agitated to rinse the specimens and remove bait residue. The water was decanted and the beetles blot-dried with paper towels before transfer to clean vials containing tissue and ethyl acetate to halt decay and maintain the beetles in a relaxed state for pinning.

Cerambycidae from fermenting bait trap

A charismatic trio of Cerambycidae from fermenting bait traps at Victoria Glades: Purpuricenus paraxillaris (left), Plinthocoelium suaveolens (center), and Stenelytrana emarginata (right).

A note about my preferred trap design. I have always used open-top buckets (previously 1-G metal, now 2-L plastic), but “window jugs” (i.e., ½-G milk or juice jugs with holes, or “windows”, cut in the sides) are also commonly used. I have not directly compared buckets with window jugs; however, I favor buckets because I believe beetles attracted to window jugs are more likely to “perch” on the trap itself rather than fall directly into the bait. I also believe that beetles, once trapped, are more likely to escape from window jugs because the window edges provide “grab” sites for beetles before they succumb. The risk of escape can be reduced if the bait surface lies well below the bottom edge of the windows, but this then limits the quantity of bait that can be used. In my experience, 500–750 mL is the minimum volume of bait that is needed to last the duration of the two-week fermentation cycle without evaporating to the point that it is not deep enough to quickly submerge beetles falling into it. Some may be concerned that open-top buckets are prone to dilution by rain, but in my experience this happens infrequently and I have not noticed diluted bait to be any less effective at attracting beetles. Rain shields, on the other hand, only serve to provide a potential perch for beetles attracted to the trap.

Plinthocoelium suaveolens

Plinthocoelium suaveolens captured in flight near its host tree, gum bumelia (Sideroxylon lanuginosum), at Victoria Glades.

A total of 558 longhorned beetles representing 16 species were collected from the traps over the course of the season (see list below). Of these, 339 specimens representing 14 species were attracted to molasses/beer, while 219 specimens representing 14 species were attracted to red wine. Ten species were represented by more than two specimens and were attracted to both bait types, the most desirable being Plinthocoelium suaveolens (41 specimens), Purpuricenus axillaris (20 specimens), P. paraxillaris (3 specimens), and Stenelytrana emarginata (6 specimens). The number of P. suaveolens collected is remarkable, considering that it was not collected during my previous trapping effort spanning several years in the 1980s. It may be significant that 1) the molasses/beer recipe used in this study was considerably more concentrated than that used in the 1980s, and 2) nearly twice as many specimens were collected in red wine (not used in the 1980s) compared to molasses/beer. I routinely examined the gum bumelia trees during my weekly visits in an attempt to find adults on their host, especially during flowering, but encountered only a single adult in flight near one of the trees—a curious result given the diurnal habits and large, conspicuous appearance of the adults. All other species collected in numbers were more attracted to molasses/beer, with the significant exception of Purpuricenus paraxillaris. Seven species taken this season were not detected with fermenting bait traps in the 1980s, bringing to 23 the number of species collected by this method in Missouri. One species, Strangalia sexnotata, is documented from fermenting bait for the first time in this study.

2015 fermenting bait trap catch

2015 fermenting bait trap catch, box 1 of 3 (click to enlarge).

2015 fermenting bait trap catch, box 2 of 3 (click to enlarge).

2015 fermenting bait trap catch, box 2 of 3 (click to enlarge).

2015 fermenting bait trap catch

2015 fermenting bait trap catch, box 3 of 3 (click to enlarge).

Longhorned beetle species and numbers taken in fermenting bait traps in 2015—most to least abundant (MB = molasses/beer, RW = red wine):

  1. Elaphidion mucronatum – 254 (MB = 176, RW = 78)
  2. Eburia quadrigeminata – 145 (MB = 73, RW = 54)
  3. Plinthocoelium suaveolens – 41 (MB = 14, RW = 27)
  4. Neoclytus scutellaris* – 32 (MB = 26, RW = 6)
  5. Parelaphidion aspersum – 26 (MB = 18, RW = 8)
  6. Purpuricenus paraxillaris – 20 (MB = 6, RW = 14)
  7. Orthosoma brunneum – 13 (MB = 8, RW = 5)
  8. Neoclytus mucronatus* – 8 (MB = 6, RW = 2)
  9. Stenelytrana emarginata* – 6 (MB = 5, RW = 1)
  10. Purpuricenus axillaris – 3 (MB = 2, RW = 1)
  11. Enaphalodes atomarius – 2 (MB = 1, RW = 1)
  12. Strangalia famelica solitaria* – 2 (MB = 2, RW = 0)
  13. Typocerus velutinus* – 2 (MB = 1, RW = 1)
  14. Xylotrechus colonus* – 2 (MB = 0, RW = 2)
  15. Elytrimitatrix undatus – 1 (MB = 1, RW = 0)
  16. Strangalia sexnotata** – 1 (MB = 0, RW = 1)

* Not previously reported at fermenting baits in Missouri.
** Not previously reported from fermenting baits anywhere.

With regards to other insects, no attempt was made to quantify their occurrence or diversity, but a few interesting specimens were collected. Elateridae (click beetles) and other beetles were notable by their absence, in contrast to the great diversity recorded from by Champlain & Knull (1932). Flower scarabs were the exception, with two Euphoria inda and a moderate series of E. sepulchralis taken only in red wine traps. The most common non-beetle insects encountered were moths, flies, and stinging wasps, for which molasses/beer seemed to be much more attractive than red wine. The majority of the wasps were Vespidae, but a few large Crabronidae (one Sphecius speciosus and two Stizus brevipennis, I think) and at least two species of Pompiliidae were collected (see box 3 image above).

The diversity of longhorned beetles collected this season was undoubtedly influenced by habitat selection for trap placement (interface between dry, post-oak woodland and dolomite glade). Different habitats would likely yield different species, although prior experience seems to suggest that traps placed in open woodlands are more productive than those placed in dense forests. Recently thinned forests may have good potential due to an abundance of dead wood from thinning operations and trees stressed by sudden exposure to sunlight. Plans are currently underway to place traps (both molasses/beer and red wine) in a variety of wooded habitats during the 2016 season.

REFERENCES:

Champlain, A.B. & H. B. Kirk. 1926. Bait pan insects. Entomological News 37:288–291 [Biodiversity Heritage Library].

Champlain, A. B. & J. N. Knull.  1932. Fermenting bait traps for trapping Elateridae and Cerambycidae (Coleop.).  Entomological News 43(10):253–257.

Frost, S. W. 1937. New records from bait traps. (Dipt., Coleop., Corrodentia). Entomological News 48:201–202 [Biodiversity Heritage Library].

Frost, S. W. & H. Dietrich. 1929. Coleoptera taken from bait-traps. Annals of the Entomological Society of America 22(3):427–436 [abstract].

Guarnieri, F. G. 2009. A survey of longhorned beetles (Coleoptera: Cerambycidae) from Paw Paw, Morgan County, West Virginia. The Maryland Entomologist, 5(1):11–22 [pdf].

MacRae, T. C. 1994. Annotated checklist of the longhorned beetles (Coleoptera: Cerambycidae and Disteniidae) known to occur in Missouri. Insecta Mundi 7(4) (1993):223–252 [pdf].

MacRae, T. C. 2000. Review of the genus Purpuricenus Dejean (Coleoptera: Cerambycidae) in North America. The Pan-Pacific Entomologist 76:137–169 [pdf].

MacRae, T. C. & M. E. Rice. 2007. Distributional and biological observations on North American Cerambycidae (Coleoptera). The Coleopterists Bulletin 61(2):227–263 [pdf].

© Ted C. MacRae 2015

Insect Identifications and Etiquette

I’ve been a student of insects for most of my life, and of the many aspects of entomology that interest me, field collecting and identification remain the most enjoyable. My interest in beetles first began to gel during my days at the university (despite a thesis project focused on leafhoppers), and early in my career I settled on wood-boring beetles (principally Buprestidae and Cerambycidae) as the taxa that most interested me. To say that species identification of these beetles can be difficult is an understatement, but I was fortunate to have been helped by a number of individuals—well-established coleopterists—who freely shared their time and expertise with me during my early years and pointed me in the right direction as I began to learn the craft. Some of the more influential include colleagues that have since passed (e.g., Gayle Nelson, John Chemsak, Chuck Bellamy, and Frank Hovore) and those that, thankfully, continue with us (e.g., Rick Westcott and Henry Hespenheide).

It has been a little more than 30 years now since I began studying these beetles, and due in great part to the help I received early on and the motivation that it inspired within me, I have gained a certain amount of proficiency in their identification as well. Not surprisingly, I too regularly receive requests from people looking for help with identifications. I rarely turn down such requests (in fact, I don’t think I have ever turned one down)—it not only helps my own research but also, occasionally, allows me to fill a gap or two in my collection. More importantly, however, it is my duty—I benefited greatly from those who shared their expertise with me, so it’s only fair that I continue by their example.

As common a practice as this is among collectors, it seems odd that there are few written guidelines on the etiquette of requesting and providing identifications. Note that this is something different than borrowing specimens for study, which has its own set of expectations and responsibilities. As someone who has both requested and received requests for specimen identifications for a long time now, I have my own thoughts about reasonable expectations in this regard. Perhaps you, too, will find these thoughts useful the next time you contemplate asking somebody to identify your specimens (or accepting a request to do so).

Guidelines for requesting identifications

  1. Always ask permission to send specimens before doing so. ‘Nuff said.
  2. When you do send specimens, read  and follow the guidelines suggested to avoid creating additional work for the identifier who must repair specimens damaged in shipment.
  3. Leave extra room in the specimen box. While tightly packed specimens minimize shipment size and can reduce cost, it also increases risk of damage during shipment due to ‘bumping’ or during removal from the box for ID. More importantly, it allows little or no room for the addition of identification labels to specimens. Additionally, many identifiers find it helpful to remove all of the specimens from a box and group them by related taxa to facilitate identification. The reassembled specimens may require more space than they did in their original arrangement.
  4. Send the entire available series of specimens. A common practice among those sending specimens for ID is to hold back specimens from a series and send only one or a few examples. Whether this is to, again, minimize the size of the shipment, confirm a provisional ID, or safeguard specimens perceived as desirable, it nevertheless prevents the identifier from having access to the range of data and variability represented in the series. This is important if the series contains 1) multiple species, 2) previously undocumented distributions or ecological data, or 3) unusual morphological variants. An exception to this is when very long series of specimens are available and sending the entire series would be unwieldy and/or unnecessary. In this case, the identifier should be informed that only a partial series of specimens was sent.
  5. Allow retentions. It doesn’t happen often, but sometimes individuals have balked at my requests to retain specimens that proved useful for my studies. This is poor etiquette, as it shows little respect for the value of the service being provided by the person making the identifications. More common is to allow retention of examples from a series, but not singletons. This also, in my opinion, is poor etiquette. I remember one of my early sendings to Gayle Nelson that contained a single specimen of Agrilus audax, a very rare North American buprestid known by only a handful of specimens. Not surprisingly, Gayle did not have this species in his collection, and while I, too, was a student of the group I didn’t hesitate to give this specimen Gayle—established and well-respected expert of the family that he was. To this day the species remains unrepresented in my collection, yet I have never second guessed that decision due to the value of what I gained in his respect and mentorship in the years since. Most identifiers are both humble and sparing in their requests for retentions.¹
  6. Allow time for identifications. Individuals with expertise in a given group are generally few in number, and those willing to provide identifications may be fewer still. As a result, they usually have a number of boxes on hand at any one time awaiting identification. Get an idea from them at the start of how long they expect it will be before they can complete the task. If the projected timeline passes and you don’t hear back from them, an inquiry is fine, but be polite and understanding.

¹ A corollary to this asking for specimens in exchange for specimens retained. An exchange involves two parties sending each other specimens that mutually benefit each other’s collections. Identifications are a service provided by one party that benefit the requester. To suggest an exchange as ‘payment’ for retained specimens ignores the value of the service being provided by the identifier

Guidelines for providing identifications

  1. Once specimens are received, protect them from damage as you would your own collection. Maintain them in a protective cabinet or check them regularly to ensure that dermestid pests do not gain a toehold.
  2. Provide the identifications in as timely a manner as possible. This is not always easy, especially for those willing to accept a large number of requests and who may find themselves inundated with boxes awaiting identification. If you cannot provide identifications relatively quickly, be honest with the requestor regarding how long you expect the identifications to take. If it does take longer, provide an update to the requestor and give them the option to have the specimens returned or confirm that they are okay with the delay.
  3. Add your identification label with your name and date (year) to at least the first specimen in the series. Even better is if you can add a small, pre-printed ID label to every specimen in the series, but this can be difficult if the number of specimens and/or diversity of species is large. If there are specimens with prior identifications that you disagree with, turn the prior ID label upside-down, replace through an existing pin hole, and add your ID label. I disagree with the practice of folding prior ID labels—not only could I be wrong, but this unnecessarily damages something with historical value, especially if new pin holes are added to the label. Always place your ID label below any existing labels (i.e., label order should reflect their sequence of placement—oldest labels nearest the specimen and newest labels furthest away).
  4. Keep retentions to a minimum. I generally ask to retain specimens only when they significantly improve the representation in my collection or provide significant new data—i.e., un- or under-represented species, undocumented distributions or ecological data, etc. The bar for singletons is even higher—usually only if they are completely absent from my collection (with ~65% of U.S. Buprestidae now represented in my collection, this is an increasingly uncommon occurrence).
  5. Following #4, provide an accounting of retained specimens. Minimally, a list of species and their number should be given, and my preference is to provide label data as well (especially if requested). I once sent a batch of beetles (in a family in which I do not specialize) to an expert for identification, and when I received them back it was obvious that a number of specimens had been retained (perhaps 1/3 of the total number). When I wrote to the identifier and asked for an accounting (remember, I was only asking for an accounting—I did not have a problem with the retentions themselves), I received a rather terse reply from the individual stating that he did not ‘have time’ to provide this. Needless to say, this level of dismissiveness was not appreciated, and I have since found another more agreeable researcher with expertise in that family to send specimens for identification.
  6. When you are ready to return the specimens, read  and follow it’s suggested guidelines to avoid causing damage to the specimens whose care you were entrusted.

Again, these guidelines are written from the perspective of a private individual sending and receiving specimens for identification. Scientists at institutions may have additional or differing guidelines on this subject, but in any case these guidelines should be communicated to and understood by individuals requesting identifications before any material is sent.

If you have additional suggestions or comments on how these guidelines can be improved I would appreciate hearing them.

© Ted C. MacRae 2015

Buprestidae type specimens at Fundación Miguel Lillo, Argentina

During my most recent visit to Argentina this past February and March, I had the chance to go behind the scenes and visit the entomology collection at Fundación Miguel Lillo, Instituto de Entomología, Tucumán. It’s always a treat to visit any entomology collection—public or private—at any location. When the collection has holdings of Buprestidae, so much the better. Much to my delight, however, this collection not only had holdings of Buprestidae (not surprisingly representing primarily Argentine species), but also a small collection of type specimens designated by Antonio Cobos Sanchez (1922–1998). Cobos was one of the 20th century’s most prolific students of Buprestidae, with publications in the family spanning the period from 1949–1990 (coincidentally, 1990 being the year of my very first buprestid publication!). I was graciously allowed to photograph these specimens, some of which present interesting nomenclatural situations that are worthy of comment. These are presented below with my notes.

Jose xx & Ted MacRae

Looking at the insect collection at Fundación Miguel Lillo, Argentina.


Sufamily POLYCESTINAE

Tribe TYNDARINI

Tylauchenia golbachi Cobos, 1993 (currently placed in Oocypetes)

Tylauchenia golbachi Cobos, 1993. The species was moved to the genus Ocypetes.

Lapsus calami or mislabeled type specimen? Cobos (1973) described Tylauchenia golbachi from Argentina (now placed in the genus Ocypetes), stating the type locality as “6 kms. N. de Belén, 1.240 m. alt., Catamarca, Argentina (Willink, Terán y Stange coll., con trampa de Malaise, 1-15-I-1970…)”. The specimen above bears the holotype label, but the locality label clearly shows that it was collected in Tucumán rather than Catamarca and that the collector’s name is Guanuco rather than the above stated names. Interestingly, in the same publication Cobos gives the allotype female collection data as “San Pedro de Colalao, Tucumán, Argentina (Coll. Guanuco, 9-III-1949)”. At first I thought this might actually be the allotype rather than the holotype; however, 1) the specimen clearly bears a holotype label, and 2) it is also clearly a male based on the dissected genitalia preserved on the label below the specimen. There are two possible explanations, both of which make it difficult to determine what is the true type locality: 1) the holotype and allotype specimens are correctly labeled, but Cobos simply transposed their label data in his publication describing the species, making Tucumán the true type locality, or 2) the holotype and allotype locality labels were switched at some point and the true type locality is Catamarca, as stated in the publication in which the species is described. This latter possibility is more serious, as in addition to the doubts it generates regarding the type locality it also raises concern about the integrity of the holotype specimen. The latter explanation, however, seems less likely, as it is more difficult to imagine a scenario where only the locality label but not the others was switched than to imagine a transposition of label data in the publication. Sadly, at this point, there seems no easy way to determine which of the two explanations is correct.

Subfamily CHRYSOCHROINAE

Tribe DICERCINI

Lampetis tucumana monrosi Cobos (nomen nudum?)

Lampetis tucumana “monrosi” Cobos (ms name?)

A manuscript name? Cobos never actually proposed a subspecies “monrosi” for Lampetis tucumana (Guérin-Méneville & Percheron, 1835) (the name on the separate box label is misspelled). He did use the name for two other buprestid taxa: Tetragonoschema monrosi Cobos, 1949—now regarded as a synonym of T. argentiniense (Obenberger, 1915), and Anthaxia monrosi Cobos, 1972—now placed in the genus Agrilaxia. The holotype label on the specimen clearly states “Lampetis tucumana monrosi” in Cobos’ handwriting, so one can only presume that Cobos had identified this specimen as representing a distinct subspecies but never followed through and actually described it.

Ectinogonia (Pseudolampetis) fasciata metallica Cobos, 1969

Psiloptera (Pseudolampetis) fasciata metallica Cobos, 1969. Pseudolampetis was later considered a subgenus of Ectinogonia but is now regarded as a full genus.

Oh, what a tangled web we weave! Cobos (1969) originally described this taxon as a subspecies of Psiloptera (Pseudolampetisfasciata Kerremans, 1919. Moore (1986) moved Pseudolampetis to a subgenus of Ectinogonia, which resulted in two taxa in the latter genus bearing the name “metallica“—Ectinogonia (Pseudolampetisfasciata metallica (Cobos, 1969) and Ectinogonia metallica Fairmaire, 1856—the latter now considered a synonym of E. speciosa (Germain, 1856). In taxonomic nomenclature, two taxa in the same genus cannot bear the same name—a situation known as homonymy. In such cases, the older name has priority and the younger name, in this case Cobos’, must be replaced. This was done by Bellamy (2006), who proposed the new name moorei for this subspecies, resulting in the name Ectinogonia (Pseudolampetis) fasciata moorei Bellamy, 2006. To bring some level of absurdity to the situation, the subgenus Pseudolampetis was subsequently raised to full genus rank, being listed as such in the recent world catalogue (Bellamy 2008), and since Cobos’ name was not originally proposed within the genus Ectinogonia it no longer competes with Germain’s name in that genus. As a result, there is no homonymy and Cobos’ original name must once again stand as Pseudolampetis fasciata metallica (Cobos, 1969), while Bellamy’s replacement name must be regarded as unnecessary. This fact seems to have been overlooked when Pseudolampetis was raised to genus rank, as Cobos’ taxon is still listed in the world catalogue as “Pseudolampetis fasciata moorei (Bellamy, 2006)”! This situation is a perfect example of just how complicated these situations can be to identify, track, and update. The type locality for the unique female is given as “Chagual, 1.200 metros de altitud, Rio Marañón, en el Perú, VIII-1953 (B. Fernández leg.)”.

Subfamily BUPRESTINAE

Tribe STIGMODERINI

Conognatha rufiventris weyrauchi Cobos, 1969

Conognatha rufiventris weyrauchi Cobos, 1969. The taxon is now considered a synonym of Conognatha abdominalis Waterhouse, 1912.

Insufficient grounds. Cobos (1969) regarded this specimen from Peru as subspecifically distinct from Conognatha rufiventris Waterhouse, 1912 from Brazil based on a suite of subtle character differences and named the taxon Conognatha rufiventris weyrauchi in honor of Prof. W. Weyrauch, who made made the holotype specimen available to him for study. Moore & Lander (2010) considered that the taxon did not represent C. rufiventris, but rather was a uniquely colored specimen of Conognatha abdominalis Waterhouse, 1912. The holotype is a male with the type locality given as “del Valle de Chatichamayo, a 1.200 m., en Peru (J. Schuiike leg.)”.

Conognatha amphititres Cobos, 1958 (syn. of Buprestis amoena Kirby, 1818; currently placed in Conognatha)

Conognatha amoena amphititres Cobos, 1958. The taxon is now considered a synonym of C. amoena (Kirby, 1818).

Insufficient grounds—part II. Cobos (1958) regarded this specimen from Brazil as subspecifically distinct from C. amoena (Kirby, 1818—originally described in the genus Buprestis) based on subtle characters and gave it the name Conognatha amoena amphititres (no etymology was given for the subspecies name). Moore & Lander (2006) regarded these differences as insufficient for subspecies status and placed the taxon as a synonym of the parent species. The holotype is thought to be a female with the type locality given as “Rio de Janeiro (Brasil)”.

Tribe CHRYSOBOTHRINI

Colobogaster weyrauchi Cobos, 1966

Colobogaster weyrauchi Cobos, 1966

Cobos (1966) described Colobogaster weyrauchi from Peru and named it after the collector, relating it to the widespread Colobogaster cyanitarsis Gory & Laporte, 1837. The type locality was given as “Pucallpá, 200 m. alt., Perú (W. Weyrauch coll. I-1948)”.

Subfamily AGRILINAE

Tribe CORAEBINI

Dismorpha grandis Cobos, 1990

Dismorpha grandis Cobos, 1990

Cobos (1990) described Dismorpha grandis from Argentina in his very last buprestid publication, stating that the species had the appearance of an enormous D. irrorata (Gory & Laporte, 1839) (thus, the name “grandis“). The holotype is a male with the type locality given as “Bemberg, Misiones, Argentina (Exp. Hayward-Willink-Golbach: 12-29-I-1945)”.

Tribe AGRILINI

Diadorina golbachi Cobos, 1974 (monotypic)

Diadorina golbachi Cobos, 1974 (monotypic)

Cobos (1974) described Diadorina golbachi from Argentina as the only member (and thus the type species) of the new genus Diadorina (the genus is still regarded as monotypic), naming it in honor of the collector. The holotype specimen is a female with the type locality given as “La Tigres, Santiago del Estero, Argentina (R. Golbaeh coll. 11-16-1-1970)”.

Tribe TRACHEINI

Pachyshelus huallaga Cobos 1969 (correct spelling is huallagus)

Pachyshelus huallaga Cobos, 1969

Cobos (1969) described and named this species after the river at the type locality in Peru. He related it to Pachyschelus atratus Kerremans, 1896 from Brazil and northern Argentina, stating that it differed by its distinct and less brilliant coloration and other features. Since the genus name is considered masculine, the correct species name is “Pachyschelus huallagus Cobos, 1969″. The unique holotype is a female with the type locality given as “Tingo María, Rio Huallaga, 700 metros de altitud, Peru, X-1946 (W. Weyrauch leg.)”.

Pachyschelus weyrauchi Cobos, 1959

Pachyschelus weyrauchi Cobos, 1969

Cobos (1969) described Pachyschelus weyrauchi from Ecuador and named it in honor of its collector. He related the unique male to Pachyschelus aeneicollis (Kirsch, 1873) from Peru and Bolivia, citing differences in coloration, body shape, and surface sculpture. The type locality was given as “El Puyo, 900 metros de altitud, Ecuador, 10-IV-1958 (W. Weyrauch leg.)”.

There are two additional Buprestidae type specimens in the collection (Colobogaster pizarroi Cobos, 1966 and Hylaeogena cognathoides Cobos, 1969), but they are in another drawer that we did not find immediately and, thus, I did not have a chance to photograph them. My apologies!

REFERENCES:

Bellamy, C. L. 2006. Nomenclatural notes and corrections in Buprestidae (Coleoptera). The Pan-Pacific Entomologist 81(3/4):145–158 [pdf].

Bellamy, C. L. 2008. A World Catalogue and Bibliography of the Jewel Beetles (Coleoptera: Buprestoidea). Volume 2: Chrysochroinae: Sphenopterini through Buprestinae: Stigmoderini. Pensoft Series Faunistica No. 77, pp. 626–1260, Pensoft Publishers, Sofia-Moscow [details & links].

Cobos, A. 1966. Notas sobre Bupréstidos neotropicales. XV: Tres especies nuevas de Colobogaster Sol. (Coleoptera). EOS, Revista Española de Entomología 41(2-3):205–214 [pdf].

Cobos, A. 1969. Notas sobre Bupréstidos neotropicales XVII. Especies y subespecies nuevas (Coleoptera). EOS, Revista Española de Entomología 44(1968):19–43 [pdf].

Cobos, A. 1958. Tercera nota sobre Bupréstidos (Ins. Coleoptera) neotropicales descripciónes y rectificaciónes diversas. Acta Zoologica Lilloana 15:83–102 [pdf].

Cobos, A. 1973. Revisión del género Tylauchenia Burm., y afines (Coleoptera, Buprestidae). Archivos del Instituto de Aclimatacion 18:147–173 [pdf].

Cobos, A. 1974. Notas sobre Bupréstidos neotropicales, XIX. El género Amorphosternus H. Deyrolle y afines. Archivos de Instituto de Aclimatación 19:65–81 [pdf].

Cobos, A. 1990. Revisión del género Dismorpha Gistel (Coleoptera, Buprestidae). Revista Brasileira de Entomología 34(3):539–559 [pdf].

Moore Rodriguez, T. 1986. Contribución al conocimiento de los Buprestidos neotropicales (Coleoptera: Buprestidae). Revista Chilena de Entomología 13:21–29 [BioStor].

Moore Rodriguez, T. & T. Lander. 2010. Revision du genre Conognatha. Edition Magellanes 24:1–172 [introduction and generic discussion in French and Spanish; keys to species in English, French and Spanish] [order information].

© Ted C. MacRae 2015

I got Thomas Shahan to image my Chrysochroa corbetti!

Chrysochroa-corbetti-TwitterThose who follow me on Twitter know that I attended Entomology 2014 last month in Portland, Oregon. As with other scientific conferences, live tweeting of the talks and associated events was all the rage. I may not have been the most prolific “tweeter”, but I did do my share, and one of my tweets involved a rather spectacular preserved specimen of the jewel beetle, Chrysochroa corbetti. The quick iPhone snapshot attached to the tweet was sufficient to prove that the beetle is pure eye candy, but still it did not do full justice to its stunning beauty:

Fortunately, while I was at the meetings I ran into Thomas Shahan—already an icon among insect macrophotographers for his seemingly impossible portraits of jumping spiders, tiger beetles, and other insects. I had planned to spend a couple of days in Salem after the meetings to visit my friend and longtime buprestophile Rick Westcott. As it happens, Thomas is currently a Digital Imaging Specialist at the Oregon Department of Agriculture where Rick spent his entire career as an entomologist. When I showed my specimen to Thomas, he kindly agreed to make some focus-stacked images of the specimen using his lab’s photographic setup. I think you can now agree that this is one of the most spectacular jewel beetles around, and I think you’ll also agree that these images by Thomas are perhaps the most stunning of this oft-photographed species. Be sure to check out the last photo—a 10× close-up of the dorsal elytral detail at the interface between the green and blue areas. Simply stunning!

Chrysochroa (Chroodema) corbetti (Kerremans, 1893) | Thailand

Chrysochroa (Chroodema) corbetti (Kerremans, 1893) | Thailand

Chrysochroa corbetti lateral view

Chrysochroa corbetti lateral view

Chrysochroa corbetti ventral view

Chrysochroa corbetti ventral v

Chrysochroa corbetti dorsal elytral detail (10X)

Chrysochroa corbetti dorsal elytral detail (10X)

© Ted C. MacRae 2014

Proof that it’s possible to ship large, pinned beetles safely!

Miscellaneous Buprestidae from Dan Heffern

Miscellaneous Buprestidae from Dan Heffern

Those who have followed this blog for awhile know that I’ve been on a bit of a rant during the past few months about the way pinned insect specimens are packed and shipped. This has been prompted primarily by the receipt of several damaged insect shipments, some of the more egregious examples of which are shown here and on Facebook. In all of these cases, damage could have been prevented had the specimens simply been packed and shipped using standard best practices.

I do not wish, however, to give the impression that every insect shipment I receive is damaged. The photo above represents a shipment I just received from Dan Heffern, who is kindly gifting to me some of the excess Buprestidae that he has in his collection in order to make room for his more beloved Cerambycidae. This shipment was especially prone to damage because of the number of large, heavy-bodied specimens it contains. Nevertheless, it arrived safe and sound because of the attention paid by Dan to securing the specimens in place. Note the liberal use of brace pins around each specimen—the larger the specimen, the more brace pins. In addition, the pinning box features a double-foam layer. Double-foam holds specimens much more securely in place than does a single layer, and while I didn’t mention it in my original post it’s a good idea for shipments containing large, heavy-bodied specimens. One drawback of double-foam is that it pushes labels on the pin up close to the specimen, but re-positioning labels on pins is certainly better than having to reattach broken body parts on specimens!

My thanks to Dan for this fine shipment and for paying such great attention to its packing to ensure receipt in the best condition possible!

© Ted C. MacRae 2014

Receiving a shipment of insects for identification…

…is like Christmas all over again!

Unopened shipping box

Unopened shipping box

The sight of a newly delivered box sitting outside my office brings on a rush of excitement. The sight of an enormous box is even more exciting. I know what’s inside is gonna be good, but I don’t know how good. Will there be rare species I haven’t seen before? Will there be specimens representing new (and, thus, publishable) state records or host associations? By the same token, the bigger the box, the more nervous I get. Shipping pinned insect specimens can be risky, and the potential for damage to the specimens increases as the size of the shipment increases—it all depends on how well they were packed (and a little bit of luck!). The prominent “Fragile” labeling, detailed description of the contents, and up arrow indicators are all good first signs.

Opened shipping box w/ paperwork

Opened shipping box w/ paperwork

I remain optimistic as I open the shipping box and see foam peanuts filling the box almost, but not completely, to the brim to allow a little bit of shuffle for shock absorption. The specimen boxes are also completely hidden under the top layer of foam peanuts, suggesting there is enough vertical clearance inside. Lastly, paperwork placed inside the shipping box and on top of the cushioning ensures that the shipment can be delivered even if the outer shipping label is damaged or lost.

Inner shipping boxes

Inner shipping boxes

Below the top layer of foam I find two inner shipping boxes. I am a little concerned by the lack of clearance between the inner shipping boxes and the sides of the outer shipping box—ideally there should be a foam-filled gap of at least a couple of inches to allow some lateral shock absorption. I am also concerned that the two inner shipping boxes are not also bound to prevent bumping against each other, although the lack of space between them and the outer shipping box probably makes this point moot.

Opened inner shipping boxes

Opened inner shipping boxes

Inside the inner shipping boxes are very nicely wrapped specimen boxes. I’m not sure the inner wrapping to cushion the specimen boxes from each other accomplishes all that much other than to increase the size of the inner shipping boxes, which in turn decreases the clearance between the inner and outer shipping boxes. I would have rather seen the specimen boxes bound tightly together into a small unit to have additional space between them and the outer shipping box.

Unopened specimen boxes

Unopened specimen boxes

Seven classic insect specimen shipping boxes—the excitement (and nervousness) mounts as I prepare to open them and get my first look at the enclosed specimens.

Opened specimen boxes

Opened specimen boxes

A fine selection of gorgeous jewel beetles—mostly from Colorado but with a good number of specimens collected from countries around the world. But uh-oh, no inner false lids! A false lid rests directly on top of the pins of the specimens inside and is held in place by cushioning between the false and true lids. False lids are essential in shipments of any size to keep the pinned specimens, especially heavy-bodied ones, from working their way loose from the foam and bouncing around inside the specimen box during shipment. Fortunately, all of the specimens stayed put in most of the specimen boxes, …

Shipping damage

Shipping damage

…but one or two of the really heavy-bodied specimens did work their way loose in a couple of the boxes. As a result, there was some minor damage in the form of broken tarsi and antennae. The damage, however, is not great, and with fine-tipped forceps and a little bit of clear finger nail polish I should be able to effect decent (if not perfect) repairs. To the shipper’s credit, they made extensive use of brace pins on each side of heavier-bodied specimens in all of the boxes—this probably served to keep the damage as minimal as it was.

Although I salivate looking at the specimens—nearly 800 in all, I must set aside my desire to dive right into them and turn my attentions back to a previously received shipment (also numbering in the several hundreds). As soon as I finish that shipment, I’ll start working on this one, but I suspect that while I’m working on it I will receive another shipment that, like this one, competes newly for my attentions.

Copyright © Ted C. MacRae 2014