Volume I, no. 1, January 1992Table of Contents
Winter pitfall trapping? -
George Keeney
Winter Pitfall Trapping?For most collectors, the end of summer signifies the time to hang up the net and the time to start curating the year's catch. With the first wintry blast of December, one longs for an early spring. Some are content to wait. Others develop an ever growing "collector's itch", anxiously awaiting the annual rites of spring. Aside from collecting infested wood and peeling bark, I scratch my itch by pitfall trapping in what some consider to be the most unproductive time of year. Granted, one doesn't get a large volume of insects that can be typical in spring and summer trapping, but it can add diversity to your collection. You may get species that are out only in the late fall, winter and early spring. Beetles which I have commonly taken pitfalling at this time of year include various carabids i.e. Carabus, Sphaeroderus, Scaphinotus, Pterostichus and Amara; staphylinids and the dung scarab, Geotrupes egerei. One species, Nicrophilus pettitii, which is normally only rarely collected and often difficult to find, actually is taken fairly frequently in winter trapping. It belongs to the family Agyrtidae, long classified as a Silphidae but now considered to be allied with the Leiodidae. Peck (1981) gives the distribution and biology of this flightless carrion feeder. The above mentioned beetles are but a few of the different kinds you can get. My experience has been limited to southern Ohio, so I would suggest trying different geographical localities and habitats. I'd be interested in knowing the results if anyone gives it a try. [results can be sent to OC-editor] Setting up a winter pitfall trap is basically the same as you would do any other time of the year, but with a few additional considerations. The pitfall container itself should be a sturdy, unbreakable plastic container as frost-heaving during freeze/thaw cycles could crack containers that are too rigid or brittle. Certain plastics may become brittle when cold. It you don't know about your particular container, place one in a deep-freeze for a couple of hours, then try bending it while cold. Also fill the container with water and freeze it to see if it will split as the ice expands, in case it inadvertently becomes filled with water and subsequently freezes. I have found that 1/2 gallon or 1 quart deli containers, like the kind that potato salad comes in, to fit the bill quite nicely. Ideally, the container Should be 15-20 cm deep and 15-20 cm in diameter. If an inner bait container is to be used, perhaps you should choose towards the higher end of the range. I cover my traps with a 30X30 cm, square of scrap plywood or masonite as a rain shield. (see figure on next page) When the trap is completely assembled with screening the shield also keeps raccoons from reaching directly into the trap. The shield is suspended 5-8 cm over the container by supporting it on short dowel rod pieces, sticks or small stones. After placing the rain shield, I cover the entire trap with 60X60 cm square of chicken coop wire. This mesh size provides no barrier to insects. This is secured, tautly, at the corners with 6-7 Inch gutter nails. When in place and covered with some leaves and twigs, it not only provides a pitfall that is fairly secure from raccoons, skunks and other similar scavengers, but also disguises it from the casual human observer. I seem to have more trouble with hunters, mushroomers and hikers than raccoons. Often times I may have a few minutes of trouble finding it myself! Try to remember local landmarks. With little underbrush foliage, uncamouflaged traps can be easily spotted. In my opinion trap security is a must and becomes increasingly important in areas of high human and/or scavenger activity. This not only protects your investment in time, money, and prevents lost specimens, but is also the ecologically sound thing to do. Secure traps prevent the poisoning of pets and wildlife and contamination of the water supplies, particularly if ethylene glycol or "antifreeze" is used as a preservative in the trap. I prefer to use ethylene glycol because it is easy to obtain in large amounts, and it's relatively inexpensive. Its chemical stability allows for reuse in the trap provided it has not become too diluted or soiled. Simply sieve the insects from the fluid and replace the fluid in the trap. The fact that it is "antifreeze" is useful during winter. Just remember to soak the specimens in alcohol for a couple of weeks to remove the residual antifreeze. If you do not remove it properly, the specimen will simply harden up and may appear greasy. Also keep animals away from the antifreeze, as it is sweet tasting and very toxic to animals. Ethyl and isopropyl alcohol (70%) have much lower freezing points than water (-117 & -89 degrees C, respectively, when absolute). Trouble is that alcohol evaporates rapidly, necessitating that traps be checked rather frequently. The addition of glycerin would prevent specimens from hardening in the event the alcohol evaporates entirely. Winter temperatures may sharply reduce evaporation rates. Formalin degrades rapidly, polymerizes at temperatures below 13 degrees C and is just a nasty caustic chemical. I consider it unsuitable for winter use. It may be suitable for summer use but I don't know if it exhibits any attractiveness or repellency to beetles. The addition of sodium chloride (salt) to water lowers the solution's freezing point. This coupled with its preservative properties may give concentrated brine solutions some distinct advantages. Salt is readily available at low cost and certainly much less toxic than ethylene glycol. One drawback is that salt is corrosive to insect pins. Be sure to gently rinse the specimens in fresh water and soak in alcohol for a couple of weeks changing the alcohol a few times to remove the salt residues before pinning. A couple of changes may be necessary as sodium chloride is not extremely soluble in 70% alcohol. Failure to do so may result in salt blooms on the specimens and rusted, broken pins. All of the above mentioned chemicals have been used in various combinations. It is apparent that there could be many different combinations, many equally satisfactory. A lot of information on trap preservatives seems to be anecdotal and in fact there may be some of you that have your own "recipes". This topic will be discussed more comprehensively at a later date. The use of bait in the traps depends upon your purpose in trapping. Ecological studies may preclude their use. They could lead one to false assumptions about population densities, particularly it the insect is highly motile and attracted from over a large area. If you prefer to use a bait and "get more bug for the buck", there are a number of things to choose. Typical baits used may be dead mice or rats, liver or dung. Use your imagination, for different materials will attract different kinds and numbers of insects. I have found fermented beans to be very attractive to both carrion and dung feeders. The bean bait remains attractive for over a month, unlike carrion and dung. Because of the potentially lower temperature during the trapping period, whatever bait is to be used should be ripened at room temperature 47 days in a closed container. When fermenting beans, cover them with an amount of water equal to twice the volume of the dry beans as they will absorb a large amount of the water. I always prefer soil to inoculate the material with microorganisms, just to get things rolling. Place the bait in an unbreakable container that is appropriately sized to fit within the outer container. A one pint container is proportioned nicely to a 20 cm deep X 20 cm diameter outer container. Cover the jar securely with nylon mesh or window screen to prevent the beetles from entering the bait and making a messy catch. Do not contaminate the bait with the preservative. The amount of preservative should come no higher than one-half the height of the bait jar. Be sure to put enough fluid in to kill and cover the catch though. If you want to learn more about pitfall trapping and trap designs I have given some references. Morril (1975) Shows how to make a very simple and inexpensive trap from various plastic cups. An early, complex model is described by Fitcher (1941). Wojcik et. al. (1972) shows a less complicated model but one which is labor intensive and needs specialized parts. Alter looking at their designs, you may get some ideas and find the parts around the home to design your own style of trap. In conclusion, remember that trap counts will be influenced by temperature, even though these beetles may be active in winter. Extremes of cold will still suppress the activity of cold-tolerant beetles so don't be disappointed in the low numbers during cold snaps. The bait will basically remain "refrigerated" and as soon as warmer winter temperatures return, so will the beetles. -George Keeney 100 Year Old Collection Back in ActionThe Ashland county Historical Museum (ACHM) houses an insect collection which contains approximately 12,000 specimens of Coleoptera. This collection was assembled circa 1900 (earliest is 1843 most recent 1850) by Thomas Thornburg, a horticulturist and an amateur beetle collector. The collecting done locally by T. Thornburg was greatly augmented by his trading with collectors and purchases of exotics from dealers. A large number of the specimens in the collection, and those of greatest age, are from a purchase of an already old European collection. Historically significant names and places arise in the collection. J. N. Knull, known for his work on Cerambycidae and F. H. Snow, founder of the U of Kansas Entomology Department are both contributors to the collection. There is evidence that some of the specimens were collected by A.R. Wallace of evolution theory fame and many have dates and locations which coincide with Livingston and Stanley expeditions. According to the grandson of T. Thornburg his grandfather did in fact receive packages from these expeditions. The collection houses examples of many of the world's most dramatic beetles. It covers all regions but is far from synoptic in regard to species. Like much of the collecting done during the period represented in this collection data is weak and often absent. When I first discovered this collection it was in a state of disrepair. The beetles have since been restored. This involved re-pinning of all specimens, re-lining of cases, identifying or correction of identification and the creation of a computerized data bank of all information on the specimens. As of now the re-pinning and re-lining are complete. Identification and data list have been done as noted below. I would invite anyone interested in seeing the collection to contact me or stop by the museum during its open hours (April 1 thru December 28 Wednesday, Friday, and Sunday, 1 pm till 4 pm). Anyone interested in working on any of the beetles should contact me. Specimens are available for loan, and in some cases, trade. [chart goes here] Most other families are represented in the remaining drawers by one to 20+ specimens. Approximately 10,000 specimens of Lepidoptera and an assortment of other orders are also in the holdings but are presently under restoration. * USNM Standard drawer
- Kip Will An Unusual Method for Collecting BuprestidsDuring the summer of 1976 I had the opportunity to work at a YMCA camp, Camp Glen Taylor, near Mifflin in Richland County, Ohio. My duties as the nature lodge counselor allowed for a good deal of time for collecting in the nearby woods and fields. One afternoon, as I was collecting cicindelids in an old, seldom-used baseball diamond, I noticed a medium-sized wasp flying toward the bare infield from the direction of a nearby wooded area. From its direct path of flight in a slow, labored manner it was obvious that the prey it was carrying was quite a load for the wasp. Upon landing, the wasp disappeared quickly down its burrow with its prey before it could be inspected more closely. As I began examining the area I counted more than one hundred similar burrow openings in an area of approximately 1000 square feet. Occasionally a wasp would be seen leaving its burrow or sitting in the entrance of the burrow. Within several minutes, another wasp, laden with prey, flew into the area from the direction of the woods. I swept the wasp from the air, jostled it in the net to force it to release its prey, and then released the wasp. In the net was a specimen of the buprestid species, Chrysobothris femorata (Olivier). Within a couple of hours, I netted and released nearly thirty wasps, stealing their cargo from them. All of the wasps captured were carrying buprestid beetles, although three different species of buprestids were represented. Chrysobothris femorata (Olivier) was most commonly taken by the wasps, this species constituting 21 of 28 specimens taken. Six specimens of Dicerca divaricata (Say) were collected, as well as a single specimen of the rarely collected Buprestis rufipes (Olivier). Several interesting observations were noted. Upon capture and release, all of the wasps flew directly back toward the wooded area, none flying toward the nesting area. Whether this was a predator avoidance behavior, by flying away from the burrows toward more heavy cover or whether it was a return to the search behavior is unclear, due to the limited time spent observing the wasps' behavior in depth. Although all of the prey beetles were buprestids, the three species differ markedly in both morphology and host plant associations. Chrysobothris femorata (Olivier) is relatively small (8-15mm, a brassy charcoal in color, and frequently uses oaks and hickories as its larval host. Dicerca divaricata (Say) is larger (15-20mm), coppery in color, and uses a variety of hardwood hosts. Buprestis rufipes (Olivier) is large (20-25mm), bright green with yellow stripes, and utilizes maple and beech as its hosts. The wasp may locate the beetles by detecting them by their quick, jerky motions as they move along the branches or trunks of the host trees. This type of movement is the only trait common to the three species of beetles in a structural or behavioral sense. One other possibility is that the wasps were using chemical stimuli to detect the beetles. The most likely would be sexual pheromones given off by the females, but even these would be expected to be species-specific for the beetles. The pheromones of the Buprestidae may be generally similar enough across the family to allow the wasps to detect a basic buprestid "scent ". Had the beetles been sexed it could have given some hint as to the answer of this question, but enough of the specimens have been exchanged over the years as to make an assessment of the proportion of males to females impossible. Unfortunately, the wasps were not well identified and have since been lost to some event of history. Only a general identification was made, the wasps belonging to the family Sphecidae, most likely being a member of the subfamily Philanthinae or Crabroninae. In the summer of 1985, a similar but less extensive area of burrows was observed at Lake Hope State Park in Vinton County, Ohio. These were situated on a barren slope along a roadway through a wooded area. Although the wasps seen were superficially similar to the species observed in 1976, they were provisioning their burrows exclusively with beetles of the family Curculionidae. This was probably indicative that the wasps seen in Vinton County were of a different species than that of the wasps seen in Richland County. Although this type of situation is not encountered often, it can provide a productive opportunity for the collector to take species that are not easily collected in numbers by other methods. Any collector who has attempted to hand-pick buprestids from their host plants on a hot summer day knows that it can be an exercise in futility. In comparison, a wasp burdened with a paralyzed beetle of a size nearly equal to its own is unlikely to evade even the most inept net-swinger. Although it would be easy to work an area of burrows repeatedly, it only makes good ecological sense to sample an area, and then leave the wasps alone to do their thing. Overly molesting an area could have detrimental effects on the wasp population, and should be taken into consideration by the collector. - R. A. Androw Spider Beetles in OhioPtinidae, spider beetles, comprise about 55 genera and 600 species worldwide. There are about 55 species known in North America including 10 introduced species. The majority of species are found in the drier parts of the subtropical and temperate zones, although this may be a collecting artifact, due to the relatively unknown fauna of the tropics. Most species are scavengers and are found in a variety of habitats such as mammal and bird nest, hymenopteran nest, leaf litter, stored products and other places with relatively dry dung or organic debris, such as caves or tree holes. There are also a number of woodboring species and still others associate with ants. I will tell you more about their unusual habits in a forthcoming issue of this exiting new newsletter. A number of species closely associated with stored products have become spread by commerce throughout favorable areas. Unfortunately, these make up the majority of our fauna in Ohio, we do have three or perhaps four native species. Most of these introduced and native species belong to the genus Ptinus. The native species range in size from 2 to 4 mm while introduced species can reach lengths of 5 mm. I am trying to collect native species to rear and produce larvae and pupae. They have proved so far to be an elusive family of beetles to collect and I would be forever thankful to my fellow beetle enthusiast to be on the lookout for these beasts (even introduced pest species). Keep your eyes open on your blacklight sheet, for this may be the best way to collect some of the native species. Other habitats previously described are also productive places to look for them. [Information about the ptinids you collect can be sent to OC or the author-editor] - Keith Philips Immature Beetles: the Rest of the StoryHistorically coleopterists have concentrated their efforts on adults and have all but ignored the immature stages. Consequently little is known about the immature stages of most beetles. Presently it is almost impossible to identify a field caught beetle larva to species. This need not be so. Increasingly we find that there is much to be gained by studying characteristics of more than one life stage. Often the habits of adults and larvae stand in stark contrast to each other. The adult beetle Is essentially geared towards reproduction while the larva is concerned principally with feeding. More often than not, the morphology of the larva and the adult share little in common. Thus two radically different manifestations of a single genotype team up for survival. Many students of Coleoptera find themselves preoccupied with building a collection. The impetus to do so generally stems a fascination with the adult form. I freely admit that it was the incredible diversity of adult beetles that resulted in my decision to study them. Eventually, I became fascinated with immature beetles and insect rearing. Insect larvae and pupae can be every bit as interesting as adults. Most amateur coleopterists who rear insects do so in hopes of procuring an adult specimen which can be pinned. While the idea of rearing beetles to obtain the immature stages may sound foreign, I can think of no better way that an amateur may potentially benefit the systematic community. There are two methods for obtaining immature beetles: 1) procuring them in the field and 2) culturing adults. Larval beetles occur in a plethora of microhabitats and may or may not occur alongside adults. They are sometimes easier to collect than adults especially when the adults are short lived. Since there are almost no keys to species of larval beetles, it may be worthwhile to keep larvae alive and rear to an adult stage. If one is successful in doing this, the larval exuvium should be preserved. It should be either placed in a vial or mounted on a card and pinned beneath the adult specimen. Culturing beetles, while challenging, can be profitable. In addition to obtaining with certainty the immature stages of a given species, Insect cultures also yield adults which can augment specimens in your own collection and provide additional materials for exchange. Immature beetles are best preserved in 70% ethyl alcohol. Whenever possible, all immature beetles should be placed in boiling water for about a minute before they are placed in alcohol. This will denature enzymes present which would normally accelerate the decay process and allow the alcohol to penetrate and preserve muscles and internal structures. While it is true that storage of immature insects does lend itself to frequent inspection, as does pinning, this should not deter one from making a larval collection. Provided that care is taken to prevent evaporation of the preserving fluid, immature beetles should remain in good shape indefinitely. Also there is never a danger of them becoming dermestid fodder. - Peter Kovarik Bembidion obtusum range extensionIn the September - October 1991 issue of The Entomological Review, E. R. Hoebeke, J. K. Liebherr and R.T. Bell report on the expansion of the known range of Bembidion obtusum in the eastern United States and specifically into Ohio. Specimens from Ohio were collected by H.J. Lee of Cleveland. This species, native to central Europe, is reported to have been caught on "open clayish ground". It has been taken both by pitfall and leaf litter as well. As the afore mentioned authors noted the B. obtusum shows dimorphic hind wings and are therefore very interesting to look at for their dispersal pattern. Keep a lookout for this species and let OC know if you get them in the coming year. - K. W. Will & R. A. Androw Tiger Beetle Colors[Reprinted from "Tiger Hunt" by Tom Schultz in Natural History, May 1991] The colors of the tiger beetle are produced by the surface of the insect exoskeleton, which under extreme magnification resembles a waffle. The hundreds of thousands of tiny pits are lined with transparent reflecting layers, less than three ten-thousandths of an inch, which can only be seen with the aid of an electron microscope. The thickness of the layers in any given pit determines what wavelength of light or color, the pit will reflect. Each layer reflects all the wavelengths of the incoming sunlight, but only the light waves from a very narrow portion of the spectrum produce colors we see. As the waves bounce off the multiple reflecting layers, only one wavelength will emerge from the pits in phase. The waves that are in phase will reinforce one another; the rest emerge out of phase, canceling one another out. (This is analogous to what happens when two sets of waves traveling across the surface of the ocean meet: if the troughs and crest of one set coincide with those of the other, they enhance each other; but if the crests of one wave train coincide with the trough of the other, the net effect is no waves at all---they cancel each other out.) Which wavelength emerges from a pit is determined by the thickness of its layers. The dimensions of the reflecting layers are determined by the cells that secrete new exoskeleton when the beetle molts from a pupa to an adult. Below the reflectors, these cells also deposit the black pigment melanin, which, like the backing of a mirror, intensifies the color by absorbing any nonreflected light. The white markings on the tiger beetle forewings are areas where the reflectors and melanin are lacking. - Tom Schultz |