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

Posted in Buprestidae, Coleoptera, Fabaceae, Scarabaeidae | Tagged , , , , , , , , | Leave a comment

2015 Texas Collecting Trip iReport—Fall Tiger Beetles

This is the fourth in a series of “Collecting Trip iReports”—so named because I’ve illustrated them exclusively with iPhone photographs. As I’ve mentioned in previous articles in this series (2013 Oklahoma2013 Great Basin, and 2014 Great Plains), I tend to favor my iPhone camera for general photography—i.e., habitats, landscapes, miscellaneous subjects, etc.—during collecting trips and save my full-sized dSLR camera only for those subjects that I want high-quality macro photographs of. iPhones are not only small, handy, and quick but also capable (within reason) of quite good photographs (see this post for tips on making the most of the iPhone camera’s capabilities). This keeps the amount of time that I need to spend taking photos at a minimum, thus allowing more time for the trip’s intended purpose—collecting! Those photos form the basis of this overall trip synopsis, while photos taken with the ‘real’ camera will be featured in future posts on individual subjects.

Last year during late September and early October I travelled to eastern and central Texas. This trip was all about fall tiger beetles, in particular certain subspecies of the Festive Tiger Beetle (Cicindela scutellaris) and Big Sand Tiger Beetle (Cicindela formosa) found in that area that I had not yet seen. I enjoy all collecting trips, but fall tiger beetle trips are among the most enjoyable of all—cooler temperature, a changing landscape, and charismatic subjects that are both fun and challenging to find and photograph. This trip was no different, with spectacular weather during the entire week and, for the most part, great success in finding the species/subspecies that I was after. At this point I’d like to acknowledge the help of several people—David Hermann (Ft. Worth, Texas), David Brzoska (Naples, Florida), and Steve Spomer (Lincoln, Nebraska), who generously provided information on species and localities. My success at finding these beetles was due in large part to the information they provided.


Day 1 – Cobb Hollow

My car

Little question about what I am doing out here.

After driving 700 miles from my home near St. Louis, I arrived at the first stop of trip—Cobb Hollow in north-central Texas. This small creek lined with deep, dry sand is close to Forestburg (Montegue County)—the type locality of Cicindela scutellaris flavoviridis, a beautiful, all-green subspecies with the elytra suffused golden-yellow.  The habitat looked very promising from the start, and it wasn’t long before I found the first tiger beetle of the trip—a gorgeous, red nominate Big Sand Tiger Beetle (Cicindela formosa formosa). Not long after that I found the first Cicindela scutellaris flavoviridis, and over the next few hours I would find a total of nine individuals. Despite the extensive habitat along the creek the beetles were quite localized, occurring primarily in two dry sand areas within a mile west of the bridge. This spot is actually near the northern limit of the subspecies’ distribution, and several of the individuals showed varying influence from nominate scutellaris with the elytra tending to be more red than yellow-green. There was a diversity of other tiger beetles here as well—C. formosa formosa was the only one that was common, but I did find also a few individuals each of Tetracha carolina, Cicindelidia punctulata, Cicindela splendida, and C. repanda. A very cool place.

Cobb Hollow from bridge

View of Cobb Hollow east from the bridge

Sand bar along creek

Dry sand deposits line the creek.

Robber fly with bumble bee prey

I watched this robber fly snag a bumble bee in mid-flight.

Ted MacRae at Cobb Hollow

Looking down onto the creek from the bridge.


Day 2 – Stalking the Limestone Tiger Beetle

Today was all about looking for the Limestone Tiger Beetle, Cicindelidia politula. I have collected this species previously at several sites in Erath and Somervell Counties, Texas (west of Ft. Worth) and featured photographs from that trip. However, since I would be passing through the area on my way south I decided to spend a day looking for it again and, hopefully, collecting a few more specimens. Cicindelidia politula is related to the much more common and widespread Punctured Tiger Beetle, C. punctulata, but is shiny blue-black with the elytral markings absent or limited to the apices and the abdomen red. I visited several localities—two new ones for me in Erath County and another I had visited previously in Somervell County, with habitats that ranged from rocky clay to white limestone exposures along roadsides and even limestone gravel.

I found a fair number of individuals at the first site (1.7 mi SW Bluff Dale, Jct US-377 & FM-1188), which had a finely ground limestone substrate. Most of the individuals were flushed from the base of clumps of bunch grass and captured when they landed in more exposed situations.

Limestone habitat for Cicindelidia politula

Cicindelidia politula habitat—1.7 mi SW of Bluff Dale.

The beetle had also been reported along the roadsides at the second location (0.4 mi E Jct FM-2481 on CR-539), but the only individual I saw here was on a very coarse crushed limestone 2-track leading off of the main road.

Limestone habitat for Cicindelidia politula

Cicindelidia politula habitat—0.4 mi E Jct FM-2481 on CR-539.

The species was most numerous at the third site in Somervell County (3.4 mi SE Jct US-67 on CR-2013). I collected ten individuals and saw probably that many more on white limestone exposures along the roadside and along a dirt road cut along the base of the hill to the NE side of the highway. Most of the beetles in the latter area were seen along the scraped dirt road (at left in 2nd photo below), although presumably the beetles also utilized the undisturbed, surrounding habitat.

Limestone habitat for Cicindelidia politula

Cicindelidia politula habitat on white, limestone exposures along the roadside.

Limestone habitat for Cicindelidia politula

Cicindelidia politula habitat on white limestone hillside and scraped dirt road.

Catching the beetles at this last locality was challenging—the adults are fast and flighty, and the rough, rocky habitat made it difficult to clamp the net over the beetle and pounce on top of the rim before they were able to find a gap and escape. With practice I found my catch efficiency increased a little bit if I slowly approached the beetle and then made an assertive swing with the net right when the beetle began to fly—the trick is learning how to tell when they are ready to fly (and “assertive” is the key word!). Tiger Beetle Stalker; however, does not quit!

Tiger beetle stalker!

Tiger Beetle Stalker!


Day 3 (Part 1) – Pedernales Fall State Park

This was another locality where Cicindela scutellaris flavoviridis had been recorded. I came here to find this subspecies even though I had seen it two days previously at Cobb Hollow, because that latter population showed some slight intergradation of characters from nominate C. scutellaris and I wanted to get field photographs of a “pure” population. I was pretty excited when I saw extensive dry sand habitat lining the upper bank area along the Perdenales River; however, I found no tiger beetles of any kind after extensive searching through that habitat. I did note the area seemed dry and reasoned that perhaps timely rains had not yet triggered emergence of C. scutellaris, C. formosa, and other sand-loving fall tiger beetles. I did find a small area of wet sand right along the water’s edge where three species of Cicindelidia could be seen: C. ocellata rectilatera, C. trifasciata ascendens, and C. punctulata. I’ve photographed all of these species before, so I didn’t try to spend any time doing so here. However, combined with the species seen the previous two days, this made a total of ten species seen on the trip so far. Although I didn’t find the beetle I was looking for, I marveled at the beauty of the area, especially the Pedernales River with its hard, conglomerate bedrock and mini shut-ins and spent quite a bit of time here taking photographs.

Perdenales River

The Perdenales River is the centerpiece of the state park.

Schistocerca americana or nitens

Schistocerca americana or S. nitens (ID courtesy of Matt Brust).

Perdenales River

Shut-ins are extensive along the Perdenales River.

Poecilognathus sp.

Bee flies (family Bombyliidae), prob. Poecilognathus sp. (ID courtesy Rob Velten).


Day 3 (Part 2) – Lick Creek Park

Another of the Festive Tiger Beetle subspecies that I wanted to look for was Cicindela scutellaris rugata. I had several localities from which this solid blue-green subspecies has been recorded, and this site was the nearest of those that I planned to visit. The drive from Pedernales State Park was longer than I anticipated, so I didn’t get to this spot until close to 6 p.m. At first I worried that I wouldn’t have enough time to even find suitable habitat, but that was no problem as I quickly found the Post Oak Trail and its perfect open, post oak woodland with deep sand substrate. By all accounts the beetles should have been all over the trail but they weren’t. As with the previous site, the area was quite dry as evidenced by the wilted plants along the trail side, and I also note that the previous record from here was on Oct. 23rd—more than three weeks later. Despite the fact that I didn’t find any tiger beetles, I did see a young timber rattle snake (Crotalus horridus) crossing the trail late in the hike—I took a quick shot with the iPhone (see below) and then broke out the big camera and was able fire off a few shots before it left the trail and headed for cover. (Several people walking the trail came upon us, and they were all—happily—more than willing to oblige my requests to stay away until I was finished.)

Sand woodlant habitat for Cicindela scutellaris rugosa

Post oak woodland with dry sand substrate seems to be perfect for Cicindela scutellaris rugata.

Wilted American beautyberry (Callicarpa americana)

Wilted American beautyberry (Callicarpa americana).

Timber rattlesnake (Crotolus horridus)

A youngish (prob. ~32″ in length) timber rattlesnake (Crotolus horridus) was a treat to see.


Day 4 – East Texas cemeteries

Cemeteries are often great places to look for tiger beetles because they tend to be located on parcels of land with low agricultural value that were donated by landowners to local churches. Older cemeteries especially tend not to be highly maintained and, thus, offer excellent habitat for tiger beetles. My goals for this day were Cicindela scutellaris rugata and the gorgeous Cicindela formosa pigmentosignata. I had records of both from a couple of cemeteries in eastern Texas (Sand Flat Pioneer Cemetery in Henderson and Morris Chapel Cemetery in Van Zandt Counties) and found good numbers of both along sandy 2-tracks and sparsely to moderately vegetated sand exposures in and around the cemetery grounds. I don’t have any iPhone photographs to share of either of these species, but I did spent a lot of time with the big camera and got a number of photos of each that I am quite pleased with—I’ll share those in future posts. The cemeteries themselves were haunting and poignant, with some headstones dating back to the late 1800s.

Sandy 2-track habitat for Cicindela scutellaris rugata & C. formosa pigmentosignata

Sandy 2-track habitat for Cicindela scutellaris rugata & C. formosa pigmentosignata at Sand Flat Pioneer Cemetery, Henderson County, Texas.

 

Ant mound

Pogonomyrmex sp. poss. barbatus tend their nest entrance (ID courtesy of Ben Coulter).

Sand Flat Pioneer Cemetery

Oldest section of Sand Flat Pioneer Cemetery.

Died Nov 10, 1874

Fallen, but not forgotten—yet (died Nov 10, 1874).

Oldest headstones (late 1800s)

Oldest headstones (late 1800s) at rest under the shade of huge, red-cedar trees.

Oldest person (106 yrs old)

The oldest person died at 106 years of age (born in 1804).

At Morris Chapel Cemetery I found C. formosa pigmentosignata and C. scutellaris rugata on sparsely vegetated deep dry sand 2-track north of the cemetery. I did also manage to get field photos of the former before it got too hot and they became too active. There were also a few of the latter in the open sandy ground just outside the northwestern edge of the cemetery. As with Sand Flat Pioneer Cemetery, I spent a bit of time in the cemetery proper to look at the headstones—the oldest headstone also being the most poignant; a one and a half-year old boy who died in 1881.

Sandy 2-track habitat for Cicindela scutellaris rugata & C. formosa pigmentosignata

Sandy 2-track habitat near Morris Chapel Cemetery.

Morris Chapel Cemetery

A large, spreading post oak shades pioneers at rest.

Died 1881 (age 1½ yrs)

A poignant headstone (died 1881 at 1½ years of age).

After finishing up at Morris Chapel Cemetery I returned to Sand Flat Cemetery to see if I could get more field photographs before the beetles bedded down for the night. The sun was still up when I arrived a little before 6 p.m., but the shadows were long and no beetles were seen. Not one to waste an opportunity, I broke out the big camera anyway and started photographing a large species of bee fly (family Bombyliidae) that was perching on the ground and on the tips of plains snakecotton (Froelichia floridana).

Undet. bee fly

Bee fly (family Bombyliidae), poss. Poecilanthrax lucifer? (ID courtesy Alex Harman).


Day 5 (Part 1) – Cowtown Bowman Archery Club

With both specimens and good field photos of Cicindela scutellaris rugata and C. formosa pigmentosignata in hand, I returned my attention to C. scutellaris flavoviridis. Again, I did already have specimens in hand from Cobb Hollow, but most of them showed some degree of intergradation with nominate C. scutellaris and I was hoping to see some “pure” individuals. Failing to find it at the more southerly locations (Pedernales State Park and Lick Creek Park), I had one more location in Tarrant County where the subspecies had been recorded—a sand borrow pit near the entrance of Cowtown Bowman Archery Club. Once again I searched the area thoroughly for a couple of hours during mid-morning but did not see the subspecies or any other tiger beetles. Conditions were overcast and cool (72°F), but I do not think this explains the absence of adults. Rather, I think I was on the early side of the season and they just hadn’t started emerging at this site.

While I was at the site I found several tiger beetle larval burrows in a moderately vegetated area near the deeper sand deposits that were occupied by Tetracha carolina, so I used the “stab” or “ambush” method to collect several 3rd instars for an attempt at rearing. For those of you who are not familiar with this technique, a knife is set at a 45° angle with the tip in the soil about 1″ from the edge of the burrow. Then you wait, sometimes for quite a while, until the larva reappears at the top of the burrow and STAB the knife assertively into the soil to block the larva from retreating. The larvae are extremely wary with excellent vision and will usually drop back down immediately when they see you, so you have to be ready and act quickly. Once the retreat is blocked, a simple twist of the knife to expose the larva is all that is needed. I prepared larval habitats by placing native soil with as intact a top layer as possible in plastic critter carriers, made a starter hole for each larva with a pencil, dropped each larva into one of the holes, and then pushed the soil to seal the burrow entrance. This prevents the larvae from crawling right back out of the starter burrow, which can result in them encountering and fighting each other. The larvae will eventually reopen the burrow entrance, but after being sealed inside for a while they usually accept the burrow and further modify it to suit their needs.

 

Sandy grassland habitat for Tetracha prob. carolina

Sandy grassland habitat for Tetracha carolina.

Larval burrows (lower left) can be recognized by their clean, almost perfectly round, beveled edge. The presence of fresh soil diggings cast to one side (upper right) indicates the burrow is occupied by an active larva.

Tetracha prob. carolina larval burrow

Tetracha carolina larval burrow with cast soil diggings.

Using the “stab” or “ambush” method to collect larvae. One must have patience to successfully use this method.

"Stab 'n; grab" method to collect tiger beetle larvae (Tetracha prob. carolina)

Using the “stab” or “ambush” method to collect tiger beetle larvae.


Day 5 (Part 2) – Cobb Hollow (epilogue)

Although I had found Cicindela scutellaris flavoviridis at this site on the first day of the trip, I had not taken any field photographs in hopes of finding a more “pure” population at one of the more southerly locations. That did not happen, so I returned to Cobb Hollow on this last day in the field to get field photographs from the population there. Temperatures were a bit cooler (mid-70s) and cloud cover was variable, actually sprinkling when I arrived mid-afternoon but eventually clearing. This seemed to have no detrimental effect on adult presence, and it may have actually helped as I was able to photograph the very first individual that I found to my heart’s content. I collected that individual and the next three that I saw by hand and found two more over the next hour—all on the same deep, dry sand bars west of the bridge where I had seen them previously. Curiously, Cicindela formosa was strangely absent from these same areas where they had been so numerous a few days earlier.

Habitat for Cicindela formosa formosa and C. scutellaris flavoviridis

Deep, dry sand deposit where most of C. scutellaris flavoviridis were seen.

On the east side of the bridge I collected two more Tetracha carolina in the same moderately vegetated sandy clay spot as last time, then went on to the furthest dry sand bar where I found and photographed (but did not collect) a single C. formosa (only one shot before it took off). I also found a female green lynx spider (Peucetia viridans) sitting on her egg mass and got some nice macro photos as well as this iPhone shot (talk about a face only a mother could love!).

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Female green lynx spider (Peucetia viridans) atop her egg mass.


I hope you’ve enjoyed this collecting trip iReport. Stay tuned for true macro photographs of the tiger beetles and other insects/arthropods that I photographed on this trip in more subject-specific posts. You are also welcome to leave feedback in the comments below.

Ted MacRae w/ field collecting equipment & camera

© Ted C. MacRae 2016

Posted in Acrididae, Arachnida, Araneae, Asilidae, Bombyliidae, Cicindelidae, Coleoptera, Diptera, Formicidae, Hymenoptera, Orthoptera, Reptilia, Vertebrata | Tagged , | 7 Comments

Super duper June bugs

Last June, after spending the day collecting insects at Sand Hills State Park in south-central Kansas with Mary Liz Jameson, Jeff Huether and I setup our blacklights at the edge of the dunes. We were hoping to attract males of the genus Prionus, following a hunch that maybe the dunes—a popular historical collecting site—would prove to be the habitat for the enigmatic Prionus simplex (known only from the type specimen labeled simply “Ks.”). We knew it was a long shot, made even longer by a bright moon and the unseasonably cool temperatures that settled over the dunes as the sun dipped below the horizon, and in the end no Prionus would be seen. We did see, however, some other interesting insects, one of the more interesting being males of Hammond’s lined June beetle—Polyphylla hammondi. Almost immediately after sunset a number of these large, chunky-bodied beetles resembling super-sized versions of their far more diverse and commonly encountered relatives in the genus Phyllophaga (May beetles) began arriving at the lights—each one noisily announcing its visit by its loud, buzzing, flight and bumbling thud onto the ground nearby.

Polyphylla hammondi

Polyphylla hammondi | Sand Dunes State Park, Kansas

I’ve encountered beetles in the genus Polyphylla only occasionally over the  years, almost always at night as a result of their attraction to lights. The genus is most diverse in the southwestern U.S., and many species are found only in specific sand dune habitats (Young 1988, LaRue 1998). Their large size, relatively more restricted distributions, and less common occurrence make them interesting enough, but what made this encounter particularly interesting for me was the way the beetles—all males—held their fan-like antennae splayed out. Male Polyphylla have greatly enlarged antennae that they use to detect sex pheromones emitted by the female (Lilly and Shorthouse 1971). Many female Polyphylla are flightless, especially those restricted to sand habitats, and are rarely collected, and for some species they still remain unknown. In fact, the best way to find females is to listen for the sound of the males hitting the ground or vegetation once they have located a female (Skelley 2009).

Polyphylla hammondi

Male with antennae splayed to detect female pheromone.

It was clear to me that these males were actively searching for females. The greatly elongated antennomeres provide lots of surface area for sensory pores to detect female pheromones at low concentrations. I’d not seen this before and didn’t know how long it would last—many beetles have narrow windows of activity for mating that can be affected or restricted further by environmental cues such as temperature. I figured I’d better get some photographs on the spot while I could, and this was a smart decision as it wasn’t too long after I took these photos that the males stopped coming to the lights and those that were already there became inactive and no longer held their antennae so impressively splayed.

Polyphylla hammondi

Males cease activity after sunset.

In a recent paper describing a new western species of the genus, La Rue (1998) provided detailed notes on behavior that probably pertain to other sand dune inhabiting species as well. Males were observed to begin flying in late afternoon, making rapid, irregular flights several meters above the sand surface. However, as dusk approached, their flights became less erratic and more purposeful as they flew rapidly upwind and then returned in a slow zig-zag flight (indicative of osmoclinotaxic orientation). Mating occurred after they located a female sitting on the sand and alighted within a few centimeters of her. Several males were attracted to each female, further supporting the use of pheromones by the female to attract males. Males were also attracted abundantly to lights after dusk and ceased activity shortly to several hours after sunset, presumably because females cease releasing pheromone to attract them and burrow back into the sand.

REFERENCES:

LaRue, D. A. 1998. Notes on Polyphylla Harris with a description of a new species (Coleoptera: Scarabaeidae: Melolonthinae). Insecta Mundi 12(1–2):23–37 [pdf].

Lilly, C. E. & J. D. Shorthouse. 1971. Responses of males of the 10-lined June beetle, Polyphylla decemlineata (Coleoptera: Scarabaeidae), to female sex pheromone. The Canadian Entomologist 103:1757–1761 [abstract].

Skelley, P. E. 2009. A new species of Polyphylla Harris from peninsular Florida (Coleoptera: Scarabaeidae: Melolonthinae) with a key to species of the pubescens species group. Insecta Mundi 0085:1–14 [pdf].

Young, R. M. 1988. A monograph of the genus Polyphylla Harris in America north of Mexico (Coleoptera: Scarabaeidae: Melolonthinae). Bulletin of the University of Nebraska State Museum 11(2):vi+115 pp. [BioQuip preview].

© Ted C. MacRae 2016

Posted in Coleoptera, Scarabaeidae | Tagged , , , , , , , | 9 Comments

North America’s most recognizable longhorned beetle

One of the more impressive insects that we found during our visit to Sand Hills State Park in south-central Kansas last June was Plectrodera scalator, the cottonwood borer. Large and robust (in fact, the only larger species in the family are the prionid root borers and their kin), their striking checkered pattern of white pubescence on a glossy black body makes them perhaps the most recognizable of all North American longhorned beetles (Linsley & Chemsak 1984). The very robust body of this individual, along with the relatively shorter antennae (only about as long as the body) identify it as a female—males are generally smaller and less robust with the body slightly tapering and the antennae distinctly longer than the body.

Plectrodera scalator

Plectrodera scalator (Fabricius, 1792) | Sand Hills State Park, Kansas

The white coloration on the body of these beetles is not a cuticular pigment (which is rather rare in beetles and is most often associated with species found in white sand habitats, e.g., certain tiger beetles), but instead a result of dense mats of microscopic white setae. The patterns formed by these mats are apparently as unique to each individual as fingerprints are to humans (Yanega 1996), making these beetles at once immediately recognizable as a species yet distinctive as individuals.

Plectrodera scalator

Adults of this species are found most often on cottonwood.

These are said to be common beetles in their range across the eastern two-thirds of the country, especially so in the Great Plains where their favored host, cottonwood (Populus deltoides), is especially abundant. Despite this, I have encountered this species only a handful of times in more than 3o years of searching. I know they’re out there, even in my home state of Missouri where I recorded 154 specimens collected in the state and deposited in various collections (MacRae 1994). It was not until around 2000 that I even saw my first ones (on a cottonwood tree in a homeowner’s yard just across the Mississippi River in Illinois), and in fact this one was actually found by Mary Liz Jameson, who had accompanied us to the field that day. It makes me wonder if their coloration, so strikingly conspicuous when isolated against a clean, blue sky background, might actually afford some type of cryptic protection against the normal backdrop of foliage and branches on which they are normally found—a phenomenon that I call “conspicuous crypsis” and which I have noted for other longhorned beetles (e.g., Acanthocinus nodosus). Perhaps, with this species at least, I have not yet set my search image to notice them.

Plectrodera scalator

Large, robust size and a distinctive checkered pattern of black and white makes these beetles among the most recognizable longhorned beetles in North America.

REFERENCES:

Linsley, E. G. and J. A. Chemsak. 1984. The Cerambycidae of North America, Part VII, No. 1: Taxonomy and classification of the subfamily Lamiinae, tribes Parmenini through Acanthoderini. University of California Publications in Entomology 102:xi + 1–258. [preview]

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]

Yanega, D. 1996. Field Guide to Northeastern Longhorned Beetles (Coleoptera: Cerambycidae). Illinois Natural History Survey Manual 6, x + 174 pp. [preview]

Posted in Cerambycidae, Coleoptera | Tagged , , , , , , , | 18 Comments

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

Posted in Cerambycidae, Coleoptera, Disteniidae | Tagged , , , , , , , , , | 34 Comments

North America’s most “extreme” jewel beetle

When Chuck Bellamy passed away two years ago, he left behind a remarkable legacy of study on the family Buprestidae (jewel beetles) that includes not only his insect collection—surely one of the best in the world in terms of representation of genera and species in the family—but also his extensive library of primary literature. Both of these assets, built over a period of decades, are now housed in the California State Collection of Arthropods at the CDFA Plant Pest Diagnostics Laboratory in Sacramento, California. Chuck, however, was not just a jewel beetle collector and taxonomist—he was also a skilled photographer, focusing (pun intended) largely, though not exclusively, on his beloved jewel beetles. Digital cameras were still far in the future when Chuck began photographing these beetles, and as a result the bulk of his photographic legacy exists in the form of 35mm slides. I was the fortunate recipient of his slide collection, numbering in the thousands, and have been slowly scanning his slides into digital format with the goal to eventually make them available to the larger community of buprestid workers. Some of his best photos were published in a memorial issue of The Coleopterists Bulletin (2014, volume 68, number 1), and I featured a few additional photos in this post shortly before the publication of that issue. There remain slides, however, of many additional species, a large number of which surely represent the only field photographs of live adults. As I convert his slides to digital format, I hope to share some of the more interesting here.

For the first of these featured species, I can think of no better one than Lepismadora algodones. This tiny little jewel beetle is the only representative of the genus, which was not even known until 1986 when it was discovered by Mimi & Rob Velten in the Algodones Sand Hills of southeastern California. The species and genus were described the following year (Velten & Bellamy 1987), making Lepismadora the most recently discovered new genus of jewel beetle in the U.S. The recentness of its discovery is remarkable, since southern California in general and the Algodones Sand Dunes in particular were thought to have been relatively well collected at the time of the beetle’s discovery. Also remarkable is the distant relationship of this monotypic genus to any other North American species; its closest known relative being the genus Eudiadora—known only from Argentina (Bellamy 1991).

Lepismadora algodones

Lepismadora algodones Velten, in Velten & Bellamy, 1987 (Coleoptera: Buprestidae)

Even more remarkable, however, are its highly localized distribution and extreme habitat. The entire type series (one male holotype and 159 paratypes) and all individuals collected since its description have been found only in a single old canal on the west side of the Algodones dunes. Summer temperatures in the dunes routinely reach in excess of 110°F and are even higher in the depressed canal where the beetles are found. Astoundingly, the adults are active only during the hottest hours of the day (ca. 10 a.m. to 2 p.m.), during which time they can be found on the flowers and foliage of fanleaf crinklematTiquilia plicata (Boraginaceae). The reason for the beetle’s highly restricted distribution is a mystery, as the plant on which the beetles are found is rather widespread across the southwestern U.S. and northwestern Mexico. A final mystery is the still unknown larval host plant—it could be T. plicata, but it could just as likely be something completely different.

Algodones Dunes

Old canal on the west side of Algodones Sand Hills, type locality of Lepismadora algodones.

I moved to California a few years after the species was described and, of course, soon set out to find it for myself. I had driven to southern California from my home in Sacramento to meet the late Gayle Nelson (another important mentor of mine), who told me where to find the beetle and what the host plant looked like but also warned me about the extreme heat I would encounter. His advice was to hike the canal until I had half a bottle of water, then turn around and hike back. Mindful of his advice, I arrived at the dunes the next day around mid-morning, filled my water bottle and hydrated myself as much as I could, and climbed down into the canal. The heat was overpowering—more so down in the canal and far beyond anything I had ever experienced to that point, and after quickly recognizing the host plants I began tapping their tiny, prostrate branches over my beating sheet and looking for the beetles. I went as far as I could down the canal, perhaps 200 yards, before I had to turn around, but I had not yet seen any beetles and was starting to lose hope. I continued to tap host plants on the way back, though by then not really expecting to see anything. About halfway back I saw something laying on the ground a short distance ahead. As I approached I saw it was a small plastic vial with a white cap, and when I picked it up I saw inside a dried out T. plicata twig and a dead adult beetle—unmistakably L. algodones! While excited to have found the species, it was at the same time a bit unsatisfying for the specimen to be one that somebody else had collected before me and then lost (for all I know, it could have been Chuck Bellamy, considering that the beetle was apparently intended to be kept alive, possibly for photography!). I slipped the vial into my pocket, started tapping branches again, and found three additional adults in the immediate vicinity of where I had found the vial (and doing much to soothe my dissatisfaction with the first specimen). Those would be the only specimens that I would find that day, though I would succeed in finding another individual on a subsequent visit two years later.

REFERENCES:

Bellamy, C. L. 1991. A revision of the genus Eudiadora Obenberger (Coleoptera: Buprestidae). Proceedings of the Entomological Society of Washington 93(2):409-419 [Biodiversity Heritage Library].

Velten, R. K. & C. L. Bellamy. 1987. A new genus and species of Coroebini Bedel from southern California with a discussion of its relationships in the tribe (Coleoptera, Buprestidae). The Coleopterists Bulletin 41(1):185–192 [pdf].

© Ted C. MacRae 2015

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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

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