New Curriculum Kits


Virginia Tech Insect Collection Curriculum Kit

Last Saturday at Hokie Bugfest, we launched our Curriculum Kits (above). These kits contain material for incorporating insect biodiversity into your K-12 science curriculum. The contents include insect specimens (butterfly, beetle, fly, wasp, and katydid), a Golden Guide to Insects, a Rite-in-the-rain field notebook, collection vials, 5x and 10x magnifying lenses, a VTEC sticker, and a companion field guide. The companion field guide follows the Virginia Science Standards of Learning for grades K-12, and many of the activities in the guide are adaptable to the different grade levels (download here).

If you are K-12 teacher in Virginia interested in including insect biodiversity into your curriculum, the kits are available on loan, and contact Dr. Paul Marek for more details (email: pmarek@vt.edu).

The field guide was adapted from F. Taylor, A. Raflo, L. Sharp (1997) Model Inquiries into Nature in The Schoolyard, The MINTS: Book: An Inquiry Field Guide to the Natural History of Southwestern Virginia Schoolyards (CC-BY-NC-SA 4.0)

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A checklist of bee species in the Virginia Tech Insect Collection


Sweat bee, Augochlora pura mosieri Cockerell, 1922 (family Halictidae)

By Katie Williamson, Derek Hennen, and Paul Marek

What’s one of the first things that comes to your mind when describing a bee? A fuzzy, flying insect? Pollination? Or even how the insect is capable of a rather feisty and painful sting? A member of the order Hymenoptera (“bees, ants, wasps, and hornets”) and the superfamily Apoidea, these mostly-vegetarian bees originated from a lineage of carnivorous, meat-eating wasps such as Cicada Killers, Jewel Wasps, and Mud Daubers. While social bees, such as bumble bees and honey bees, are well known, many species of bees are solitary and do not socialize as much with each other. So what makes a bee a bee? And how diverse are bees? Understanding the evolutionary history of this group can answer these questions.

There are seven families of bees (Andrenidae, Halictidae, Apidae, Mellitidae, Colletidae Megachilidae, and Stenotritidae) and 20,000 known species. Bees are estimated to have first emerged during the Cretaceous Period, and since then, have speciated into tens of thousands of species, each with a morphology that matches their environment. For example, while a Bumble bee (Apidae) may be completely covered with branched hairs, a Sweat bee (Halictidae) possesses minute branched hairs that are nearly impossible to see unless one examines the bee up close. Their branched hairs, a major feature that distinguishes bees, trap pollen grains thereby allowing the bee to gather and transport this protein-rich source of food to their nest. Some bees even have a special built-in pollen basket called a corbicula that they pack with pollen, and still other bees such as the cuckoo bees, have lost pollen collecting structures and evolved to steal other’s pollen.


Cuckoo bee, Nomada superba Cresson, 1863 (family Apidae)

Here at the Virginia Tech Insect Collection, we have almost 200 identified species of bees and over 2,000 specimens. We have been collecting bees for over a century, and the oldest specimen in the collection is a Carpenter bee from 1902. Most of the specimens were collected in the United States, particularly Appalachia, and are in the family Apidae, but other bee families are well-represented in our natural history collection. One notable species is the Rusty-patched Bumble bee (Bombus affinis), whose key defining feature is a distinct, orange (rusty) patch of hair on the insect’s abdomen. The specimens in the collection were collected before its recent designation as an endangered species, and can be used in the future by other scientists researching the biology and conservation of the species. Indeed, DNA from specimens preserved in natural history collections have been used to study and mitigate population declines in bees.

All the bee species in the Virginia Tech Insect Collection are listed below, organized by family. Links associated with each species lead to the Symbiota Collections of Arthropods Network, an online database that provides associated collection data and photographs for each bee specimen in the collection. The bees in the VTEC are a result of a century of hard work by entomologists who collected, prepared, and identified specimens in the collection.


Squash bee, Peponapis pruinosa Say, 1837 (family Apidae)

By studying the bee specimens preserved in the VTEC, researchers are able to answer scientific questions about evolution, morphology, and ecology. Indeed, bees in our collection even retain the pollen that they gathered from the last flower they visited. Palynologists (scientists who study pollen) are able to identify plants from looking at the pollen stuck to the bee. These studies further enhance not only our understanding of bees, but what factors affect their behavior, evolution, and ecological contributions of these amazing animals.

ANDRENIDAE, the Mining bees
Andrena accepta Viereck, 1916
Andrena aliciae Robertson, 1891
Andrena alleghaniensis Viereck, 1907, “Alleghany Andrena”
Andrena barbara Bouseman & LaBerge, 1979, “Barbara’s Miner”
Andrena bisalicis Viereck, 1908
Andrena brevipalpis Cockerell, 1930
Andrena carlini Cockerell, 1901, “Carlin’s Miner”
Andrena carolina Viereck, 1909
Andrena commoda Smith, 1879, “Advantaged Miner Bee”
Andrena confederata Viereck, 1917, “Southern Miner”
Andrena cornelli Viereck, 1907, “Azalea Miner”
Andrena crataegi Robertson, 1893, “Hawthorn Andrena”
Andrena cressonii Robertson, 1891, “Cresson’s Andrena”
Andrena dunningi Cockerell, 1898, “Dunning’s Miner”
Andrena erigeniae Robertson, 1891, “Spring Beauty Andrena”
Andrena fenningeri Viereck, 1922
Andrena forbesii Robertson, 1891, “Forbe’s Andrena”
Andrena heraclei Robertson, 1897
Andrena hippotes Robertson, 1895
Andrena illini Bouseman & LaBerge, 1979
Andrena imitatrix Cresson, 1872
Andrena mariae Robertson, 1891
Andrena miserabilis Cresson, 1872, “Miserable Andrena”
Andrena nasonii Robertson, 1895, “Nason’s Andrena”
Andrena perplexa Smith, 1853
Andrena personata Robertson, 1897
Andrena pruni Robertson, 1891, “Cherry Miner”
Andrena rugosa Robertson, 1891, “Rugose Andrena”
Andrena sayi Robertson, 1891
Andrena spiraeana Robertson, 1895
Andrena tridens Robertson, 1902
Andrena vicina Smith, 1853, “Neighborly Miner”
Andrena violae Robertson, 1891, “Violet Miner”
Andrena wilkella (Kirby, 1802), “Wilke’s Mining Bee”
Calliopsis andreniformis Smith, 1853
Pseudopanurgus Cockerell, 1897

HALICTIDAE, the Sweat bees
Agapostemon radiatus (Say, 1837)
Agapostemon sericeus (Förster, 1771), “Silky Striped-Sweat Bee”
Agapostemon splendens (Lepeletier, 1841), “Brown-Winged Striped-Sweat Bee”
Agapostemon texanus Cresson, 1872, “Texas Striped-Sweat Bee”
Agapostemon virescens (Fabricius, 1775), “Bicolored Striped-Sweat Bee”
Augochlora pura (Say, 1837), “Pure Green Augochlora”
Augochlora pura mosieri Cockerell, 1922
Augochlora pura pura (Say, 1837)
Augochlorella aurata (Smith, 1853)
Augochlorella gratiosa (Smith, 1853)
Augochlorella persimilis (Viereck, 1910)
Augochlorella striata (Provancher, 1888)
Augochloropsis metallica (Fabricius, 1793)
Augochloropsis metallica metallica (Fabricius, 1793)
Augochloropsis sumptuosa (Smith, 1853)
Dialictus Robertson, 1902
Halictus confusus Smith, 1853, “Confusing Metallic-Furrow Bee”
Halictus ligatus Say, 1837, “Ligated Furrow Bee”
Halictus parallelus Say, 1837, “Parallel Furrow Bee”
Halictus rubicundus (Christ, 1791), “Orange-Legged Furrow Bee”
Lasioglossum anomalum (Robertson, 1892), “Anomalous Metallic-Sweat Bee”
Lasioglossum apocyni (Mitchell, 1960)
Lasioglossum arcanus (Sandhouse, 1924)
Lasioglossum bruneri (Crawford, 1902), “Bruner’s Dialictus”
Lasioglossum callidum (Sandhouse, 1924)
Lasioglossum coeruleum (Robertson, 1893)
Lasioglossum coreopsis (Robertson, 1902)
Lasioglossum coriaceum (Smith, 1853)
Lasioglossum cressonii (Robertson, 1890) “Cresson’s Dialictus”
Lasioglossum forbesii (Robertson, 1890)
Lasioglossum foxii (Robertson, 1895)
Lasioglossum fuscipenne (Smith, 1853)
Lasioglossum illinoense (Robertson, 1892)
Lasioglossum imitatum (Smith, 1853)
Lasioglossum leucozonium (Schrank, 1781)
Lasioglossum lineatulum (Crawford, 1906), “Lineated Metallic-Sweat Bee”
Lasioglossum lustrans (Cockerell, 1897)
Lasioglossum macoupinense (Robertson, 1895)
Lasioglossum nelumbonis (Robertson, 1890)
Lasioglossum obscurum (Robertson, 1892)
Lasioglossum pectorale (Smith, 1853)
Lasioglossum pilosum (Smith, 1853)
Lasioglossum platyparium (Robertson, 1895)
Lasioglossum pruinosum (Robertson, 1892), “Pruinose Metallic-Sweat Bee”
Lasioglossum quebecense (Crawford, 1907)
Lasioglossum sopinci (Crawford, 1932)
Lasioglossum tegulare (Robertson, 1890), “Epaulette Metallic-Sweat Bee”
Lasioglossum truncatum (Robertson, 1901)
Lasioglossum versans (Lovell, 1905)
Lasioglossum versatum (Robertson, 1902)
Lasioglossum viridatum (Lovell, 1905)
Lasioglossum zephyrum (Smith, 1853), “Zephyr Dialictus”
Sphecodes banksii Lovell, 1909
Sphecodes heraclei Robertson, 1897
Sphecodes ranunculi Robertson, 1897

COLLETIDAE, the Masked or Plasterer bees
Colletes americanus Cresson, 1868
Colletes compactus Cresson, 1868, “Aster Cellophane Bee”
Colletes eulophi Robertson, 1891
Colletes inaequalis Say, 1837, “Unequal Cellophane Bee”
Colletes latitarsis Robertson, 1891, “Broad-footed Cellophane Bee”
Colletes thoracicus Smith, 1853, “Rufous-Chested Cellophane Bee”
Hylaeus affinis (Smith, 1853), “Eastern Masked Bee”
Hylaeus mesillae (Cockerell, 1896), “Mesilla Masked Bee”
Hylaeus modestus Say, 1837, “Modest Masked Bee”

MEGACHILIDAE, the Leafcutter or Mason bees
Anthidiellum Cockerell, 1904, “Rotund Resin Bees”
Anthidium maculifrons Smith, 1854, “Spot-Fronted Wool-Carder Bee”
Anthidium manicatum (Linnaeus, 1758), “European Woolcarder”
Anthidium oblongatum (Illiger, 1806), “Oblong Woolcarder”
Coelioxys moesta Cresson, 1864
Coelioxys octodentata Say, 1824, “Eight-toothed Cuckoo-leaf-cutter”
Coelioxys rufitarsis Smith, 1854, “Red-footed Cuckoo-leaf-cutter”
Coelioxys sayi Robertson, 1897, “Say’s Cuckoo-leaf-cutter”
Coelioxys slossoni Viereck, 1902
Heriades carinatus Cresson, 1864
Heriades variolosus (Cresson, 1872)
Hoplitis pilosifrons (Cresson, 1864)
Hoplitis producta (Cresson, 1864)
Hoplitis spoliata (Provancher, 1888)
Lithurgus Berthold, 1827
Megachile brevis Say, 1837, “Common Little Leaf-cutter Bee”
Megachile campanulae (Robertson, 1903), “Bellflower Resin Bee”
Megachile frigida Smith, 1853, “Frigid Leafcutter”
Megachile gemula Cresson, 1878, “Small-handed Leaf-cutter Bee”
Megachile inimica sayi Cresson, 1878, “Hostile Leaf-cutter Bee”
Megachile latimanus Say, 1823, “Broad-handed Leafcutter Bee”
Megachile mendica Cresson, 1878, “Flat-tailed Leaf-cutter Bee”
Megachile onobrychidis Cockerell, 1908, “Western Little Leaf-cutter Bee”
Megachile perihirta Cockerell, 1898, “Western Leafcutter Bee”
Megachile petulans Cresson, 1878, “Petulant Leaf-cutter Bee”
Megachile pugnata Say, 1837, “Pugnacious Leaf-cutter Bee”
Megachile relativa Cresson, 1878, “Golden-tailed Leafcutter Bee”
Megachile rotundata (Fabricius, 1793), “Alfalfa Leafcutter Bee”
Megachile texana Cresson, 1878, “Texas Leaf-cutter Bee”
Megachile townsendiana Cockerell, 1898
Osmia albiventris Cresson, 1864
Osmia atriventris Cresson, 1864, “Blueberry Bee”
Osmia bucephala Cresson, 1864, “Bufflehead Mason Bee”
Osmia conjuncta Cresson, 1864
Osmia cornifrons (Radoszkowski, 1887), “Horned-face Bee”
Osmia georgica Cresson, 1878
Osmia lignaria Say, 1837, “Blue Orchard Mason Bee”
Osmia lignaria lignaria Say, 1837
Osmia pumila Cresson, 1864
Osmia subfasciata Cresson, 1872
Osmia taurus Smith, 1873, “Taurus Mason Bee”
Osmia texana Cresson, 1872, “Texas Osmia”
Osmia virga Sandhouse, 1939
Paranthidium jugatorium (Say, 1824)
Stelis lateralis Cresson, 1864

APIDAE, the Cuckoo, Carpenter, Digger, Bumble, and Honey Bees
Anthedonia compta (Cresson, 1878)
Anthophora abrupta Say, 1837, “Abrupt Digger Bee”
Anthophora bomboides Kirby, 1837, “Bumble-bee Digger”
Anthophora bomboides bomboides Kirby 1837
Anthophora pacifica Cresson, 1878, “Pacific Digger”
Anthophora terminalis Cresson, 1869, “Orange-tipped Wood-digger”
Anthophora urbana Cresson, 1878, “Urbane Digger”
Anthophora ursina Cresson, 1869, “Ursine Digger”
Apis mellifera Linnaeus, 1758, “European honey bee”
Bombus affinis Cresson, 1863, “Rusty patched bumble bee”
Bombus ashtoni (Cresson, 1864), “Ashton’s Cuckoo Bumblebee”
Bombus auricomus (Robertson, 1903), “Black-and-gold Bumble Bee”
Bombus bimaculatus Cresson, 1863, “Two-spotted Bumble Bee”
Bombus borealis Kirby, 1837, “Northern Amber Bumble Bee”
Bombus citrinus (Smith, 1854), “Lemon Cuckoo Bumble Bee”
Bombus fernaldae (Franklin, 1911), “Fernald’s Cuckoo Bumble Bee”
Bombus fervidus (Fabricius, 1798), “Golden Northern Bumble Bee”
Bombus griseocollis (DeGeer, 1773), “Brown-belted Bumble Bee”
Bombus impatiens Cresson, 1863, “Common Eastern Bumble Bee”
Bombus nevadensis Cresson, 1874, “Nevada Bumble Bee”
Bombus pensylvanicus (De Geer, 1773), “American Bumble Bee”
Bombus perplexus Cresson, 1863, “Perplexing Bumble Bee”
Bombus sandersoni Franklin, 1913, “Sanderson’s Bumble Bee”
Bombus vagans Smith, 1854, “Half-black Bumble Bee”
Bombus variabilis (Cresson, 1872), “Variable Cuckoo Bumble Bee”
Bombus vosnesenskii Radoszkowski, 1862, “Yellow-faced Bumble Bee”
Ceratina calcarata Robertson, 1900, “Spurred Ceratina”
Ceratina dupla Say, 1837, “Doubled Ceratina”
Ceratina strenua Smith, 1879, “Nimble Ceratina”
Eulaema Lepeletier, 1841
Habropoda laboriosa (Fabricius, 1804), “Blueberry Digger”
Holcopasites calliopsidis (Linsley, 1943)
Melissodes agilis Cresson, 1878, “Agile Long-horned Bee”
Melissodes bidentis Cockerell, 1914
Melissodes bimaculata (Lepeletier, 1825), “Two-spotted Longhorn”
Melissodes bimaculata bimaculata (Lepeletier, 1825)
Melissodes communis Cresson, 1878, “Common Long-horned Bee”
Melissodes denticulata Smith, 1854
Melissodes fimbriata Cresson, 1878
Melissodes rustica (Say, 1837)
Melissodes trinodis Robertson, 1901
Melitoma taurea (Say, 1837), “Chimney Bee”
Nomada articulata Smith, 1854
Nomada composita Mitchell, 1962
Nomada imbricata Smith, 1854
Nomada luteoloides Robertson, 1895
Nomada maculata Cresson, 1863, “Spotted Nomad Bee”
Nomada superba Cresson, 1863, “Superb Nomada”
Nomada vegana Cockerell, 1903
Peponapis pruinosa (Say, 1837), “Pruinose Squash Bee”
Psithyrus Lepeletier, 1833, “Cuckoo Bumble Bees”
Ptilothrix bombiformis (Cresson, 1878), “Hibiscus Bee”
Svastra obliqua (Say, 1837), “Sunflower Bee”
Tetralonia Spinola, 1839
Triepeolus lunatus (Say, 1824), “Cuckoo Bee”
Triepeolus remigatus (Fabricius, 1804)
Triepeolus simplex Robertson, 1903
Xenoglossa strenua (Cresson, 1878), “Large Squash Bee”
Xylocopa tabaniformis Smith, 1854, “Horsefly-like Carpenter Bee”
Xylocopa virginica (Linnaeus, 1771), Eastern Carpenter Bee”

Further reading about native bee diversity in the eastern United States

Appalachian Beekeeping Collective
Bees of Blandy Experimental Farm, Virginia
Center for Biological Diversity, Native Bee Diversity
Couvillon Bee Lab at Virginia Tech
Discover Life’s Bee Identification
James Wilson, Extension Apiculturist at Virginia Tech
Native Bee Inventory and Monitoring Lab
Native Bees, Jha Lab, University of Texas at Austin
New York Times, Plan Bee: The Rise of Alternative Pollinators
O’Rourke Lab at Virginia Tech
University of Minnesota, Bee Lab

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Curiosities from the cabinets: Lycid beetle mimicry ring


Lycid beetle mimicry ring (A. Lucidota, Lampyridae; B. Limonia, Limoniidae; C. Chaetodactyla, Ptilodactylidae; D. Mesopteron, Lycidae; E. Austrolimnophila, Limoniidae; F. Calopteron, Lycidae; G. Correbidia, Erebidae; H. Calopteron, Lycidae; I. Correbia, Erebidae; J. Lycomorpha, Erebidae; K. Correbidia, Erebidae). Scale bar: 5 mm.

A few months ago, while browsing the unsorted material in the VTEC, Grant and I discovered this group of orange and black insects collected from a small patch of rainforest near Turrialba, Costa Rica, by Dr. Robin Andrews (Department of Biological Sciences) in 1966. The beetles, flies and moths are distantly related, and yet they appear similar with the same orange and black color pattern. The similarity is not by chance, but a result of convergent evolution by mimicry.

The insects are members of a mimicry ring with lycid beetles as the model species. Also known as net-winged beetles, because the pronounced veins in their elytra give a net-like appearance, lycids are toxic and defended with pyrazines and lycidic acid (Moore et al., 1990; Eisner et al., 2008). The veins, which are turgid and easily ruptured, contain the toxins that spill out when the beetle is handled by a predator. It’s been shown experimentally that the beetles are rejected by spiders, lizards, and birds (Darlington, 1938; Eisner et al., 2008).

Historically, it’s likely the lycid beetles were on the (evolutionary) scene first. Over time predators learned to associate their distinctive coloration with being noxious, a.k.a. “aposematic”. The predators’ perception of prey as toxic may also include the smell of the beetle too since pyrazine is aromatic. Quoting Eisner et al. (2008), the implicit message of pyrazine may be, “desist, lest you are willing to risk an unpleasant mouthful”. This cocktail of defense chemicals including a mixture of a highly toxic chemical and an aromatic chemical is also known in other insects (Gullan & Cranston, 2014).

Later in time, those flies and moths that tended to appear more similar to the lycids persisted in nature. If the lycids and their aposematic signal was so effective at deterring predators then other species—even if they were non-toxic—would converge as a result of mimicry.

It is interesting to note the two larger moths labelled G and K have very accurate coloring to match the beetles including a black stripe in the middle of an orange pronotum; whereas the smaller moth, labeled I, is less precise because it has an orange stripe on a black pronotum. This is because predators are less likely to attack smaller mimics as they are less profitable as prey items (Penney et al., 2012). Therefore predators exert less selection for accuracy of mimic prey.

References

Darlington, P. J. (1938). Experiments on mimicry in Cuba, with suggestions for future study. Ecological Entomology, 87(23), 681-695.

Eisner, T., Schroeder, F. C., Snyder, N., Grant, J. B., Aneshansley, D. J., Utterback, D., Meinwald, J., & Eisner, M. (2008). Defensive chemistry of lycid beetles and of mimetic cerambycid beetles that feed on them. Chemoecology, 18(2), 109-119.

Gullan, P. J., & Cranston, P. S. (2014). The insects: an outline of entomology. John Wiley & Sons.

Moore, B. P., Brown, W. V., & Rothschild, M. (1990). Methylalkylpyrazines in aposematic insects, their hostplants and mimics. Chemoecology, 1(2), 43-51.

Penney, H. D., Hassall, C., Skevington, J. H., Abbott, K. R., & Sherratt, T. N. (2012). A comparative analysis of the evolution of imperfect mimicry. Nature, 483(7390), 461-464.

 

 

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Bressler longhorned beetle collection


The longhorned beetle, Typocerus zebra (Olivier, 1795)

In May, Dr. Ellen Brown of Fredericksburg, Virginia donated the collection of her husband Dr. Barry Lee Bressler (1936 – 2017) to the Virginia Tech Insect Collection. Dr. Bressler, who was born in Reading, Pennsylyvania, was a retired Naval Surface Warfare Center (NSWC) physicist and adjunct professor of physics at Virginia Tech. Dr. Bressler graduated from Virginia Tech with a PhD in physics for his work on Fermion-boson couplings.

Bressler, an ardent naturalist, collected beetles and other insects in eastern Virginia. His collection is primarily made up of longhorned beetles of the family Cerambycidae, but other taxa are represented too including weevils, ants, and flies. The collection represents a valuable synoptic collection of the family Cerambycidae in eastern Virginia. Of the more than 1,000 specimens, there are several uncommonly encountered and rare specimens including Saperda imitans Felt & Joutel, 1904; Centrodera sublineata LeConte, 1862; and Lepturges pictus (LeConte, 1852).

Student curator in the collection, Grant Schiermeyer, who is curating the collection, identifying species, and incorporating the material into the VTEC states, “this collection of cerambycids contains a great amount of the diversity of longhorn beetles in this eastern US and will be a significant addition to the VTEC by adding specimens that are underrepresented or rare.”

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Rusty patched bumble bee


Bombus affinis Cresson, 1863, the Rusty patched bumble bee (#VTEC000000590)

On Tuesday, the U.S. Fish and Wildlife Service designated the Rusty patched bumble bee as endangered under the Endangered Species Act. The specimen pictured above is one of 31 individuals that we have databased in the Virginia Tech Insect Collection. The Endangered Species Act of 1973 (ESA) implements means to protect and conserve species. The ESA is important not only because it works to protect individual species, but by doing so it also works to look after the habitat in which the species lives.

The Rusty patched bumble bee is a native pollinator, once widespread through the eastern United States, and over the past 20 years entomologists have found that it has declined over 90% of its previous distribution. This is alarming not only because the species is an unique and irreplaceable piece of our country’s natural heritage, but because pollinators provide us with free pollination services. Bumble bees allow plants to flower, produce fruit and seeds for the next generations of life. Unfortunately, pollinators including the Rusty patched bumble bee are rapidly declining primarily as a result of habitat loss. The native meadows, forests, and homes of the bees are vanishing.

Protection of the Rusty patched bumble bee and its habitat where it lives is important. The process of listing an endangered species is lengthy and requires detailed study demonstrating the species habitat or range is threatened, has been over-utilized by humanity (e.g. over-harvested), highly impacted from disease or predation, or other human-made factors that affect the species very existence.

Unfortunately, the “rusty” red patches on the back of the bumble bee fade over time, which are faintly visible in the specimen pictured above. The U.S. Geological Survey’s Bee Inventorying and Monitoring Lab has some wonderful images of B. affinis below where you can see the reddish patches on the bee’s abdomen (images CC-Zero by Sam Droege).

Bombus affinis, F, side, sky meadows, virginia_2014-09-22-18.05.02 ZS PMax
Bombus affinis, F, Sky meadows sp, virginia, back_2014-09-22-17.48.35 ZS PMax
Bombus affinis, F, face, big sky meadows, virginia_2014-09-22-17.57.06 ZS PMax

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Zygaena angelicae


A zygaenid moth, Zygaena angelicae Ochsenheimer, 1808

Specimen #VTEC000000632 is a handsome zygaenid moth from Mnichovice (Czech Republic) collected by V. Kubelka. Thanks to  volunteer Charity Hall for digitizing several trays of zygaenid and arctiine moths!

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Professor Smyth’s Butterflies

IMG_4790s
Papered insects of Ellison Smyth, founding head of VT Biological Sciences

Did you know that the first football coach of Virginia Tech, Professor Ellison Smyth, was also a butterfly collector? In fact, Professor Smyth was also one of the first instructors in Biological Sciences, and later the first dean of the faculty at Virginia Tech (1902-06).

Professor Smyth grew up in Charleston, South Carolina, where as a boy enjoyed being outdoors collecting butterflies and birds. As a 10 year old, he amassed a collection of birds, butterflies and other insects. The collector’s bug (so to speak) had already taken hold, and he was hooked on natural history. After completing an undergraduate degree, he enrolled in law school at the University of Virginia. Smyth was hired as a clerk at a law firm in Charleston, South Carolina, and later as a partner. However, he admitted that pursuing law was only to satisfy his folks and later quit the firm to pursue biology completely. The lifelong love of biology and natural history was too strong to ignore! Smyth was hired as an adjunct professor at the University of South Carolina and then a professor of biology at Virginia Tech where he taught until 1925. (From the UNC Herbarium, which holds some plant specimens collected by Smyth in the late 1800s.)

I recently found some of Professor Smyth’s butterfly specimens stuffed away in an old drawer in the Virginia Tech Insect Collection. These are commonly known as “papered” specimens. Within each folded paper triangle is a single butterfly or other large winged insect (e.g., dragonfly or katydid) that was placed directly into the paper once collected. The paper helps keep the wings in good shape for pinning and spreading the specimen later on. Because many of the triangles were made from newspapers, there are some classic old-timey ads on some of them!

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A Diana fritillary, Speyeria diana (Cramer, 1777) collected by Smyth in 1907

I’ve peeked in a few of the paper envelopes and there are some spectacular butterflies that await preparation. While mostly local Appalachian species, for example the rare Diana fritillary (above), there are a number of exotic butterflies that Smyth collected in Africa and traded with colleagues. I am now carefully pinning and spreading Professor Smyth’s collection, and will share the discoveries that I find in a following post.

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Milkweed pollinia

Back in April I posted an image of a Sphex sp. wasp with some odd structures on its legs. At the time I suspected the weird things were fungal growths. After some additional research (i.e., I found a photo of a bee with similar things on its legs while surfing the Internet), it seems that those structures are milkweed pollinia and not parasitic growths.

Milkweeds (Asclepias spp.) are herbaceous perennials that tend to have brightly colored clusters of flowers with abundant nectar. There are over 100 described species and subspecies of milkweed in North America. The monarch butterfly, Danaus plexippus (L.), may be the most well-known visitor to milkweed flowers, but milkweeds attract a large suite of butterflies, flies, beetles, bees, and wasps. Many people grow milkweeds for pollinators and other beneficial insects that feed on the plant’s copious nectar.

Asclepias_syriaca_-_Common_Milkweed
Common milkweed (Asclepias syriaca L.)
By Fritzflohrreynolds (Own work)
[CC BY-SA 3.0], via Wikimedia Commons.

Unlike most flowers, milkweeds do not produce loose pollen. Waxy masses of milkweed pollen are grouped into sacs called pollinia. Orchids are the only other group of plants known to use pollinia. Bees don’t collect pollinia to use as food for their larvae the way they do with the loose pollen of other flowers. Nectar is a pollinator’s only reward for visiting milkweed flowers, but there’s a lot of nectar to be had on milkweed. The design of the milkweed flower is all about attracting pollinators with nectar and ensuring that the pollinators pick up and move pollinia to a different flower to complete pollination. Let’s take a look at the unusual structure of the milkweed flower to understand that process.

Each individual milkweed flower has an attractive star-shaped corona on top of a short central column and with outward flaring petals below the column. In the photo below of common milkweed (Asclepias syriaca L.), the corona is a lighter pink than the lower petals. Insects will find nectar on the top of the flower in those stars.

Common_Milkweed_(1035856056)
Closeup of common milkweed (Asclepias syriaca L.)
flowers. By Jason Hollinger (Common Milkweed
Uploaded by Amada44) [CC BY 2.0], via Wikimedia Commons.

The botanical illustration below shows a milkweed flower from a lateral perspective. The star-shaped corona of a single flower (A) has five hoods (h) with a  corolla (c) of lower reflexed petals. A closeup of a flower (B) shows a slit-like opening (f) between each hood formed by the fusion of the anthers at their top and bottoms. Each slit leads to the stigmatic chamber, where the pollinia are housed. Each pollinium (C) is a paired, winged structure with a central body (d).

Entomology_-_with_special_reference_to_its_biological_and_economic_aspects_(1906)_(14781610264)
Botanical illustration of individual milkweed flower.
By Internet Archive Book Images [No restrictions]
via Wikimedia Commons.

A visiting insect trying to reach the nectar offered at the top of the star-shaped corona will slip one of its legs or another appendage inside the anther slits between the hoods. The pollinia inside the stigmatic chamber sticks to the insect’s setae or tarsal claws. By pulling its leg out of the slit, the insect extracts the pollinia and carries them off to another milkweed flower. Again the insect’s leg or another appendage will likely slip inside a slit while feeding. The flower will be pollinated successfully if the donor pollinia remain in the recipient anther slit. [For more information on this process and in greater detail, see Betz et al. (1994) and Borders and Lee-Mäder (2014). Both Eye on Nature and Robert Klip at Ohio State have nice blog posts on milkweed pollination and great closeup photos of pollinia, too.]

Melissodes_trinodus,_f,_foot,_polynia_of_milkweek,_aa_co,_md_2014-07-09-19.33.21_ZS_PMax_(14436615847)
Closeup of a Melissodes trinodis Robertson (Hymenoptera: Apidae)
leg with attached pollinia. By USGS Bee Inventory and Monitoring
Lab from Beltsville, Maryland, USA [CC BY 2.0], via Wikimedia Commons.

Any insect large enough to acquire and transport the pollinia can serve as an efficient pollinator of milkweed (Ivey et al. 2003). Some insects are not robust enough to remove their legs from the anther slits with the attached pollinia and are trapped to die there if they don’t lose their appendage first. Monarchs and other butterflies with their long, delicate legs are probably not as efficient in removing and transporting the pollinia as larger, more robust insects such as our katydid hunter (Sphex pensylvanicus L., seen below) with pollinia on its legs. Milkweed pollinia must be particularly sticky to the touch to ensure attachment to visiting pollinators. Those on the front legs of our specimen have remained attached to it since it was collected on 12 Aug 1952 by H. Jordan in Blacksburg, VA.

Sphex_pensylvanicus_edited
Photo taken by Pat Shorter of the Marek Lab, Blacksburg, VA.

References

Betz, R.R., R.D. Struven, J.E. Wall, and F. B. Heitler. 1994. Insect pollinators of 12 milkweed (Asclepias) species. P. 45-60 in T.B. Bragg and J. Stubbendieck (eds.) Proc. Of the Thirteenth North American Prairie Conference. Univ. of Nebraska, Lincoln, NE.

Borders, B. and E. Lee-Mäder. 2014. Milkweeds: a conservation practitioner’s guide. The Xerces Society for Invertebrate Conservation, Portland, OR.

Ivey, C.T., P. Martinez, and R. Wyatt. 2003. Variation in pollinator effectiveness in swamp milkweed, Asclepias incarnata (Apocynaceae). Am. J. Botany 90:214-225. doi:10.3732/ajb.90.2.214

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New Insect Cabinets!

Our new insect cabinets are here! We now have museum-quality, archival cabinets specially engineered for the protective storage of our pinned specimens. The double-door cabinets are over 7’ tall and hold 48 Cornell drawers. They are made from heavy gauge steel and weigh almost 450 lbs. each. (I was relieved when we decided to hire a moving company to bring the cabinets into the collection room rather than attempt the job ourselves.)

CabinetRow1           OpenCabinet1

The cabinets, along with the new archival Cornell insect drawers and unit trays, were purchased with funding from our grant from the National Science Foundation. Our Collections in Support of Biological Research (CSBR) grant (DBI# 1458045) has allowed us to upgrade the storage infrastructure of our pinned insect collection and secure it for the future.

Many of the old cabinets used in the collection were manufactured to order by inmates in the Virginia Department of Corrections in the 1960s. These older cabinets rarely matched in height, the doors tended to warp and jam, and they were painted a military olive green. Why do I mention the color of these old cabinets? Well, dark interiors make it difficult to visually check the bottom of the cabinets for signs of insect infestation. It’s very hard to see a small dead insect, insect exuvia, or any frass on a dark surface. Our new cabinets have a glossy white finish inside and out, which will make it much easier to inspect the cabinets for things that should not be there.

OpenCabinet2

Our old cabinets were by no means tightly constructed to exclude pests that could damage the collection, but that was acceptable at the time. Fumigants such as naphthalene, paradichlorobenzene (both used in mothballs), and dichlorvos-impregnated strips (often called “pest strips”) were the primary means of protecting insect specimens from attack by dermestid beetles or other stored product pests. A certain measure of air flow into and within the cabinet was beneficial as it allowed the volatile insecticides to drift among the insect drawers. Mothballs were excellent repellents but didn’t always kill established pest populations; pest strips were very effective at eliminating pests. Collections often had these compounds inside each drawer as well as inside the cabinets for maximum protection. [See  Collateral Damage: Unintended consequences of vapor-phase organic pesticides, with emphasis on p-dichlorobenzene and naphthalene, a paper by Kathryn A. Makos and Catharine A. Hawks (both of the National Museum of Natural History, Smithsonian Institution) for more discussion on the use of fumigants in natural history collections. )

So why don’t we still use these materials in insect collections? Most fumigants have been largely phased out of modern insect curation due to safety concerns about carcinogenic and other toxic effects on mammals (such as entomologists). In addition, naphthalene and paradichlorobenzene, when used simultaneously in the same cabinet, combine together to produce a liquid that can damage specimens. Naphthalene and paradichlorobenzene vapors can also recrystallize on specimens, which is also very undesirable.

Nowadays we primarily use exclusion techniques to prevent unwanted insects in the collection. Our new cabinets are tightly constructed with gaskets to be nearly air-tight. The Cornell drawers also very close fitting (to the extent that they can be hard to open, but we’re not going to complain about that).

We can use a sticky card at the bottom of each cabinet to trap any mobile insect inside the cabinet. However, this technique doesn’t monitor insect activity that might be limited inside a particular drawer. Also, many insects tend to move up when trying to escape and may fall on the lid of the top drawer when dead, rather than all the way to the bottom of the cabinet where the sticky card is placed. Frequent visual inspection of both the inside of the cabinet and in each drawer is a more effective method of detecting storage problems in a collection, as long as it is done on a regular schedule by someone who knows what signs to look for. Should we detect any sign of insect infestation in a cabinet, we can freeze insect drawers at -20º C for 3 days to kill immature and adult insects. Each drawer in the suspect cabinet should be frozen and the entire cabinet wiped down and vacuumed thoroughly.

Mold may become a greater problem than dermestid beetles now that insect collections can be stored in air-tight units. Ironically, naphthalene flakes and balls somewhat helped protect insect collections against the growth of mold by absorbing moisture from the air. Mold spores are ubiquitous and freezing will not kill them. To avoid problems with mold in a collection, all specimens must be thoroughly dried before adding them to an insect drawer.

We have a dehumidifier in the collection room that’s typically set for 35% moisture to help limit mold growth. We are also exploring the potential use of portable dehumidifiers that passively absorb moisture from the air. Portable dehumidifiers are often used in gun safes, closets, boat cabins, and other small areas where moisture gets trapped. These dehumidifiers contain beads that passively absorb moisture from the air. When saturated, as indicated by a color change to the beads, the units can be plugged into an outlet and “recharged” by drying out the beads for a number of hours. These would be ideal to place in each insect cabinet for extra protection against humidity and mold growth.

However, the low-tech solution against humidity in the cabinets would be to use humidity indicator cards. These flat cards can be hung on the cabinet door and visually checked each time the cabinet is opened. Increasing humidity levels in a cabinet are indicated by a color change on the card. If humidity is detected, then all the drawers in that cabinet should be visually inspected immediately for mold growth. One of the portable dehumidifiers could then be placed in the cabinet to remove the unwanted moisture, but if it’s humid enough to need a dehumidifier in the cabinet, then clearly all the drawers should be removed from the cabinet until the humidity has been corrected.

Hopefully the precautions we’re taking with tight seals and dehumidifiers will help avoid dermestid beetles and fluctuating humidity levels in our collection. However, these are common curation issues in natural history collections and they do occur from time to time. The real key to keeping a collection safe is vigilant monitoring on a regular basis. Administrators might find the idea of relying on a high-tech device to detect problems appealing, but in reality, several well-trained curators can monitor a collection much more effectively. Plus, we also identify specimens, lead tours, make labels, pin raw material, and remediate any problems once we’ve detected them!

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Things I’ve Found While Transferring Specimens

We’ve been transferring our pinned insect specimens into new unit trays for several weeks now, an activity best described as time-consuming. When finished, we’ll have replaced our current mix of old, hard-bottomed and various types of soft-bottomed unit trays in VTEC with new, archival unit trays.

Pinned insects are so very fragile that it’s extremely easy to knock off a leg or other appendage unless you stay very focused on the task. When the specimens are in good shape and don’t require much additional work, then I might finish transferring all the specimens in five insect drawers by the end of the day. However, if the specimens aren’t already properly oriented on the pin, if the specimens are very small, if a leg drops off and I need to glue it to a card pinned under the label, if there’s a large mix of unidentified material that I need to sort to some crude taxonomic order…well, those are the really long days. So far I’ve come to dread insect drawers containing tipulids, phorids, syrphids, and or tettigoniids as working on them seems to take an eternity. And yesterday I accidentally jabbed a minuten pin holding a Culex sp. mosquito into my finger and was honestly surprised by the amount of blood this produced. (Even in death, mosquitoes are associated with blood loss.)

But one benefit to spending a lot of time looking at the specimens closely is that I see things that I might not have noticed with only a cursory glance in an insect drawer. For instance, I’ve found a  number of specimens that I believe were collected by my thesis advisor when he was a student here at Virginia Tech in the 1960s. There isn’t anything particularly noteworthy about the material he deposited into the collection, but there’s a sense of history when you personally know the researcher who collected a particular insect 50 years ago.

In the category of “Overly honest collection information” we have this poor carabid beetle:

Carabid_Gymshorts

I also found a hackberry emperor butterfly (Asterocampa celtis) with a collection label that claimed the butterfly had “attacked the collector.” Hackberry butterflies are attracted to human sweat and are sometimes very enthusiastic about doing so. Unfortunately the specimen was so badly damaged by dermestid beetles that it was beyond saving.

And then there is this specimen of the katydid hunter Sphex pensylvanicus L.:

Sphex_pensylvanicus_edited

Notice the lighter-brown “sea-weedy” material on the tarsi? There’s a lot of it on the front legs (click on the photo for a larger image). We think these may be the thalli of a parasitic ascomycete fungus, possibly in the Order Laboulbeniales or a similar group. While some Laboulbeniales are associated with ants, I haven’t been able to find any published reports of these fungi on other Hymenoptera. These structures could be from a different type of ectoparasitic fungi or something else entirely, perhaps something new to science. Natural history collections are full of unrecognized species and hidden data waiting for someone to spot them, as described in this recent article from The Atlantic. I have no doubt that VTEC holds its share of hidden discoveries waiting for someone to bring them to light.

Sadly, I don’t have the time right now to investigate this specimen any further as our new insect cabinets have just arrived! Our immediate goal now is to arrange the collection into an updated phylogenetic order in the new cabinets. Sometimes curation is more about housekeeping than scientific research.

(Special thanks to Pat Shorter, an undergraduate member of the Marek Lab, for taking the photo of our infected Sphex wasp! She also took the photos of the Cissites auriculata beetle in our previous post.)

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