Preservation and Curation of Insects
Preservation
Specimens of insects and arthropods, if properly preserved and cared for, can last hundreds of years. Any given specimen carries an enormous potential to inform us about itself and the time and place of collection. Maintaining any specimen for many years carries a cost. Proper preservation ensures a high quality specimen, which increases the quality of information the specimen contains, and increases the value of the maintenance of the specimen. A good specimen takes up just as much room as a bad one. The same as five dirty, rusty, junk cars take up just as much space as five clean, shiny, perfectly restored cars.
Detailed information about general and specific types of preservation can be found within the publications recommended on the Collecting Insects page of this wiki. Below is an overview appropriate for general preservation and curation.
The best long term and short term preservation strategies for insects and arthropods vary among orders. Here is a general list with preferred preservation type.
| Order | Adult: Temporary (Field) Preservation | Adult: Permanent Preservation | Immature Preservation | Comments* | |
|---|---|---|---|---|---|
| 1 | Arachnids (Spiders, Mites, Etc.) | Ethanol (80%) | Ethanol (80%) | Ethanol (80%) | Never allow to dry |
| 2 | Protura | Ethanol (80%) | Ethanol (80%) | Ethanol (80%) | Never allow to dry |
| 3 | Collembola (springtails) | Ethanol (80%) | Ethanol (80%) | Ethanol (80%) | Never allow to dry |
| 4 | Diplura | Ethanol (80%) | Ethanol (80%) | Ethanol (80%) | Never allow to dry |
| 5 | Microcoryphia (jumping bristletails) | Ethanol (80%) | Ethanol (80%) | Ethanol (80%) | Never allow to dry |
| 6 | Thysanura (silverfish) | Ethanol (80%) | Ethanol (80%) | Ethanol (80%) | Never allow to dry |
| 7 | Ephemeroptera (mayflies) | Ethanol (80%) | Ethanol (80%) | Ethanol (80%) | Never allow to dry |
| 8 | Odonata (dragonflies and damselflies) | Dry, kill jar | Pinned or enveloped | Ethanol (80%) | Do not kill or preserve adult in fluid |
| 9 | Orthoptera (grasshoppers and crickets) | Dry, kill jar; ethanol (80%), may fade color | Pinned | Ethanol (80%) | |
| 10 | Grylloblattodea | Dry, kill jar; ethanol (80%) | Ethanol (80%) | Ethanol (80%) | |
| 11 | Mantophasmatodea | Ethanol (80%) | Ethanol (80%) | Ethanol (80%) | |
| 12 | Phasmatodea (walkingsticks) | Dry, kill jar | Pinned | Ethanol (80%) | |
| 13 | Mantodea (preying mantids) | Dry, kill jar | Pinned | Ethanol (80%) | |
| 14 | Blattodea (cockroaches) | Dry, kill jar; ethanol (80%) | Pinned | Ethanol (80%) | |
| 15 | Isoptera (termites) | Ethanol (80%) | Ethanol (80%) | Ethanol (80%) | Never allow to dry |
| 16 | Plecoptera (stoneflies) | Ethanol (80%) | Ethanol (80%) | Ethanol (80%) | Never allow to dry |
| 17 | Dermaptera (earwigs) | Dry, kill jar; ethanol (80%) | Pinned | Ethanol (80%) | |
| 18 | Embiidina (webspinners) | Ethanol (80%) | Ethanol (80%) | Ethanol (80%) | |
| 19 | Zoraptera | Ethanol (80%) | Ethanol (80%) | Ethanol (80%) | Never allow to dry |
| 20 | Hemiptera (true bugs) | Dry, kill jar; ethanol (80%) | Pinned (hard bodied specimens); ethanol (80%) (soft bodied specimens); slides (some aphids and scales) | Ethanol (80%) | |
| 21 | Thysanoptera (thrips) | Ethanol (80%) | Ethanol (80%); slide | Ethanol (80%) | Never allow to dry |
| 22 | Psocoptera (Barklice) | Ethanol (80%) | Ethanol (80%); slide | Ethanol (80%) | |
| 23 | Phthiraptera (biting and chewing lice) | Ethanol (80%) | Ethanol (80%); slide | Ethanol (80%) | Never allow to dry |
| 24 | Hymenoptera (wasps, bees, ants) | Dry, kill jar; ethanol (80%) | Pinned | Ethanol (80%) | Do not kill or preserve adult bees in fluid |
| 25 | Neuropterida (fishflies, snakeflies, lacewings) | Dry, kill jar; ethanol (80%) | Pinned; Ethanol (80%) | Ethanol (80%) | |
| 26 | Strepsiptera (twisted-wing parasites) | Dry, kill jar; ethanol (80%) | Pinned; Ethanol (80%) | Ethanol (80%) | |
| 27 | Coleoptera (beetles) | Dry, kill jar; ethanol (80%) | Pinned | Ethanol (80%) | |
| 28 | Lepidoptera (butterflies, moths) | Dry, kill jar | Pinned | Ethanol (80%) | Never kill or preserve adults in fluid |
| 29 | Trichoptera (caddisflies) | Dry, kill jar; ethanol (80%) | Ethanol (80%), best; pinned (must be spread like moths) | Ethanol (80%) | |
| 30 | Siphonaptera (fleas) | Ethanol (80%) | Ethanol (80%); slide | Ethanol (80%) | |
| 31 | Mecoptera (scorpionflies) | Dry, kill jar; ethanol (80%) | Pinned; ethanol (80%) | Ethanol (80%) | |
| 32 | Diptera (flies) | Dry, kill jar; ethanol (80%) | Pinned | Ethanol (80%) | Drying adults from ethanol may require special techniques |
Table Summary: For work in the field the entire table can be summarized (most simply) in the following way: preserve everything in 80% ethanol, EXCEPT adult dragonflies, bees, butterflies and moths; flies are best killed dry but can be preserved in ethanol until they can be dried properly (usually by a museum). Once back to the lab this table may be referenced to discover which long term preservation is most appropriate for the collected specimens.
* “Never”: the specimen will be destroyed or worthless for identification purposes; “Do not”: it is best not to do this, but if this is the only option it is better to have a damaged specimen than no specimen at all.
Alcohol
Rubbing Alcohol, also called Isopropyl Alcohol and Isopropanol[1] is a commonly available alcohol that can be used to preserve specimens, but it is not recommended for long term storage. Specimens will become very brittle over a short period of time.
Ethanol, also called ethyl alcohol and grain alcohol[2] is generally the best fluid for short and long term preservation of specimens. Low concentrations of alcohol (below 70%) will not properly preserve a specimen, while high concentrations (above 90%) may cause the specimen to crush under osmotic pressure. Generally 80% (160 proof) ethanol is the best to use. In a pinch, high alcohol distilled spirits, such as 100 proof vodka or rum, can be used, but only for short periods of time until replaced by proper strength ethanol.
If there is a chance that the ethanol will be significantly diluted (for example, many specimens in the same jar, specimens are large and fluid filled, etc.) it is best to replace the ethanol once after 24-48 hours. Some large immature flies, dragonflies, beetles, and caterpillars may begin to rot internally before they become sufficiently preserved if placed directly in ethanol. Two common practices used to prevent this are: 1) inject the specimen with ethanol before immersing within ethanol; 2) bring water to a boil, take it off the heat, drop the specimen in the water and leave it for 1-2 minutes, remove specimen, pat dry, place in ethanol. Replace ethanol once after 24-48 hours.
NEVER allow specimens preserved in ethanol to dry out, unless they have been removed for pinning.
Pinning and Pointing
Pinning is a preservation technique where an insect pin is passed through the body of a specimen. As the internal organs of the specimen dry, they cause it to adhere to the shaft of the pin, preventing rotation of the specimen. Thus, the pin functions as both a permanent mount and a handle for moving specimens without touching them.
Pointing is a preservation technique where a specimen too small to pin is gently glued to the tip of a small triangle of paper (a point) which has been stuck onto an insect pin.
When appropriate, always pin a specimen. If properly cared for a pinned specimen will last for hundreds of years, resists discoloration, takes up little space, is easy to view under a microscope, easy to pass from person to person, and cheap and easy to ship.
Pinned insects are “preserved” through dehydration. Insects are covered with an exoskeleton made of chitin, a stable macromolecule. A pinned specimen will dry to the humidity level of the room and, if kept in low humidity, will undergo little to no further decomposition.
Pinned specimens are generally only damaged or destroyed in the following ways:
NEGLECT:
1. Fungal growth resulting from high humidity
2. Damage from insect pests, especially carpet beetles (Dermestidae), etc.
3. Photo-degradation from chronic exposure to sunlight
MISMANAGEMENT:
4. Mechanical breakage from rough handling
CATASTROPHE:
5. Fire
6. Flood
Pinned insects should be kept out of direct light in tightly sealed insect drawers in humidity controlled rooms. Drawers should be checked often for potential pests. If the pest management protocol calls for the use of fumigants, those should be checked often as well. Do not allow inexperienced people to handle specimens. If a collection of pinned insects is not being checked/used more than three times a year it may suffer from neglect. Donation to the nearest appropriate museum or facility with resources to properly curate the collection should be seriously considered.
Pinning and Pointing Insects requires a minimum of equipment, see Pinning Equipment (right): 1) modeling clay to put pins in; 2) pinning block, a jig used to properly space the specimen and labels; 3) points, triangular pieces of label paper upon which small specimens are glued (shown here in a stender dish); 4) fine tipped forceps to pick up, move, and arrange specimens; 5) insect pins, these come in different sizes, 00 is smallest, 7 is biggest, generally sizes 2 or 3 are best (straight pins are not appropriate for use); 6) glue used to affix a specimen to a point, clean fingernail polish may also be used.
Pinning specimens the same way minimizes handling time, allows for easier storage, and reduces damage to the specimen. By convention different groups of insects are pinned in different places. This ensures consistency among collectors over space and time. Generally insects are pinned through the “middle” (front to back, see below) just to the right of the midline (side to side). This ensures that any important characters that fall on the midline are not destroyed. However, some insects, such as butterflies and moths, are pinned through the midline to ensure that the wings rest symmetrically. See Pin Placement, right.
Grasshoppers, flies, bees and wasps, dobsonflies, and most other insects: pin through the thorax just right of the midline.
True Bugs: pin through the triangular scutellum just right of the midline.
Beetles: pin through the fore part of the right wing cover (right elytron) near its center line.
Butterflies and Moths: pin through the center of the thorax (some people pin wasps and flies through the center of the thorax, and this is acceptable but not recommended).
To pin an insect:
1. Steady the specimen with your fingers or place it on a stable surface, be careful not to crush it.
2. Place the tip of the pin at the location where it should enter the body.
3. Slide the pin all way through the specimen; be sure the pin is vertical in relation to the specimen.
4. Use the pinning block to position the specimen to the correct height on the pin.
5. Fold the legs under the body and position the wings and antennae appropriately. The goal is to have a specimen that takes up as little room as possible, without appendages sticking out awkwardly that could be easily broken off. Obviously, insects that require spreading will not fit this generalization.
Insects that are too small to pin should be pointed, see Pointing Position, right. Points are about ~5 mm long and should be made of Bristol board or heavy archival paper. Standard point punches can be purchased, or points can be cut by hand.
The way a specimen is positioned on the point is very important. Be sure you know and understand these instructions before you begin pointing specimens.
By convention the tip of the point is glued to the center of the RIGHT side of the specimen so the specimen is facing up and the pin is on the right when viewed from above (Pointing Position 1). To increase surface area for glue the tip can be bent downward with fine forceps (2). The point can go under the specimen but should not extend beyond the midline of the specimen (3). An unobstructed view of the top (1), bottom left side (3), and left side (4) of the specimen is now available. Many publications follow this convention and provide figures and photographs with the specimens oriented in this way, making comparisons between figures and specimens much easier. Proper pointing requires a bit of practice, and everyone has their particular way of doing it. But the end result should be consistent with archival standards.
Labeling
Lessons about labeling have already been presented twice in this wiki: on the Photography page and on the Collection page. This is because labeling is very important. Review previous labeling lessons if you are unsure of what information is important for a label, and the proper format for that information.
Here we will concentrate on how to make permanent labels. Field labels are ephemeral, they only need to convey information for a short period of time, from the field to the lab, and can be written on almost any convenient scrap of paper. Electronic labels attached to a photograph do not alter the way the photograph is stored, sorted, etc.
Permanent labels for pinned, pointed, fluid preserved, or slide mounted specimens differ from field or electronic labels:
1. They must be made of a substance that can last as long as the specimen, potentially hundreds of years.
2. They take up physical space, every square inch of an insect drawer costs money to buy and maintain.
3. They have a real potential to damage other specimens when specimens are being moved in a drawer.
4. They are read by humans.
5. They are generally found grouped together (a unit tray may have 50 specimens with 50 different labels made by 50 different people).
Considering the above, a permanent label should have the following qualities: 1) made of archival material; 2) as small as possible so as to not take up too much room in the collection (this is not as important for specimens stored in fluid); 3) not overly long or oddly shaped to be less likely to break other specimens; 4) font size large enough to be read by a person, with clearly defined characters (sans-serif), and with unambiguous information; and 5) information presented in a specific order so humans scanning many labels will be able to quickly find the specific specimens (for example, if among 200 specimens of species X only those from Arkansas are needed, it is easier to scan the labels if "state" is always in the top left corner of the label).
Permanent Labels for Pinned and Slide Mounted Specimens
Making good permanent labels for pinned specimens is difficult because 2) and 4) are at odds with one another; make the labels small BUT make them big enough to read. Additionally 3) and 5) come into conflict; keep the labels similar sized BUT put the same information in the same location on every label. This is difficult if one specimen was collected in Clay Co. (8 characters) and another in Montgomery Co. (14 characters).
Here is an example of a good label and label sheet:
How to make a label sheet in Microsoft Word 2007:
1. Write the label. It should be four or five lines long.
2. Change the font to Calibri.
3. Change the font size to 4 (a "4" must be typed into the font size box).
4. Zoom in until the label can be read.
5. Altering Page Margins will keep labels from being broken across columns. Different page margins are needed for labels with different numbers of lines. Thus, assuming all labels have the same number of lines (4 or 5), choose one of the following:
5 A. Label with 5 lines: change the top margin to 0.19 and all other page margins to .25 inches (under Page Setup).
5 B. Label with 4 lines: Change the bottom margin to 0.27 and all other page margins to .25 inches (under Page Setup).
6. Got to Page Layout, select Columns: More Columns. In Number of Columns type "10". Hit OK. Go back through Page Layout: Columns: More Columns and for Column #1 change Spacing to "0.01". Make sure Equal Column Width is checked.
7. Select All (Control+A), this will highlight everything.
8. Go to Page Layout and open the Paragraph Dialog Box by clicking on the button to the right of "Paragraph".
9. Under Indents and Spacing: Spacing: Line Spacing, select "Exactly" and enter "4" under "At". This reduces the line spacing so the lines are close to one another which reduces the wasted space on the label.
10. Copy and paste the label as many times as needed. To speed this up paste 10 labels singly, then copy all ten and paste that multiple times. When using one label as a template for a label with different information, be sure to change ALL the appropriate information!
The protocol above will produce 470 four-line labels or 380 five-line labels on a single sheet of label paper. DO NOT put spaces between the labels. This only wastes paper and increases the amount of time it takes to cut out the labels.
Download a Label Template file here (Microsoft Word, .doc).
Label Positions: Generally a pinned specimen will get three or four labels. Labels should be positioned as follows:
Top. Capture labels: Locality labels; labels specific to the particular project/research the specimen is associated with; etc.
Middle. Specimen Identifier labels: Database number/barcode; Museum Identifier labels; Voucher Specimen labels; etc.
Bottom. Determination labels: Identification labels and Type labels. Generally these are not removed as new or more accurate identifications are made, but can be turned upside down to indicate they are no longer valid. Generally the bottom most label is the more recent determination. Determination labels should include the name and year the identification was made (e.g. Diabrotica virgifera LeConte, Det. J. Smith 2011).
When positioning labels, make sure the top label is far enough from the specimen to be readable and is not close enough to the specimen to cause breakage. Labels should not be so close to one another that they cannot be read. A pinning block (see Pinning Equipment, #2, right) is an important tool in standardizing label height.
See Pointed and Labeled Specimens (right) for an example of specimen standardization and label placement. Note the standardized specimen position on the point, point height on the pin, label height on the pin, pin placement through the label, and distance between Locality label and Database label (right insert). Additionally note that the label takes up much more space than the actual specimen. In an extreme example, approximately 1000 specimens can be stored in one drawer (middle insert). If properly labeled and curated, many specimens need not take up much drawer space, the 16 drawers pictured in the background hold about 8000 specimens.
Permanent Labels for Fluid Preserved Specimens
Generally labels for fluid preserved specimens (see Fluid Labels, right) must contain all of the important qualities outlined above, except that the labels can be made much larger, generally font size 6. Labels should NEVER be loose in a vial or placed vertically, this will damage the specimens. Instead labels should be long enough to wrap around the inside of the vial. The locality label should face out and be located in the top 25% of the vial where it is easily readable. The identification labels should be separate from the locality label (in case the identification is incorrect or incomplete) and should be wrapped behind the locality label facing in. The identification label is read by looking through the vial.
Curation
Curation of a collection is an enormous responsibility. As has been mentioned several times, preserved insect and arthropod specimens can last for hundreds of years. The oldest known insect collection is that of English botanist Leonard Plukenet (1641–1706)[3] who pressed his insects between the pages of a book like botanical specimens. The oldest pinned insect is in the Oxford University Museum of Natural History, a Bath White butterfly, dated 1702. Curating an insect collection is a long term endeavor, to put it mildly.
Collection, preparation, and identification of specimens can be very costly in time and supplies. Housing a collection requires cabinets, drawers, etc. which can be expensive to purchase and may take up valuable floor space. These observations are not made to discourage the collection or preservation of specimens, nor are they meant to discourage the keeping of a collection. They are meant to emphasize the responsibility that falls on the shoulders of the curator. Even a small collection of a few thousand specimens may represent tens of thousands of dollars spent on travel, labor, equipment, and cabinets and thousands of man hours spent collecting, preparing, and identifying specimens. All the specimens, their value, and their potential can be lost in a few years of neglect. Indeed, there are horror stories of large, important private collections that were destroyed by ignorant family members after the collector's death ("We dumped all the bugs out because we wanted to save the bottles.").
Proper curation of an insect and arthropod collection consists of three things:
1. Preservation of specimens.
Short Term: Specimens in drawers must always be kept in a low humidity environment and must be checked at least every 3 months for pest infestations. Fluid levels of fluid preserved specimens must be checked every 12 months. Detailed information about general and specific types of preservation and curation can be found within the publications recommended on the Collecting Insects page of this wiki.
Generally the only way to assure that specimens are properly preserved is to hire a part time or full time curator.
Long Term: A known, written, funded plan must be in effect to properly deal with a collection in the event of the death of the curator, loss of funding for the collection, etc. Generally an agreement is made with a large facility that will accept the collection as a donation. Additionally an account should be established to help defer costs of transportation, curation, and supplies incurred by the accepting facility.
2. Maintaining access to specimens.
Specimens should be available for others to study, which includes regular access to the collection, lab space, a microscope, a system and records keeping for loans, and the appropriate ordering of specimens so specific taxa may be found quickly. Specimens locked away from the greater scientific community are of little value.
Generally the only way to assure that specimens can be accessed is to hire a part time or full time curator.
3. Upkeep and growth of the collection.
As our understanding of the natural world changes, the taxonomic status of a given species may change. For example a species may be moved from one genus to another, or what was once thought to be a single species may be two or more subspecies, or a complex of multiple distinct species. To reflect these changes in the collection, current literature should be regularly surveyed, identification cards should be updated, and the collection should be rearranged as necessary.
Additionally, collections should have facilities to accept new specimens. Room must be available for additional specimens of taxa already represented in the collection, and for new taxa added to the collection. A general plan for collection expansion should be devised.
Generally the only way to assure upkeep and growth of the collection is to hire a part time or full time curator.
If any of the three major aspects of curation cannot be adequately met consider working with an appropriate museum/facility as a collection is being assembled, or, if a collection already exists, consider donation of that collection to an appropriate museum/facility.
Print References
- ↑ Packard, A. S. 1890. Entomology for beginners: for the use of young folks, fruit-growers, farmers, and gardeners. 3rd Edition. Henry Holt and Company, New York. 367 pp.
First Detector Entomology Training Project
