Agrilus fuscipennis on Persimmon

Agrilus fuscipennis may not be the largest or the prettiest member of the genus occurring in Missouri (that honor is reserved for Agrilus concinnus, or “hibiscus jewel beetle”—MacRae 2004). Nevertheless,  it comes pretty darned close! Add to that the fact that it is among our most seldom encountered jewel beetles, and you can understand how excited I was to see this species on my sheet after beating a small persimmon (Diospyros virginiana) tree last weekend at Hercules Glades Wilderness in the White River Hills of extreme southwestern Missouri. In fact, I have only collected this species three times previously—all single specimens beaten from persimmon, and all back in the 1980s!

Jewel beetles are unquestionably popular among insect collectors, due no doubt in large part to their vivid, metallic colors. I think the family, however, would be even more popular were it not for the genus Agrilus. Fully one in five species of jewel beetles belongs to this genus, which at nearly 3,000 described species and counting (Bellamy 2008) is perhaps the largest genus in the entire animal kingdom. As might be expected, such hyperdiversity has resulted in taxonomic quagmire, with species limits difficult to define and many hardly distinguishable except by examination of male genitalia (MacRae 2003). Additionally, in contrast to the rest of the family which is generally recognized as containing some of the most spectacularly beautiful beetles in the world, the most species of Agrilus are small, usually less than 8 mm in length and often as small as only 4–5 mm, and also lack the vivid colors (at least, to the naked eye) for which the rest of the family is so noted.

Agrilus fuscipennis is one of several species that buck this general Agrilus theme. While not forming a discrete taxonomic group within the genus, they are all unified by the following characteristics: 1) relatively large for the genus (A. fuscipennis measures 12–14 mm length), 2) vivid red pronotum and black elytra, and 3) mine the lower trunks, crown and main roots of living rather than dead host plants. For A. fuscipennis the larval host is persimmon, and other similar species include A. vittaticollis on serviceberry (Amelanchier) and A. concinnus on wild hibiscus (Hibiscus). These other species also are not very commonly encountered, at least in my experience, perhaps partly because they are not as easily reared from their hosts as species that develop as larvae in dead wood (the latter can be easily reared by retrieving infested wood from the field and placing in containers to trap emerging adults).

In the interest of full disclosure, these photos were taken in the studio after returning home. Although the persimmon branch is real, the “blue sky” is actually just a colored index card. I prefer to photograph insects in the field, especially with insects such as tiger beetles where it is desirable to include elements of the insect’s natural habitat in the photograph. However, I don’t have a problem with studio photography if field photographs prove too difficult or time-consuming or present too high a risk of escape by a prize specimen. My normal protocol for the latter is to place the first individual in a vial and continue to search for another that I will then try to photograph in the field. If that doesn’t work then I still have the first individual as a backup for studio photographs. In the case of this beetle, I found it on the very first clump of persimmon that I beat but never saw another despite beating persimmon for the rest of the afternoon (just like the three I found separately back in the 80s)! I have plans to photograph A. concinnus later this summer on its Hibiscus host plant in southeastern Missouri—hopefully I will succeed in getting true field photographs of that species.

REFERENCES:

Bellamy, C. L. 2008. World catalogue and bibliography of the jewel beetles (Coleoptera: Buprestoidea), Volume 4: Agrilinae: Agrilina through Trachyini. Pensoft Series Faunistica 79:1–722.

MacRae, T. C. 1991. The Buprestidae (Coleoptera) of Missouri.  Insecta Mundi 5(2):101–126.

MacRae, T. C. 2004. Beetle bits: Hunting the elusive “hibiscus jewel beetle”. Nature Notes, Journal of the Webster Groves Nature Study Society 76(5):4–5.

Copyright © Ted C. MacRae 2013

Oversized, double-concave diffuser for MT-24EX twin flash

Megaloxantha purpurascens peninsulae, photographed with oversized, double-concave diffuser

This jewel beetle is, of course, Megaloxantha purpurascens peninsulae from Palawan, Philippines. I say “of course” because I’ve posted images of this same beetle on several occasions while testing out different diffuser designs for my Canon MT-24EX twin flash unit. In the most recent one, I had tried combining SoftBoxes with my oversized concave diffuser and was pleased enough with the result that I thought I might try it in the field. Well, let’s just say the extensions for the flash heads and SoftBoxes attached to them was far too clumsy for field use, and I abandoned the idea after just a couple of hours. Back to the drawing board.

Despite the problems with using the SoftBoxes in the field, I still wasn’t ready to give up on the idea of double diffusion, and I had also learned that extending my oversized diffuser out over the subject (leaving it “open”) produced better lighting than curling it back (as I had been doing). Curling the diffuser back only served to turn it into a convex diffuser, which results in more specular highlighting because the center of the diffuser is closer to the subject than the edges. A concave diffuser provides more even lighting because all parts of the diffuser are roughly the same distance from the subject. Just about that time, I saw a DIY diffuser design by Piotr Nascrecki that, in principle, resembled Alex Wild‘s tent diffuser. It was, however, much larger—like mine, and thus amenable for use with a 100mm macro lens (the macro lens I use most commonly). This resemblance to Alex’s diffuser did make me notice one missing feature—double diffusion layers. That’s when I thought, why not do the same with an oversized diffuser rather than fussing with separate diffusers attached to the flash heads? I had some Bogen Imaging filter sheets on hand (#129 Heavy Frost), so I picked up some 1-mm steel wire at the hardware store, found a Bic pen in the drawer that I could cut in half, and built the diffuser as shown in Piotr’s post. I then secured a second filter sheet above the first sheet by taping the two together along their sides, being sure to ‘bow’ the upper sheet above the bottom sheet to achieve the double diffusion effect. Here is the result (please excuse the iPhone shots):

Canon 50D with MT-24EX twin flash and oversized, double-concave diffuser.

Better view of the double diffusion layers and Piotr’s “Bic pen” attachment system.

I have big hopes that this will finally be the diffuser I’ve been looking for. For as quick a test shot as the jewel beetle photo above was, the lighting is great and the colors are vibrant—both achieved with typical post-processing. My only complaint is the slightly greater “hot spot” intensity in the lower parts of the highlights in the eyes. This is due to the flash heads sitting near the base of the diffuser, and (as Piotr recommends) a second set of Kaiser shoes will allow me to move the flash heads not only more towards the center of the diffuser but also further above it to help spread out the light throw and even out the highlights. I’ll need to play around positioning the flashes to figure out the best positions depending on the size and distance of the subject—sitting up higher as they are puts them more on “top” than in “front” of the subject, so they will need to be directed downward more than I am used to doing. Even more important, however, is field usability, and I really think this diffuser will prove to be convenient and easy to use in the field—no more gawky arms attached to the camera, the diffuser attaching quickly and easily and, just as importantly, coming off easily and storing flat in the backpack, and large enough to do the job while not so oversized that it gets in the way. Piotr says this diffuser also works well with the 65mm macro lens, so I will certainly be testing that out as well.

Copyright © Ted C. MacRae 2013

A jewel of a beetle

I really wish I had a photomicrography setup like the one that Sam Heads has at the University of Illinois for imaging preserved specimens. Alas, insect taxonomy is “just a hobby” for me, and any specimen photography I wish to do must be done with my field camera equipment. Of course, poverty prompts creativity (not that I consider a Canon 50D with an MP-E 65mm macro lens and MT-24EX twin flash unit a sign of poverty), and after a bit of tinkering and fiddling I’ve figured out a way to setup the specimen and flash units to create images of pinned specimens that I think are more than adequate for publication in taxonomic papers.

Here is one I did recently of the jewel beetle Actenodes calcaratus (family Buprestidae). This species is broadly distributed from the southwestern U.S. through Mexico and into Central America, where it breeds in dead branches of a variety of mostly fabaceous trees such as Acacia and Prosopis. During several trips to southern Mexico in recent years, Chuck Bellamy and I collected two new species of Actenodes that look very similar to A. calcaratus but differ in several important characters, primarily surface sculpture, the form and male coloration of the face, and male genitalia. A manuscript describing these two species and containing this and similar images of the new species was recently submitted for publication. Though not quite as razor-sharp as images created through focus-stacking processes, it still shows good detail and even lighting. What do you think?¹

¹ For those who find the pin head distracting, I am not a proponent of cloning out pin heads, debris, or other imperfections on images of preserved specimens in taxonomic papers. Other enhancements such as levels, sharpness, contrast, etc. are fine since these are all influenced greatly by lighting, but otherwise I believe the specimen needs to be presented exactly as it appears. A possible alternative is to remove the pin for imaging, but this presents a risk of damage to the specimen that is of questionable benefit in the case of non-type specimens—and downright irresponsible for primary types. Another alternative is to thoroughly clean and image the specimen prior to mounting, but this is rarely feasible as in most cases it is only after the specimen is mounted and studied further that its status as a new species is realized.

Actenodes calcaratus | MEXICO: Guerrero, Hwy 95, 5 km S Milpillas, 7.vii.1992, “big dead tree”, G. H. Nelson [FSCA]. Male plesiotype.

Copyright © Ted C. MacRae 2013

Backyard gems

I’ve been fortunate to have the chance to travel far and wide in my searches for insects—from the Gypsum Hills of the Great Plains and Sky Islands of the desert southwest to the subtropical riparian woodlands of the Lower Rio Grande Valley, tropical thorn forests of southern Mexico and veld of southern Africa. No matter how far I travel, however, I’m always happy to return to the Missouri Ozarks. It is here where I cut my entomological teeth so many years ago, and though I’ve now scrabbled around these ancient hills for more than three decades it continues to satisfy my thirst for natural history. Though not nearly as expansive as the Great Plains, there are nevertheless innumerable nooks and crannies nestled in the Ozarks, and I find myself constantly torn between looking for new spots (it would take several lifetimes to find them all) and going back to old favorites. Living in the northeastern “foothills” in the outskirts of St. Louis provides an ideal vantage for exploration; however, sometimes I am truly amazed at the natural history gems that can be found within a stone’s throw from my house. Some examples I’ve featured previously include Shaw Nature Reserve, home to a hotspot of the one-spotted tiger beetle, Castlewood State Park, where I found a gorgeously reddish population of the eastern big sand tiger beetle, and Victoria Glades Natural Area, site of the very first new species (and perhaps also the most beautiful) that I ever collected.

Englemann Woods Natural Area | Franklin Co., Missouri

Today I found another such area—Englemann Woods Natural Area, and at only 5 miles from my doorstep it is the closest natural gem that I have yet encountered. One of the last old-growth forests in the state, its deep loess deposits on dolomite bedrock overlooking the Missouri River valley support rich, mesic forests on the moister north and east facing slopes and dry-mesic forests on the drier west-facing slopes dissected by rich, wet-mesic forests with their hundreds-of-years-old trees. A remarkable forest of white oak, ash, basswood and maple in an area dominated by monotonous second-growth oak/hickory forests.

Steep north-facing slopes border the Missouri River valley.

It is not, however, the 200-year-old trees that will bring me back to this spot, but rather the understory on the north and east-facing slopes. Here occur some of the richest stands of eastern hornbean (Ostrya virginiana) that I have ever seen. This diminutive forest understory inhabitant is not particularly rare in Missouri, but as it prefers rather moist upland situations it is not commonly encountered in the dry-mesic forests that dominate much of the Ozarks. Stands of this tree, a member of the birch family (Betulaceae) are easy to spot in winter due to their habit of holding onto their dried canopy of tawny-brown leaves (see photo below).

Rich stands of eastern hornbeam (Ostrya virginiana) dominate the north- and east-slope understory.

Why am I so interested in this plant? It is the primary host of the jewel beetle species Agrilus champlaini. Unlike most other members of the genus, this species breeds in living trees rather than dead wood, their larvae creating characteristic swellings (galls, if you will) on the twigs and stems as they spiral around under the bark feeding on the cambium tissues before entering the wood to pupate and emerge as adults in spring. This species is known in Missouri from just two specimens, both collected by me way back in the 1980s as they emerged from galls that I had collected during the winter at two locations much further away from St. Louis. The presence of this rich stand of hornbeam just 5 miles from my home gives me the opportunity to not only search the area more thoroughly to look for the presence of galls from which I might rear additional specimens, but also to look for adults on their hosts during spring and (possibly, hopefully) succeed in photographing them alive.

Inside the “hornbeam forest.”

Another “draw” for me is the restoration work that has begun on some of the west-facing slopes in the areas. Pre-settlement Missouri was a far less wooded place than it is today, as evidenced by the richly descriptive writings penned by Henry Schoolcraft during his horseback journey through the Ozarks in the early 1800′s. At the interface between the great deciduous forests to the east and the expansive grasslands to the west, the forests of Missouri were historically a shifting mosaic of savanna and woodland mediated by fire. Relatively drier west-facing slopes were more prone to the occurrence of these fires, resulting in open woodlands with more diverse herbaceous and shrub layers. At the far extreme these habitats are most properly called “xeric dolomite/limestone prairie” but nearly universally referred to by Missourians as “glades”—islands of prairie in a sea of forest! I have sampled glades extensively in Missouri over the years, and they are perhaps my favorite of all Missouri habitats. However, it is not future glades or savannas that have me excited about Englemann Woods but rather the availability of freshly dead wood for jewel beetles and longhorned beetles resulting from the selective logging that has taken place as a first step towards restoration of such habitats on these west-slopes. The downed trees on these slopes and subsequent mortality of some still standing trees that is likely to result from the sudden exposure of their shade adapted trunks to full sun are likely to serve as a sink for these beetles for several years to come. I will want to use all the tools at my disposal for sampling them while I have this opportunity—beating, attraction to ultraviolet lights, and fermenting bait traps being the primary ones. It looks like I’d better stock up on molasses and cheap beer!

Restoration efforts on the west-facing slopes begins with selective logging.

Eastern red-cedar (Juniperus virginiana) is native to Missouri, but in our time it has become a major, invasive pest tree. The suppression of fire that came with settlement also freed this tree from a major constraining influence on its establishment in various habitats around the state, primarily dolomite/limestone glades. Nowadays most former glade habitats, unless actively managed to prevent it, have become choked with stands of this tree, resulting in shading out of the sun-loving plants that historically occurred much more commonly in the state. Untold dollars are spent each year by landscape managers on mechanical removal and controlled burns to remove red-cedar and prevent its reestablishment in these habitats. There is one habitat in Missouri, however, in which eastern red-cedar has reigned supreme for centuries or possibly millenia—dolomite/limestone bluff faces.

Craggly, old Eastern red-cedars (Juniperus virginiana) cling tenaciously to the towering dolomite bluffs.

With little more than a crack in the rock to serve as a toehold, red-cedars thrive where no other tree can, growing slowly, their gnarled trunks contorted and branches twisted by exposure to sun and wind and chronic lack of moisture. Some of the oldest trees in Missouri are red-cedars living on bluffs, with the oldest example reported coming from Missouri at an incredible 750–800 years old. There is something awe-inspiring about seeing a living organism that existed in my home state before there were roads and cars and guns. These ancient trees are now an easy drive from my house (though a rather strenuous 300-ft bushwhacking ascent to reach the bluff tops)—they seem ironically vulnerable now after having endured for so long against the forces of nature. For me, they will serve as a spiritual draw—a reason to return to this place again regardless of what success I might have at finding insects in the coming months.

This tree may pre-date Eurpoean settlement.

Adam-and-Eve orchid (Aplectrum hyemale).

Copyright © Ted C. MacRae 2013

Diffusion versus post-processing, or perhaps something even better?

One of the comments on my post Diffuser comparisons for 100mm macro lens was by Stephen Barlow, one of the original “concave diffuser” advocates, who claimed that the “dead” appearance of Photo #4 was an artifact of post-processing and not really a problem with the diffusion method itself. Heeding this comment, I reprocessed Photo #4 to see if this was really all that was needed to give it a “livelier” look by rather aggressively bumping up the brightness and contrast by 30% each (to correct for underexposure), then reducing the saturation by 10% (to correct for the effect on color caused by increased brightness and contrast), adjusted levels to a set point of 240 to add some more “high end,” and reduced highlights and shadows just a bit (10% each). Following is the original and then the reprocessed version of Photo #4:

Original post-processing

Additional post-processing.

There is no question that this additional reprocessing has greatly improved the photo. However, after I did this I got to thinking—why not try combining the two diffusers that gave the best results? Recall that the diffusion method in Photo #5 (SoftBoxes on flexible arm extenders) easily “won the vote” over Photo #4 (open concave diffuser) by a 2:1 margin (35 to 17). This may have been at least partly a result of the less than flattering post-processing of the original version of #4, but still the overall lighting effect on Photo #5 caused by the diffusion method used was quite dramatic. The only downside of the #5 method was the persistence of hot spots (albeit muted) from the flash heads and a dark background with lots of shadowing caused by light drop off (since the flash heads were mounted on the lens rather than extenders). Double diffusers are nothing new, the idea being that the first diffuser spreads the light out more before it hits the second diffuser than does a bare flash head, allowing even further diffusion of the light the reaches the subject (and background) for truly even lighting. I reasoned that using SoftBoxes on flexible arm extenders plus the concave diffuser would not only accomplish double diffusion but also allow controlled placement of the flash heads close to the specimen to maximize apparent light size and minimize light drop off. To test this I re-shot the same beetle with the same camera settings, and here is the result:

Flash heads mounted on flexible arms, diffused by SoftBoxes + open concave diffuser

My personal opinion is that this photo combines the best of both methods. While loss of light can be a problem with double diffusion, my use of extenders to place the flash heads close to the subject minimizes, or perhaps even completely negates this problem. Additionally, while subtle hot spots are still apparent, they are not nearly as apparent as in Photo #5 (SoftBox diffusers on extenders w/o concave diffuser—refresh your memory here) due to the additional diffusion, which also dramatically reduces shadowing as a result of better light throw. The hot spots are also more subtle than in #4 because of the larger apparent light size (a combination of closer flash head placement and the SoftBoxes), and is it just me or are the colors more vibrant and life-like in this photo compared to #4 (even reprocessed)? The flat colors were my biggest criticism of Photo #4, and even heavy-handed reprocessing, while helpful, didn’t completely bring it “back to life.” In contrast, the double-diffused photo required only typical post-processing to achieve a more than acceptable result—I have to believe that, all other things being equal, a photo that requires less post-processing is better than one that requires more.

Of course, using a setup like this in the studio is one thing—using it in the field is another. Both the extenders and the oversized concave diffuser are likely to make things a little clumsier in the field, and the two combined may be more clumsiness than I care to deal with. Nevertheless, the results from my test shots are certainly promising enough to give it an honest effort. Have I finally found a viable solution to diffusion in long-lens, full-flash macrophotography? We’ll find out this summer!

Copyright © Ted C. MacRae 2013

Diffuser comparisons for 100mm macro lens

I really wish I could just buy three Canon Speedlite 580EX II flash units, mount one directly on the camera, run the other two wirelessly on each side as slaves, put a nice big soft box diffuser on each of them, and be done with it! I’m beginning to think that’s the only way I’m going to get the kind of full flash insect macro photographs that I want with larger subjects that require the use of my 100mm macro lens. You know what I mean—nice, even, diffuse, vibrant light that comes at the subject from multiple directions (eliminating those annoying specular highlights in the eyes that result from more unidirectional lighting) and with enough power to allow minimal flash pulse durations (resulting in maximum motion freeze). But I can’t—the money is not in the budget, and even if it was I’d have to think seriously about the logistics of carrying and setting up in the field three Speedlites every time I wanted to photograph an (often moving) insect.

Thus, I continue trying to come up with some kind of system that makes the most of my Canon MT-24EX twin flash unit. It’s not that I don’t like this flash unit—I love it because of its light weight (good for field use) and the front-of-the-lens mounting feature that, with its dual heads, gets the flash heads closer to the subject but avoids the “flat” lighting effect of typical ring flash units. In addition, for those shooting insect macro photographs with Canon’s shorter focal length MP-E65 macro lens, the twin flash unit is probably the best choice of all, since the lens is right on top of the subject and it is relatively easy to place diffusing materials between the subject and the flash heads—Alex (Myrmecos) with his tracing paper diffuser and Kurt (Up Close with Nature) with his concave foam diffuser are two of the more successful designs out there. I use my MP-E65 lens a lot, but I use my 100mm macro lens a lot more because many of the beetles I photograph are best photographed at magnification ranges between 0.5–1.0X and, thus, are a little too large for the 65mm lens. The longer lens-to-subject distance of the 100mm lens may be helpful for working with skittish subjects, but it also creates challenges for the MT-24EX because of its relatively low power (more light drop off) and small flash heads (more specular highlighting). For the past couple of years I’ve been using a large sheet of polypropylene foam jury-rigged to the front of the lens, and while it too has functioned fairly well, I keep thinking that if I can just get the flash heads closer to the subject—each fitted with a good diffuser—then it should be possible to achieve results similar to what can be done with the 65 mm lens.

The photos below show the results of some of the ideas I’ve been working on. My main idea was to use extenders that would allow adjustable placement of the flash heads relatively close to the subject and diffuse the light from them with a modified version of the Sto-Fens+Puffers that I have tried in the past. Here is an example of the system mounted on my camera using cheap, flexible arms mounted on a plate attached to the bottom of the camera. If I decide to use this system in the field I would want to purchase much sturdier extenders (e.g. Really Right Stuff), but at only $25 these flexible arms are perfect for proof-of-concept testing. For the modified Sto-Fens+Puffers, I completed the modifications shown by Dalantech (No Cropping Zone) (I was planning to do this when I first tried the Sto-Fens+Puffers but soon found that I preferred the concave and tent designs by Kurt and Alex, at least for use with the 65mm lens). At any rate, to test the ideas I selected a very large (for long subject-to-lens distance), very shiny (for maximum specular highlighting potential) beetle from my collection (Megaloxantha purpurascens peninsulae, a stunning jewel beetle from Palawan, Philippines) and set it up for “face shots” that simulate my favorite pose for beetles in the field. Keep in mind that this was not intended to be a test of lighting for pinned specimens in the studio—that is not my interest, and there are much better approaches for doing that—but rather a proxy for the kind of lighting and diffusion I might achieve in the field. Here are the results:

#1 – flash heads mounted on lens, diffused by modified Sto-Fens+Puffers

The example above show the results obtained when using the modified Sto-Fens+Puffers with the flash heads mounted directly to the front of the lens. I didn’t try this shot without diffusers, but I doubt it would be much worse than this—specular highlighting is bad because of the small apparent light size, and overall the lighting is not very even with dark shadows and harsh highlights. This shot is a perfect example of the problems inherent in using the twin-flash with a long macro lens.

#2 – flash heads mounted on flexible arms, diffused by modified Sto-Fens+Puffers

This second shot shows the results when the modified Sto-Fens+Puffers are mounted on the flexible arm extenders and positioned as close to the subject as possible to maximize apparent light size. This was supposed to be the system that gave me the results I was looking for, but honestly I am not impressed. The highlights in the eyes are certainly larger than in the previous photo, and the overall lighting is not quite as uneven, but still the highlights are harsh and fairly sharply defined. Considering the greater difficulty in positioning the flash heads compared to lens-mounted, I have to consider the marginal improvement in lighting not worth the effort.

#3 – flash heads mounted on lens, diffused with modified Sto-Fens+ Puffers and concave diffuser (closed)

This third shot has the modified Sto-Fens+Puffers once again mounted on the lens, but also attached is my trusty concave diffuser. Honestly this combination of diffusers provides much better overall lighting and softening of the highlights compared to the previous shot, even though the flash heads are mounted on the lens rather than positioned close to the subject. Apparently the concave diffuser, though further away from the subject, still has larger apparent size and thus allows light to be transmitted to the subject from a larger apparent area. I have not normally used another diffuser between the flash heads and the concave diffuser, but my impression from this shot is that the modified Sto-Fens+Puffers do a good job of dispersing light before it hits the concave diffuser to soften the “hot spots” behind it and provide somewhat more even lighting across its surface.

#4 – flash heads mounted on lens, diffused with modified Sto-Fens+ Puffers and concave diffuser (open)

When I use the concave diffuser, I normally pull the corners back and attach them to the tops of the flash heads with Velcro to minimize light blow back (although how effective it is I really don’t know). Just for kicks, I decided to try some shots with the concave diffuser not pulled back, but left open and extending out over the subject. I did this because that actually more closely approximates how smaller versions of concave diffusers are used with the 65mm lens. The effect was not only remarkable diffusion of light, with specular highlights and hot spots almost completely lacking, but also much better lighting behind rather than just on the front of the specimen. That said, the quality of the light lacks vibrancy and seems somewhat “dead,” perhaps because of the great distance between the flash heads and the diffuser and the MT-24EX units relatively limited power. The large diffuser extending far out in front of the lens might cause problems with bumping and skittish subjects, but I am intrigued enough by this result to continue with some field testing to see what I think.

#5 – flash heads mounted on flexible arms, diffused by SoftBoxes

The final shot shows the results of another promising setup—this one again uses the flash heads mounted on flexible arm extenders to get them close to the subject, but instead of the modified Sto-Fens+Puffers I fitted each flash head with a mini SoftBox. This was not easy, as the SoftBox is designed for much larger flash heads than those of the MT-24EX, so I took another set of Sto-Fen diffusers, cut off the face, then hot-glued the SoftBox to the open Sto-Fen. Thus modified it was a simple matter to “snap” the SoftBoxes in place over the flash heads. Despite the term ‘mini’ these Soft Boxes still provide a much larger area for light transmission than the modified Sto-Fens+Puffers, and this much larger apparent light size has a dramatic effect on the overall lighting and diffusion. I’m tempted to say I like this one best. However, I do have to consider ease of function in the field—the lens-mounted Sto-Fen+Puffers and concave diffuser, either open or closed, would certainly be easier and involve no further cost (for better extenders than the cheap flexible arms I now have), but if SoftBoxes on flash heads placed close to the subject gives better results than I may have to go with it.

Will you please help me decide? I setup this little poll so you can tell me which of the systems you thought gave the most pleasing result in terms of vibrant, evenly diffused light. I can’t (to my knowledge) tell who’s voting (and if there is a way don’t tell me because I don’t want to know), so don’t let privacy concerns prevent you from adding your vote—the more voters that participate, the better information I get to help me with my decision.

Copyright © Ted C. MacRae 2013

It’s always a happy day…

…when the latest issue of The Coleopterists Bulletin arrives in my mailbox. On this occasion it was the December issue of Volume 66—nine papers and eight scientific notes filling 84 pages of beetle awesomeness. It’s pure elytral ecstasy! I presume I am like most subscribers—rapidly scanning the Table of Contents on the back cover to see if any deal directly with my preferred taxa. Yes! Two papers dealing with Buprestidae (jewel beetles), one on Cerambycidae (longhorned beetles), and one on Cicindelinae (tiger beetles)—a real bonanza. After that, a more cursory look through the rest of the Table of Contents to see what other papers look interesting enough to at least scan through.

For me the most interesting are the two Buprestidae papers, with Hansen et al. documenting new state records, larval hosts, and biological notes for 47 North American species and Westcott & Murray reporting the introduction into the U.S. of yet another Eurasian exotic (Trachys minutus) and its apparent establishment in Massachusetts. As the current “keeper” of distributional records and host plant associations for North American jewel beetles (along with Rick Westcott, Salem, Oregon), I will be busily updating my database over the next few days to reflect these new records. I am a great fan of “notes” papers such as these (and am, in fact, currently finishing a similar manuscript with co-author Joshua Basham, who is also a co-author on the Hansen et al. paper). However, I do have a few quibbles—Hansen et al. report Agrilus  quadriguttatatus as a new record for Tennessee, but it is already known from that state, and Cercis canadensis (eastern redbud) is reported as a new larval host for Anthaxia (Haplanthaxia) cyanella despite the prior records from that host by Knull (1920) and Hespenheide (1974). More puzzlingly, the authors record Agrilus lecontei celticola from locations in eastern Tennessee despite guidance from me on several occasions that this subspecies, while perhaps distinctive in Texas, transitions broadly across Louisiana and Mississippi  with the nominate subspecies. As such, material from eastern Tennessee cannot be regarded conclusively to represent this subspecies (and I remain unconvinced even that the subspecific distinction is warranted). Lastly, in recording Actenodes simi from Tennessee, the authors mention that the closest previous record is from Missouri with no specific locality mentioned (Fisher 1942), even though I recently recorded several specific locations for the species in eastern and southern Missouri (MacRae & Nelson 2003). The overall impression is that the authors are not fully versed in recent literature on Buprestidae and have instead relied exclusively on the recent Nelson et al. (2008) catalogue—known amongst buprestid workers to be incomplete and with errors—as the only source for determining the status of their records.

Among Cerambycidae, Raje et al. report the results of molecular analyses on two color forms of Sternidius alpha. This broadly distributed and highly variable species exhibits multiple color variants across its range, leading to the description of multiple subspecies that were eventually synonymized under the current name. Their analysis of the barcoding region of the cytochrome oxidase I gene, however, revealed three distinct clades among the two color forms, suggesting the potential for taxonomic significance. More work, of course, is needed from additional color morphs from different localities.

Finally, my friend Matt Brust and colleagues discuss the ovipositional behavior of numerous species of North American tiger beetles, unexpectedly finding that many oviposit only after digging some distance below the surface of the soil. This information is extremely valuable for those interested in rearing tiger beetles for description of larval stages, expanding the window of survey for species with limited temporal occurrence, and cross-breeding studies. To that end, and of greatest interest to me, they have included numerous observations from their own studies that have resulted in the development of successful protocols and rapid rearing of large numbers of larvae to adulthood.

Actually, there is one more thing… For several years now the December issue, as a bonus, has been accompanied by the Patricia Vaurie Series Monograph as a supplement to that year’s volume. This year’s issue features a revision of the scarab genus Euphoria by Jesús Orozco, and although I have not studied it carefully it looks like a robust treatment of the group. Yes, I know that scarabs are not one of my primary interest groups, but show me a coleopterist that—regardless of the group they work on— does not stop and collect these gorgeous, colorful, flower-loving beetles whenever they encounter them and I’ll show you a coleopterist that is far too restrictive in their natural history interests! Based on examination of nearly 19,000 specimens from 67 collections, the work considers 59 valid species (ten of which are described as new) distributed throughout the Western Hemisphere. Complete with keys to species and, for each, synonymy, description, diagnosis, taxonomic history, natural history, temporal occurrence geographic distribution, and—of critical importance in my opinion—full data for all specimens examined, it is everything a good revision should be. Then there are the color plates—one full page for each species—with a large dorsal habitus view, closeups of the head, male genitalia, and color variants, a temporal distribution chart, and a map of its geographical distribution. Again, while I may not be a serious student of scarabs, you can bet that I’ll be going back through my holdings of Euphoria beetles and checking them to make sure they conform to this new standard of knowledge on the group.

REFERENCES:

Brust, M. L., C. B. Knisley, S. M. Spomer & K. Miwa. 2012. Observations of oviposition behavior among North American tiger beetle (Coleoptera: Carabidae: Cicindelinae) species and notes on mass rearing. The Coleopterists Bulletin 66(4):309–314.

Fisher, W. S. 1942. A revision of North American species of buprestid beetles belonging to the tribe Chrysobothrini. U. S. Department of Agriculture, Miscellaneous Publication 470, 1–275.

Hansen, J. A., J. P. Basham, J. B. Oliver, N. N. Youseef, W. E. Klingeman, J. K. Moulton & D. C. Fare. 2012. New state and host plant records for metallic woodboring beetles (Coleoptera: Buprestidae) in Tennessee, U.S.A. The Coleopterists Bulletin 66(4):337–343.

Hespenheide, H. A. 1974.  Notes on the ecology, distribution, and taxonomy of certain Buprestidae.  The Coleopterists Bulletin 27(4) [1973]:183–186.

Knull, J. N. 1920. Notes on Buprestidae with description of a new species (Coleop.). Entomological News 31(1):4–12.

MacRae, T. C. and G. H. Nelson. 2003. Distributional and biological notes on Buprestidae (Coleoptera) in North and Central America and the West Indies, with validation of one species. The Coleopterists Bulletin 57(1):57–70.

Nelson, G. H., G. C. Walters, Jr., R. D. Haines, & C. L. Bellamy.  2008.  A Catalogue and Bibliography of the Buprestoidea of America North of Mexico.  Coleopterists Society Special Publication No. 4, The Coleopterists Society, North Potomac, Maryland, 274 pp.

Orozco, J. 2012. Monographic revision of the American genus Euphoria Burmeister, 1842 (Coleoptera: Scarabaeidae: Cetoniinae). Coleopterists Society Monographs, Patricia Vaurie Series No. 11, 182 pp.

Raje, K. R., V. R. Ferris & J. D. Holland. 2012. Two color variants of Sternidius alpha (Say) (Coleoptera: Cerambycidae) show dissimilar cytochrome oxidase I genes. The Coleopterists Bulletin 66(4):333–336.

Westcott, R. L. & T. C. Murray. 2012. An exotic leafminer, Trachys minutus (L.) (Coleoptera: Buprestidae), found in Massachusetts, U.S.A. The Coleopterists Bulletin 66(4):360–361.

Copyright © Ted C. MacRae 2013

Let’s make a deal!

Image source unknown

I recently conducted a complete reorganization of the jewel beetles (superfamily Buprestoidea) in my collection (TCMC). The primary purpose of this was to bring the nomenclature and arrangement of the collection into agreement with the recently published World Catalogue of Buprestoidea (Bellamy 2008) and accurately document the taxa represented in the collection and their numbers. In the short term this will be helpful not only in visualizing what is represented but also what is missing (particularly in North America), while longer term it lays the groundwork for the eventual donation of my collection to a public institution.

In an active, working collection, no inventory is ever fully up-to-date. In my case, the inventory includes only completely curated material that has been incorporated into the main cabinets. I still have several years worth of material in various states of curation—i.e., unmounted, mounted but unlabeled, or labeled but unidentified. That said, the main collection now contains more than 23,000 specimens of Buprestoidea representing 1,500+ species worldwide. Of the species represented, 37% are Nearctic (U.S./Canada), 22% Palearctic (Europe, North Africa, temperate Asia), 19% Neotropical (Latin America), 10% Afrotropical (Subsaharan Africa), 7% Indomalayan (tropical Asia) and 6% Australian (Australia/New Zealand). The collection also contains 492 paratype specimens representing 77 species. The inventory has been converted to  PDF and uploaded for access by the link below. It lists all of the species represented, with nomenclature updated and taxa arranged according to Bellamy (2008) and number of specimens  indicated for each. Also indicated are higher taxa not yet represented in the collection (shown in gray rather than black text) so that the collection holdings can be placed in context of a complete higher classification for the superfamily.

 Click to see full inventory of TCMC Buprestoidea

Of course, as a North American, the Nearctic fauna is the primary focus of my taxonomic and biological studies. As a result, I am keen to have the Nearctic fauna represented as completely as possible in my collection. Currently I have 75% (595) of the 790 species and non-nominate subspecies currently recognized in North America. Obviously, by now I’ve picked most of the low-hanging fruit, and the last 25% will be much more difficult to get. Many of these are truly rare species that I may never find (some are known only by the holotype), while others are more common but occur in areas that I have limited opportunity to visit. These species are also indicated in the above inventory (again, in gray text) but are also listed below for easy reference. If you have any of the species on this list, please let me know and also what you might like to receive in exchange for them. I have not only many species of Buprestidae from around the world to offer, but also beetles in other families such as longhorned beetles (Cerambycidae), tiger beetles (Cicindelinae), scarabs (Scarabaeoidea), and even non-beetles such as treehoppers (Membracidae) and cicadas (Cicadoidea). Let’s make a deal!

REFERENCE:

Bellamy, C. L. 2008. World Catalogue and Bibliography of the Jewel Beetles (Coleoptera: Buprestoidea),  Volumes 1–5. Pensoft Series Faunistica, 3125 pp.

Copyright © Ted C. MacRae 2013

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T.C.MacRae Collection Desiderata

Family SCHIZOPODIDAE LeConte 1859
Subfamily SCHIZOPODINAE LeConte 1859
Tribe SCHIZOPODINI LeConte 1859

Genus Schizopus LeConte 1858
— sallei ssp. sallei Horn 1885
— sallei ssp. nigricans Nelson 1991
Genus Dystaxia LeConte 1866
— elegans Fall 1905

Family BUPRESTIDAE Leach 1815
Subfamily POLYCESTINAE Lacordaire 1857
Acmaeoderioid lineage sensu Volkovitsh 2001
Tribe HAPLOSTETHINI LeConte 1861

Genus Mastogenius Solier 1849
— arizonicus Bellamy 2002
— puncticollis Schaeffer 1919

Tribe ACMAEODERINI Kerremans 1893
Subtribe ACMAEODEROIDINA Cobos 1955

Genus Acmaeoderoides Van Dyke 1942
— cazieri Nelson 1968
— depressus Nelson 1968

Subtribe ACMAEODERINA Kerremans 1893

Genus Acmaeodera Eschscholtz 1829
– Subgenus Acmaeodera (s. str.)
— audreyae Westcott & Barr 2007
— bryanti Van Dyke 1953
— comata LeConte 1858
— consors Horn 1878
— cubaecola Jaquelin du Val 1857
— discalis Cazier 1940
— dolorosa ssp. liberta Fall 1922
— fattigi Knull 1953
— flavosticta Horn 1878
— horni Fall 1899
— inyoensis Cazier 1940
— laticollis Kerremans 1902
— morbosa Fall 1899
— pubiventris ssp. panocheae Westcott 2001
— recticolloides Westcott 1971
— starrae Knull 1966
— subbalteata LeConte 1863
— thoracata Knull 1974
— tildenorum Nelson & Westcott 1995
— wheeleri Van Dyke 1919

Genus Acmaeoderopsis Barr 1974
— prosopis Davidson 2006
— rockefelleri (Cazier 1951)
— varipilis (Van Dyke 1934)

Genus Anambodera Barr 1974
— nebulosa (Horn 1894)
— santarosae (Knull 1960)

Polyctesioid lineage sensu Volkovitsh 2001
Chrysophana generic group [tribal level] sensu Volkovitsh 2001

Genus Beerellus Nelson 1982
— taxodii Nelson 1982

Polycestioid lineage sensu Volkovitsh 2001
Tribe POLYCESTINI Lacordaire 1857

Genus Polycesta Dejean 1833
– Subgenus Polycesta (s. str.)
— angulosa Jacquelin du Val 1857
– Subgenus Polycesta (Arizonica) Cobos 1981
— arizonica ssp. acidota Cazier 1951
– Subgenus Polycesta (Tularensia) Nelson 1997
— crypta Barr 1949

Tribe TYNDARINI Cobos 1955
Subtribe TYNDARINA Cobos 1955

Genus Paratyndaris Fisher 1919
– Subgenus Paratyndaris (s. str.)
— anomalis Knull 1937
— crandalli Knull 1941
— grassmani Parker 1947
— quadrinotata Knull 1938

Subfamily CHRYSOCHROINAE Laporte 1835
Chrysochroid lineage sensu Bellamy 2003
Nanularia generic group [tribal level] sensu Volkovitsh 2001

Genus Nanularia Casey 1909
— cupreofusca Casey 1909
— pygmaea (Knull 1941)

Tribe CHRYSOCHROINI Laporte 1835
Subtribe CHALCOPHORINA Lacordaire 1857
Texania generic group sensu Volkovitsh 2001

Genus Texania Casey 1909
— langeri (Chevrolat 1853)

Tribe POECILONOTINI Jakobson 1913
Subtribe POECILONOTINA Jakobson 1913

Genus Poecilonota Eschscholtz 1829
— ferrea (Melsheimer 1845)
— montana Chamberlin 1922
— viridicyanea Nelson1997

Dicercioid lineage sensu Bellamy 2003
Tribe DICERCINI Gistel 1848
Subtribe HIPPOMELANINA Holynski 1993

Genus Hippomelas Laporte & Gory 1837
— martini Nelson 1996
— parkeri Nelson 1996

Genus Gyascutus LeConte 1858
– Subgenus Gyascutus (s. str.)
— jeanae (Nelson 1988)
— pacificus (Chamberlin 1938)

Genus Barrellus Nelson & Bellamy 1996
— femoratus (Knull 1941)

Subtribe DICERCINA Gistel 1848
Dicerca generic group sensu Volkovitsh 2001

Genus Dicerca Eschscholtz 1829
— dumolini (Laporte & Gory 1837)
— hornii nelsoni Beer 1974
— lugubris LeConte 1860
— mutica LeConte 1860
— sexualis Crotch 1873
— spreta (Gory 1841)
— tuberculata (Laporte & Gory 1837)

Subfamily BUPRESTINAE Leach 1815
Buprestioid lineage sensu Volkovitsh 2001
Buprestinioid branch sensu Volkovitsh 2001
Tribe BUPRESTINI Leach 1815
Subtribe TRACHYKELINA Holynski 1988

Genus Trachykele Marseul 1865
— fattigi Knull 1954
— opulenta Fall 1906

Subtribe BUPRESTINA Leach 1815

Genus Buprestis Linnaeus 1758
– Subgenus Buprestis (Cypriacis) Casey 1909
— intricata Casey 1909
— prospera Casey 1909
– Subgenus Buprestis (Knulliobuprestis) Kurosawa 1988
— fremontiae Burke 1924
– Subgenus Buprestis (Stereosa) Casey 1909
— apricans Herbst 1801
— decora Fabricius 1775

Anthaxioid lineage sensu Volkovitsh 2001
Anthaxiinioid branch sensu Volkovitsh 2001
Tribe ANTHAXIINI Gory et Laporte 1839

Genus Anthaxia Eschscholtz 1829
– Subgenus Anthaxia (Haplanthaxia) Reitter 1911
— carya Wellso & Jackman 2006
— caseyi ssp. sublaevis Van Dyke 1916
– Subgenus Anthaxia (Melanthaxia) Rikhter 1944
— barri Bílý 1995
— californica Obenberger 1914
— cupriola Barr 1971
— emarginata Barr 1971
— embrikstrandella Obenberger 1936
— exasperans Cobos 1958
— furnissi Barr 1971
— helferiana Bílý 1995
— hurdi Cobos 1958
— nanula Casey 1884
— neofunerula Obenberger 1942
— nevadensis Obenberger 1928
— oregonensis Obenberger 1942
— porella Barr 1971
— sculpturata Barr 1971
— serripennis Obenberger 1936
— strigata LeConte 1859
— subprasina Cobos 1959
— tarsalis Barr 1971
— wallowae Obenberger 1942

Tribe XENORHIPIDINI Cobos 1986
Subtribe XENORHIPIDINA Cobos 1986

Genus Hesperorhipis Fall 1930
— hyperbola ssp. californica Knull 1947
— jacumbae Knull 1954
— mirabilis ssp. mirabilis Knull 1947

Chrysobothrioid lineage sensu Volkovitsh 2001
Melanophilinioid branch sensu Volkovitsh 2001
Tribe MELANOPHILINI Bedel 1921
Subtribe MELANOPHILINA Bedel 1921

Genus Phaenops Dejean 1833
— carolina (Manee 1913)
— caseyi (Obenberger 1944)
— obenbergeri (Knull 1952)
— vandykei Obenberger 1944

Chrysobothrinioid branch sensu Volkovitsh 2001
Tribe ACTENODINI Gistel 1848

Genus Actenodes Dejean 1833
— arizonicus Knull 1927
— mimicus Knull 1964

Tribe CHRYSOBOTHRINI Gory et Laporte 1838

Genus Chrysobothris Eschscholtz 1829
– Subgenus Chrysobothris (s. str.)
— aeneola LeConte 1860
— bacchari Van Dyke 1923
— bicolor Horn 1894
— bisinuata Chamberlin 1938
— bispinosa Schaeffer 1909
— boharti Van Dyke 1934
— breviloboides Barr 1969
— caurina Horn 1886
— chamberliniana Fisher 1948
— costifrons ssp. costifrons Waterhouse 1887
— culbersoniana Knull 1943
— cupressicona Barr & Westcott 1976
— deserta Horn 1886
— dolata Horn 1886
— fragariae Fisher 1930
— grindeliae Van Dyke 1937
— helferi Fisher 1942
— hidalgoensis Knull 1951
— horningi Barr 1969
— hubbardi Fisher 1942
— idahoensis Barr 1969
— kelloggi Knull 1937
— knulli Nelson 1975
— nelsoni Westcott & Alten 2006
— oregona Chamberlin 1934
— orono Frost 1920
— paragrindeliae Knull 1943
— potentillae Barr 1969
— pseudacutipennis Obenberger 1940
— pubilineata Vogt 1949
— purpurata Bland 1864
— roguensis Beer 1967
— schaefferi Obenberger 1934
— schistomorion Westcott & Davidson 2001
— scitula Gory 1841
—sexfasciata ssp. sexfasciata Schaeffer 1919
— sloicola Manley & Wellso 1976
— smaragdula Fall 1976
— socialis ssp. apache Westcott & Barr 2007
— speculifer Horn 1886
— subopaca Schaeffer 1904
— vivida Knull 1952
— westcotti Barr 1969
— wickhami Fisher 1942

Genus Knowltonia Fisher 1935
— alleni (Cazier 1938)
— atrifasciata (LeConte 1878)

Subfamily AGRILINAE Laporte 1835
Tribe AGRILINI Laporte 1835
Subtribe AGRILINA Laporte 1835

Genus Agrilus Curtis 1825
– Subgenus Agrilus (s. str.)
— hazardi Knull 1966
– Subgenus Agrilus (Engyaulus) Waterhouse 1889
— inhabilis ssp. cuprinus Nelson 1996
— utahensis Westcott 1996
– Subgenus Agrilus (Quercagrilus) Alexeev 1998
— derasofasciatus Boisduval & Lacordaire 1835
– Subgenus Agrilus (Uragrilus) Semenov-Tian-Shanskij 1935
— granulatus ssp. mojavei Knull 1952
— sayi Saunders 1871
– Subgenus undefined
— amelanchieri Knull 1944
— arizonus Knull 1934
— audax Horn 1891
— aurilaterus Waterhouse 1889
— bespencus Barr 2008
— burkei Fisher 1917
— catalinae Knull 1940
— cercidii Knull 1937
— cochisei Knull 1948
— criddlei Frost 1920
— davisi Knull 1941
— delicatulus Waterhouse 1889
— dozieri Fisher 1918
— exiguellus Fisher 1928
— floridanus Crotch 1873
— funestus Gory 1841
— geronimoi Knull 1950
— gillespiensis Knull 1947
— hazardi Knull 1966
— horni Kerremans 1900
— jacobinus Horn 1891
— langei Obenberger 1935
— latifrons Waterhouse 1889
— montosae Barr 2008
— neabditus Knull 1935
— nevadensis Horn 1891
— nigricans Gory 1841
— obscurilineatus Vogt 1949
— olivaceoniger Fisher 1928
— ometauhtli Fisher 1938
— palmerleei Knull 1944
— parabductus Knull 1954
— pilosicollis Fisher 1928
— pseudocoryli Fisher 1928
— pubifrons Fisher 1928
— restrictus Waterhouse 1889
— shoemakeri Knull 1938
— sierrae Van Dyke 1923
— snowi Fall 1905
— torquatus LeConte 1860
— waltersi Nelson 1985
— wenzeli Knull 1934

Tribe TRACHYINI Laporte 1835
Subtribe BRACHYINA Cobos 1979

Genus Taphrocerus Solier 1833
— floridanus Obenberger 1934

Subtribe PACHYSCHELINA Böving et Craighead 1931

Genus Pachyschelus Solier 1833
— fisheri Vogt 1949
— schwartzi Kerremans 1892
— vogti Hespenheide 2003