Fire ant winged reproductives: male and female

While in Austin at the Entomological Society of America meetings, I had the chance to tour The University of Texas at Austin’s Brackenridge Field Laboratory.  Located on 82 acres of land bordering the Colorado River, the station supports studies in biodiversity, ecosystem change and natural history. A major focus of research at the station involves efforts to establish biological control agents for control of imported fire ant (Solenopsis invicta) using entomopathogens and parasitoids (e.g. phorid flies in the genus Pseudacteon). This research relies on maintaining cultures of fire ants to support rearing of the phorid fly. While time was limited and I did not have much opportunity to photograph either the ant or the fly, I did manage to quickly sneak in a shot or two of some winged reproductives that were removed from the teaming formicid mass in a rearing tray and placed on a table top for all to see (and when I say “a shot or two” I mean it. I had the chance only for one shot of the female and two of the male as they crawled crazily about and the tour leader quickly tried to move us on). I’m sure Alex Wild has all stages/forms of this species covered in spades, but the sexually dimorphic winged reproductives were new for me, and perhaps some readers of this blog as well.

Solenopsis invicta winged reproductives: male (top), female (bottom).

Solenopsis invicta winged reproductives: male (top), female (bottom).

Copyright © Ted C. MacRae 2013

ESA Insect Macrophotography Workshop

Today is the last day of the Entomological Society of America (ESA) Annual Meeting in Austin, Texas, and it has been an action packed week for me. Annual meetings such as this serve several purposes. In addition to seeing talks on a variety of subjects—in my case covering subjects ranging from insect resistance management to scientific outreach to beetle systematics—they also offer the chance to establish new connections with other entomologists that share common interests and reinforce existing ones. Of course, a major part of my interest in entomology revolves around insect macrophotography, and in recent years ESA has begun to cater to the entomological photographer contingent within the society. Last year’s meetings featured a macrophotography symposium titled, “Entomologists Beyond Borders” (for which I was one of the invited speakers), and this year featured an Insect Macrophotography Workshop led by Austin-based entomologist/photographer Ian Wright. Having done this for a few years now I figured a lot of the workshop might be review for me, but I still have much to learn and am willing to accept new ideas from any source. Besides, the workshop involved a field trip to a local habitat to try out our insect photography skills, and for a field junkie like me time in the field at an otherwise all-indoor event spanning close to a week is always welcome. The location of the meetings in Austin this year made this possible, as even in mid-November there still remain insects out and about that can be photographed if the weather cooperates (and it did).

This will be a somewhat different post than what I usually post here. Rather than featuring photos of a certain species and using them as a backdrop for a more detailed look at their taxonomy or natural history, I’m just going to post all the photos that I ended up keeping from the field trip portion of the workshop with just a comment or two about each. We went to the city’s nearby waste-water treatment facility, the grounds of which are wild and woolly enough to provide habitat for insects, and spent about an hour and a half seeing what we could find. For myself, it was a chance to photograph some insects I’ve not normally tried to photograph (i.e., dragonflies, ambush bugs) and get more practice on my blue sky technique. I did appreciate the chance to spend some time talking to Ian during while we traveled to the site and back, and I also ended up helping other participants with their camera equipment questions and technique suggestions. With that, here are the photos I took—I’ll be curious to see what readers think of this post format versus my more typical style.

Micrutalis calva

Micrutalis calva (Hemiptera: Membracidae) on silverleaf nightshade (Solanum elaeagnifolium).

Micrutalis calva

This species of treehopper is restricted to herbaceous plant hosts.

Anax junius

Anax junius (Odonata: Aeshnidae), one of the darner species of dragonfly.

Anax junius

This adult was perched on a dead twig tip and seemed to be “asleep.”

Anax junius

I clipped the perch and held it up for these “in-your-face” shots – it then awoke with a start and flew off.

Phymata sp.

Phymata sp. (Hemiptera: Reduviidae), one of the so-called “jagged ambush bugs.”

Phymata sp.

Formerly a separate family, ambush bugs are now combined with assassin bugs (family Reduviidae).

Acmaeodera flavomarginata

Acmaeodera flavomarginata (Coleoptera: Buprestidae).

Acmaeodera flavomarginata

This is one of a few species of jewel beetle in the southcentral US that are active during the fall.

Mecaphesa sp.

Mecaphesa sp. (Araneae: Thomisidae), one of the crab spiders

Mecaphesa sp.

Cryptic coloration allows the spider to lurk unseen by potential insect prey visiting the flower.

Gratiana pallidula

Gratiana pallidula (Coleoptera: Chrysomelidae) on silverleaf nightshade (Solanum eleagnifolium).

Gratiana pallidula

A type of tortoise beetle, adults “clamp” down against the leaf as a defense against predators.

Copyright © Ted C. MacRae 2013

The Darwin Beetle

Like most modern biologists, Charles Darwin ranks high on my short list of intellectual/entomological heroes. Actually, with all due respect to others on the list—Carl Linnaeus, Alfred Russell Wallace, John Lawrence LeConte, and others, Darwin sits at #1. His theory of evolution, offered more than 150 years ago to a powerfully skeptical world, continues to provide the basic framework for modern biology (as Theodosius Dobzhansky said in his 1973 paper in American Biology Teacher, “Nothing in biology makes sense except in the light of evolution”). Thus, when Max Barclay recently posted on Facebook a photograph of a beetle collected by Charles Darwin himself, it reminded me that I have yet to visit Down House in Kent (the home of Charles Darwin) or to see anything personally touched by the man whose legacy I revere more than any other. Little did I know that Max did not post the photo from The Natural History Museum in London, but from Austin, Texas where he and I were each arriving for the annual meetings of the Entomological Society of America. When I commented on his post how I would love to see a beetle collected by Darwin someday, Max replied that he had the specimen with him and that he would bring it to the meetings for me to see (and I quote, “Most fun it has had since it flew to 22-year-old Charles Darwin’s gas lamp in Tierra Del Fuego in December 1832”). Can you imagine my anticipation?! True to his word, Max found me at the opening reception, came up from behind me, and placed  the plastic, see-through box housing the specimen on the table in front of me. I recognized it instantly, but still seeing “C. Darwin” on the label almost felt like I’d just met the man himself. I asked Max if it was okay to open the box, to which he agreed, and I even dared to grab the pin head and re-position the specimen for photographs. Call me crazy, but it was as spiritual an experience as I’ve had since, well… “Mrs. Ples” stood before me!

At any rate, here is the “Darwin Beetle,” followed by proof that I really got to hold it!

Sericoides glacialis (Fabricius), collected at Tierro del Fuego in 1832 by Charles Darwin.

Scarab beetle collected at Tierro del Fuego in 1832 by Charles Darwin. Identified as Sericoides glacialis (Fabricius) by Andrew B. T. Smith in 2012 after standing for many years as ‘Sericodes Reichii Guer.’

Holding the ''Darwin Beetle''

Holding the ”Darwin Beetle”

Copyright © Ted C. MacRae 2013

Midget male meloid mates with mega mama

Pyrota bilineata on flowers of Chrysothamnus viscidflorus | San Juan Co., Utah

Pyrota bilineata on flowers of Chrysothamnus viscidflorus | San Juan Co., Utah

While looking for longhorned beetles in the genus Crossidius on flowers of yellow rabbitbrush (Chrysothamnus viscidiflorus) in southern Utah, I encountered one particular plant with numerous blister beetles (family Meloidae) on its blossoms. The orange color, two black pronotal spots, and distinctive black and white longitudinal elytral stripes leave no doubt as to its identity—Pyrota bilineata, but for good measure I sent a photo to my field mate for the trip, Jeff Huether, who confirmed its identity. I had seen singletons of this species at a few previous localities during the trip, so I was intrigued by the large numbers of individuals congregated on this single plant. As I looked at them, I saw one individual that appeared to have something stuck to the tip of its abdomen. I peered closer to get a better look and, to my surprise, discovered that it was actually a male in the act of mating. The male was tiny, only one-third the size of the female, representing about as extreme a size difference in mating insects as I’ve ever seen.

Pyrota bilineata on flowers of Chrysothamnus viscidflorus | San Juan Co., Utah

A tiny male mates with the ginormous female.

Many species of blister beetles exhibit tremendous size variability, and a unique aspect of some species’ mating behavior is the cantharidin-packed spermatophore produced by males and transferred to females during mating. (Cantharidin is a toxic defensive compound that serves as a very effective deterrent to predation.) The spermatophores are energetically “expensive” to produce and are transferred to females during relatively short-lived mating aggregations. Mating in some species may take up to 24–48 hours, thus reducing the opportunities for multiple matings, and as a result males of long-mated species end up investing rather heavily in a limited number of females compared to males that mate more often. These features lead to size assortative mating (Alcock & Hanley 1987), with males showing a preference for larger females (that are presumably more fecund) and females preferring larger males to maximize the amount of cantharidin that they receive or to ensure receipt of a spermatophore large enough to fertilize their full complement of eggs. Medium-sized individuals, likewise, would choose the largest of the remaining individuals, leaving the smallest individuals to mate among themselves. Alcock & Hanley (1987) also note, however, that not all species of blister beetles exhibit size assortative mating, even though they form large mating aggregations and individuals vary greatly in size. I have not seen any reference to size assortative mating in Pyrota bilineata; however, this example seems to suggest that the behavior is not practiced by this species. This could be due to shorter mating times (leading to more opportunities for mating) or a range of variation in body size that is not sufficient to consistently favor the behavior.

REFERENCE:

Alcock, J. & N. F. Hadley. 1987. Assortative Mating by Size: A Comparison of Two Meloid Beetles (Coleoptera: Meloidae). Journal of the Kansas Entomological Society 60(1):41–50 [preview].

Copyright © Ted C. MacRae 2013

One-shot Wednesday: The “other” hibiscus jewel beetle

Paragrilus tenuis | Stoddard Co., Missouri

Paragrilus tenuis (LeConte) | Stoddard Co., Missouri

This past summer I visited Otter Slough Conservation Area in southeast Missouri in an effort to find and photograph the stunningly beautiful Agrilus concinnus Horn, or “hibiscus jewel beetle” (MacRae 2004). I was not successful in that quest, but I did manage to snap a single photo of another jewel beetle also associated exclusively with hibiscus, Paragrilus tenuis (LeConte). This species belongs to a much smaller genus of mostly Neotropical jewel beetles that resemble the related and much more speciose genus Agrilus but differ significantly by their antennae being received in grooves along the sides of the pronotum and, for the most part, their association as larvae with stems of living, herbaceous plants rather than dead branches and twigs of deciduous trees. Only four species of Paragrilus occur in the U.S. (Hespenheide 2002), and of these only Ptenuis is known to occur in the eastern U.S. where it has been reported breeding in Hibiscus moscheutos (including ssp. lasiocarpos). I have also collected adults on H. laevis (MacRae 2006), but to my knowledge it has not yet been reared from that plant.

These tiny little beetles (~ 5 mm in length) are normally seen resting on the terminal leaves of their host plants, but they are extremely wary and quick to take flight. As a result, photographing them in situ with a short macro lens in the heat of the day is rather challenging, especially when they are not numerous. I only saw perhaps half a dozen individuals during the visit, and the photo shown here represents the only shot that I even managed to fire off. While I would have liked to have gotten a dorsal view of the beetle, this single shot is nevertheless well-focused and a rather interesting composition.

REFERENCES:

Hespenheide, H. A. 2002. A review of North and Central American Paragrilus Saunders, 1871 (Coleoptera: Buprestidae: Agrilinae). Zootaxa 43:1–28 [pdf].

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 [pdf].

MacRae, T. C. 2006. Distributional and biological notes on North American Buprestidae (Coleoptera), with comments on variation in Anthaxia (Haplanthaxiaviridicornis (Say) and A. (H.) viridfrons Gory. The Pan-Pacific Entomologist 82(2):166–199 [pdf].

Copyright © Ted C. MacRae 2013

Honey Locust Borer

Agrilus difficilis | Beaver Dunes State Park, Beaver Co., Oklahoma

Agrilus difficilis | Beaver Dunes State Park, Beaver Co., Oklahoma

Conditions for collecting didn’t look very promising when I awoke on Day 4 of my early June trip to northwestern Oklahoma. After collecting at Alabaster Caverns State Park the previous day, I had traveled a few hours further west during the evening with plans to collect at Beaver Dunes State Park the following morning. However, heavy rain during the night and lingering sprinkles during the morning had me thinking it might be a lost day. By noon, however, the rain had completely abated, and though the sky still hung low and gray I decided I had nothing to lose by at least trying. I knew quickly that I’d made the right decision, as within minutes of arriving at the park I began seeing jewel beetles (family Buprestidae) landing on my beating sheet. Hackberry (Celtis sp.) was abundant along the roadways and supporting great numbers of individuals in the genera Chrysobothris and Agrilus.

This species is associated almost exclusively with honey locust (Gleditsia triacanthos).

This species is associated almost exclusively with honey locust (Gleditsia triacanthos).

By the time I reached the back end of the campground, I’d collected rather large series of the hackberry associates when I noticed a dying honey locust (Gleditsia triacanthos) tree in one of the campsites. Honey locust (and fabaceous trees, in general) is favored by several species of jewel beetles—at least a dozen species have been recorded in the literature reared from its branches, and another dozen species have been collected on it as adults. As a result, when jewel beetles are active it’s always a good bet that some will be found on honey locust when present, especially if the trees are stressed or dying. I walked up to the tree—a fairly large one—and scanned the lower branches overhead to see if I could notice any activity. I did not, but I nevertheless placed my beating sheet underneath one of the branches, gave the branch a quick “whack” with the handle of my net, and lowered the beating sheet to have a look. To my surprise, I saw at least 50 adults of the jewel beetle species, Agrilus difficilis, sitting on the beating sheet. Because of the cloudy conditions and cool, moist air, the beetles were not very active and did not immediately zip off the beating sheet as they would have had the day been sunnier and warmer, so I was able to rather easily collect a decent series of the beetles without any trouble. I had never seen the beetles so numerous, however, so I continued to beat a few more branches—each yielding just as many adults as the previous. I was astonished by the fact that the beetles were so abundant on the branch, yet I had not seen them even when I specifically looked for the presence of jewel beetles in the branches prior to beating them. Taking another look at the branches, I was able to visually detect just a few individuals, and those only with great difficulty, until I pulled the branch down and was able to look at it up close.

Relatively large size, coppery-purple color without spots on the elytra, and the presence of lateral white patches distinguish this species.

Large size, coppery color, no spots on elytra, and presence of lateral white patches distinguish this species.

Honey locust became a rather popular landscape ornamental tree in the eastern U.S. after the development of thornless cultivars, and while at first the tree seemed to be relatively free of insect pests, A, difficilis has proven to be one of several insects that have adapted to these landscape plants and occasionally cause economic damage. Trees in urban landscapes are often planted in suboptimal sites and suffer from stress to a much greater degree than their native counterparts, and the beetles take advantage of the lowered defensive capabilities of these stressed trees to gain entry. Larvae mine beneath the bark and damage the cambium layer, interfering with movement of water and nutrients. Trees in later stages of attack usually exhibit branch dieback and D-shaped holes in the trunk and main branches where adults have emerged from the tree. In severe cases infestation by this species can result in death of the tree. As mentioned above, there are twelve other species of jewel beetles that have been reared from the wood of honey locust. All of these have been reared only from dead wood rather than living trees, but adults of these species might, nevertheless, be encountered on living trees. They include three species (A. egeniformisA. fallax, and A. pseudofallax) that might be confused with A. difficilis; however, the latter is easily distinguished from these and other congeners by its relatively large size, coppery color with purple luster, absence of any spots or pubescent lines on the elytra, and distinctive patches of white pubescence along the sides. As with most wood boring beetles, chemical control of the adults or larvae is usually not feasible once an infestation has already begun—the best method to avoid infestations in landscape trees is proper site selection and optimal care to prevent stress that reduces the ability of the tree to fend off attack.

Copyright © Ted C. MacRae 2013

How to pack and ship pinned insect specimens

Even though I don’t work in a museum, sending and receiving pinned insects is a routine activity for me. As a collector of beetles with some expertise in their identification, I’ve had opportunity to exchange with or provide IDs to other collectors from around the world. Of course, the extreme fragility of dried, pinned insect specimens makes them vulnerable to damage during shipment, especially when shipped overseas. While properly labeled, pinned insect specimens have no monetary value, the scientific information they represent is priceless, and every attempt should be made to protect them from damage during shipment. Sadly, despite our best efforts damage is sometimes unavoidable, as even packages marked “Fragile” can be subject to rough or careless handling. More often than not, however, I have received shipments in which the contents suffered damage that could have been avoided had the sender paid more attention to packing the shipment in a manner that gave it the best possible chance of arriving safely. Here I offer some general tips on the best way to pack and ship pinned insect specimens for shipment. While these remarks are broadly applicable to pinned insects in general, they are given from the perspective of a someone who collects beetles—specimens of which are relatively small to moderate in size, hard-bodied, and compact in form. Insects from other groups, especially those with large, fragile species such as Lepidoptera and Orthoptera, may require additional precautions to minimize the risk of damage.

  1. Select a sturdy specimen box with a firm pinning bottom. The size of the box should be selected appropriate for the number of specimens—i.e., do not select a large box for only a few specimens or tightly pack too many specimens in too small a box,  Modern polyethylene foams used in pinning trays seem sufficiently firm to hold pinned specimens during shipment as long as they are at least ¼” thick—thicker foams, of course, will hold even more firmly but often “push” the labels on the pinned specimens up against each other, necessitating additional labor to reset them. The box should have a tight-fitting lid that can be set firmly in place. Pin the specimens into the box, making sure the pins are set completely through the foam and taking care not to overpack the specimens within the box too tightly (body parts, especially antennae and tarsi, should never overlap) that could result in damage to them or adjacent specimens during removal. Ideally the specimens should fill the box completely, but if they do not then fill the empty space with blank pins to avoid large, blank areas of foam bottom without pins. Here is an example of a filled specimen box:

    Pinned insects in specimen box ready for packing.

    Pinned insects in specimen box ready for packing.

  2. Use brace pins for large or heavy specimens. This is one of the most common mistakes I see! In the example above, several of the larger species are surrounded by brace pins to keep them from rotating on their pins and damaging neighboring specimens. At least two pins should be used—I place them against the elytra on each side behind the hind legs, and very long or heavy specimens should be further braced by additional pins on each side of the thorax to further ensure they are fully immobilized. Although not shown in this example, specimens with very heavy heads (large mandibles, etc.) should be even further immobilized with additional pins at the head. Here is a closeup view of some of the specimens in the above box that have been further secured with brace pins:

    Large specimens are further immobilized with brace pins.

    Large specimens are further immobilized with brace pins.

  3. Use an inner lid with padding to hold it firmly against the specimens. An inner lid lies on top of the specimens underneath the specimen box lid to keep the specimens securely seated in the foam and prevent them from “working” their way out. Some specimen boxes designed for shipping, such as the examples shown in these photos, come with an inner lid that is hinged on a long side. If the specimen box lacks an inner lid, one should be fashioned from cardboard or heavy card stock. The advantage of an attached inner lid is that it will not move inside the box, so if an inner lid must be fashioned it is essential to trim it so that it fits precisely within the box to minimize the potential for movement. I like to draw an outline on the cardboard with the specimen box and cut on the lines, then shave off extra material from each side to shape it to the inside perimeter of the box. Either way, make a “pull tab” out of adhesive tape and attach it to the inner lid to allow easy removal during unpacking. If the inner lid when set in place does not seat firmly against the outer lid, extra padding material such as paper towels should be placed on top of the inner lid to ensure that it sits firmly against the specimens when the outer lid is set in place. The specimen box with inner lid in place, pull tab attached, and extra padding placed on top is shown below:

    Cover the inner lid with padding to secure it firmly against the specimens.

    Cover the inner lid with padding to secure it firmly against the specimens.

  4. Seal closed specimen box with tape or rubber bands. The outer lid of the specimen box should be secured in place so that it does not “work” its way loose. Some people use tape, which is effective but must be cut if the box is opened for inspection, leaving the lid unsecured afterwards. I prefer to use sturdy rubber bands, which can be removed for inspection and then easily replaced afterwards. Some specimen boxes come equipped with metal tabs or hoops that fit through slots on the outer lid and that can be bent over to secure the lid in place. In my experience, these often break off after repeated use, so rubber bands or tape are a good insurance policy for such boxes. Another common practice is to wrap specimen boxes in packing paper or place them inside plastic, Zip-Lock bags. This was necessary in the days when excelsior shavings were often used as a packing material around the specimen box, which contained shavings that could work their way into the specimen box and cause damage. With the ready availability of modern packing materials such as foam peanuts there should no longer be any reason to use excelsior shavings. Still, wrapping or sealing inside a plastic bag can’t hurt if it is desired. A closed specimen box with rubber bands securely in place is shown in the photo below:

    Specimen box sealed with rubber bands

    Specimen box sealed with rubber bands

  5. Place an address label on the specimen box. This will ensure that the shipment does not get tossed into the “dead mail” pile if the outer address label is lost or destroyed (I’ve left the label off in these examples to ensure privacy of the recipient).
  6. Secure multiple specimen boxes tightly together. If multiple specimen boxes are shipped together, they should be secured tightly together so that they cannot “bump” into each other during shipment. As mentioned before, tape works but might end up being cut for inspection, so I prefer to use large rubber bands. String can also be used to tie the boxes together, but unless the inspection agent is handy with knots the boxes may not get tied back together. The two specimen boxes included in the shipment I used for this example, secured tightly together, are shown below:

    Multiple boxes should be bound tightly together.

    Multiple boxes should be bound tightly together.

  7. Pack specimen box inside an oversized shipping boxShipping box size selection is critical! The shipping box should not only be sturdy but also big enough to accommodate specimen boxes with at least 3–4 inches below and 2–3 inches on top and each side of the specimen box. This space is necessary to allow the packing material to function not only as cushioning but also in “shock absorption.” My preferred packing material is foam peanuts, since it doesn’t settle during shipment and the amount used can be tailored precisely to the needs of an individual box. The photo below shows the pinning boxes resting on a 4-inch layer of foam peanuts with at least 2–3 inches of space on the sides and above:

    Place specimen boxes inside a sturdy shipping box with plenty of room on all sides.

    Place specimen boxes inside a sturdy shipping box with plenty of room on all sides.

  8. DO NOT OVERPACK! This is the most common mistake people make! The packing material needs to serve two purposes: 1) provide a crush zone to protect from direct damage, and 2) provide shock absorption to protect from damage by impact jarring. The specimen box actually needs to be able to move slightly within the closed shipping box. If it cannot, energy from impacts is transmitted in full to the specimens inside, greatly increasing the risk that heavier body parts (especially the head/pronotum) will be jarred off the specimens. This not only results in damage to the broken specimen, but the dislodged body parts then act as “wrecking balls” that bounce and tumble inside the specimen box, destroying all of the specimens within their reach. After placing a 3–4-inch layer of packing in the bottom of the shipping box, I like to set the specimen box(es) on top of the foam in the center of the shipping box and fill the shipping box with additional foam peanuts to within about 1″ of the top. Avoid the temptation to fill the box to the brim, or to “settle” the foam peanuts and add a few more, as this will result in a tightly packed box that does not protect the specimens as well as a more loosely packed box. To test, close the flaps on top of the box and give the box a light up-and-down “shake”—you should feel the specimen box bounce slightly inside. If it does not, remove a small amount of packing peanuts and repeat the test. If you cannot remove enough packing peanuts without exposing the top of the specimen box inside, your shipping box is too small and you should select a larger size. The photo below shows the shipping box filled with packing peanuts to the proper level:

    Shipping box ''almost'' filled with packing material.

    Shipping box ”almost” filled with packing material.

  9. Label the package “FRAGILE”. Whether this is actually helpful or invites abuse by some passive aggressive handler is a matter of debate, but I am of the opinion that a majority of shipping personnel will actually treat the package with a little more respect if they see this label, especially with the disclosure that the contents are preserved insects with no commercial but extreme scientific value. Additionally, disclosure of such information may actually be required by some destination countries, so it’s a good idea to label packages as a matter of routine practice. I like to place one label on top of the shipping box and additional labels on all four sides. BioQuip Products sells moisture-activated adhesive labels as shown below, or similar labels can be designed in a word processing program and printed on blank adhesive labels; however, the latter should be covered with clear tape to prevent them from peeling off of the shipping box during transit.

    Place a fragile sticker on top and all four sides.

    Place a fragile sticker on top and all four sides.

Much of what I have written here I learned as a graduate student, based on a much more detailed article by Sabrosky (1971) that provides additional suggestions for extremely rare and valuable specimens, advice regarding the different postal classes available for international shipments, and a list of “Don’ts” under any circumstances.

Disclaimer: I am an amateur—albeit a highly practiced one, and there may be additional suggestions or advice from professional collection managers and museum curators that would be highly welcomed in the comments below  should it be offered.

REFERENCE:

Sabrosky, C. W. 1971. Packing and shipping pinned insects. Bulletin of the Entomological Society of America 17(1):6–8 [preview].

Copyright © Ted C. MacRae 2013