Responses to Questions Posed on the Entomo-l e-mail List
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Odonata Information Network: http://www.afn.org/~iori/
All material © 1994-1997 Terry Morse and The Respondents
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935 SW 10th St. #6
Newport, OR 97365 USA
oBrian Armitage, Ohio Biological Survey, Columbus, Ohio, USA (BA)
oPaul Bardunias, University of Kansas, Lawrence, Kansas, USA (PB)
oStephen Brooks, Natural History Museum, London, UK (SB)
oRob Cannings, Royal British Museum, Victoria, British Columbia, Canada (RC)
oEverett D. (Tim) Cashatt, Illinois State Museum, Illinois, USA (TC)
oSoowon Cho, University of Maryland, Maryland, USA (SC)
oEric Coombs, Oregon Department of Agriculture, Salem, Oregon, USA (EC)
oSid Dunkle, Collin County Community College, Plano, Texas, USA (SD)
oPaul Florence, Louisville, Kentucky, USA (PF)
oBill Mauffray, International Odonata Research Institute, Gainesville, FL, USA (BM)
oRichard L. Orr, Columbia, Maryland, USA (RO)
oDennis Paulson, Slater Museum of Natural History, University of Puget Sound, Tacoma, Washington, USA (DPa)
oDave Pehling, Washington State University, Everett, Washington, USA (DPe)
oHugh Robertson, University of Illinois, Urbana, Illinois, USA (HR)
oKarla Segelquist, Snow Entomological Museum, University of Kansas, Lawrence, Kansas, USA (KS)
Respondents' comments are identified by the initials in parentheses following their names and affiliations above. Comments I have inserted in other peoples' responses are in square brackets and initialed TM.
A brief discussion of ways to prepare aesthetically pleasing and scientifically useful specimens, based on my recent experiences
More information from Tim Cashatt on acetone preservation and DNA analysis.
The questions: In netting dragonflies, are dark- or light-colored net bags best, or does it matter?
Is an 18" (46cm) diameter net preferable to a 15" (38cm) net, and a long handle to a short one? How do you assess the tradeoffs between speed and ease of swing versus reach and area of the net opening?
BA hypothesizes that a dragonfly might remember a white net longer than a green one when you swing and miss, hence being made more wary on subsequent attempts (SC concurs with this). He admits, however, that he uses a white net and hasn't tried green.
SB prefers a dark bag (black or green; he uses black), because it will be less visible to the dragonfly. The drawback of the dark bag is that it is harder to find your specimen in it (SD comments that this is an important consideration when working in low light conditions, i.e. tropical rain forests). The bag should be deep to ensure that the dragonfly isn't able to fly out when you flip the rim over to trap the dragonfly in the bag. SB prefers a 15" net with a 4' (1.2m) handle, considering speed of swing more critical than the wide opening, and recommends the sport of badminton as practice for netting dragonflies.
SC prefers a green net with large mesh-size to reduce air resistance as you swing. His rule of thumb is a mesh just small enough that the smallest dragonfly you might catch can't quite get its head through the mesh. He recommends a bamboo handle, because its light weight makes it much easier to swing fast.
RC recommends an 18" net and an aluminum handle of collapsible 12" (30cm) segments (BioQuip #7312AA) . While many people prefer dark bags, he says he doesn't worry about it. [Bioquip's 3-section Tropic Net, #7324A-C which comes in 2' (60cm) sections, and with an optional carrying bag, #7324D, is also quite convenient and has worked well for me as I bicycle between collecting sites - TM]
SD states that, while green nets are probably better for the wariest dragonflies, he prefers a white net because of the ease of finding small specimens in them under low-light conditions. He surmises that a white net in the open might seem to be a cloud to a dragonfly, offsetting the presumed lesser conspicuousness of a dark bag. He laments that no one has studied the question scientifically. [Shouldn't be too hard to do in an entomology lab - give half the class light nets and half dark at random and see which group catches more (don't tell them why some have light and some dark nets until after the fact); let the students do the analysis as an exercise - TM].
As to net size, SD believes that the biggest bag you can swing rapidly is best (i.e. 18" is better than 15"). He prefers a bag with 1/8" mesh to allow fast air flow while still retaining damselflies, and tough enough to withstand briars. He hasn't found any material with these properties in the USA. He says that there are some excellent nets made in Japan, but they are quite expensive. In Dragonflies of the Florida Peninsula ... he says he uses a modified fish-landing net with a 50 cm diameter hoop and a 2 m handle.
PF, who has been collecting dragonflies for the past 2 summers, prefers a 15-18" net, and finds the smaller one easier to swing. He has never tried a green bag. Though finding that dragonflies sometimes seem wary of a white bag, he observes that the same may also be true of dark bags.
RO recommends a light-weight green aerial bag, 18" diameter, on an 8' (2.4m) handle for active dragonflies. He feels that the increased coverage of an 18" net is more important than the very slightly greater weight of the larger bag. He suggests that the green net is less visible to the dragonfly, but agrees that it is harder to find small specimens in a dark bag.
Ken Tennessen (1994), while collecting Gomphids in North Carolina, noticed that if he held his net out low with the white bag held tight against the handle, patrolling males would attempt to land on the bag and could be easily caught with an upswing of the net. The males did not appear to be attracted to an all-black bag and minimally so to a green bag with white rim. [I've had Sympetrum land on the white rim of my green net, and quite often on my light clothing as well. Recently, one landed on the white`knuckle of my index finger while I was jotting field notes about it and used the knuckle as a hunting perch for 30 minutes before taking off after a passing Sympetrum and not returning. - TM]
1) Use a long-handled aerial net at least 18" diameter. Dark net may be preferable to a light one.
2) Watch awhile before swinging at a patrolling dragonfly. This will allow you to position yourself most advantageously. Choose a concealed location if possible. Move deliberately, keeping the net as inconspicuous as possible until the moment of truth.
3) Swing at fast-flying, agile species from behind as they fly by; many will easily dodge a net swung head-on.
4) Place the live specimen in a glassine envelope (3.5" x 2.25" is best), the wings together above the back. Place pairs caught in tandem in the same envelope if possible. If too large to go together in a single envelope, indicate that they were in tandem on both envelopes. Write the collection number cross-referenced to field notes or full collecting information on the envelope in ink which will not dissolve and run in acetone, if they are to be so treated.
1) In the field, keep the envelopes containing live dragonflies in as cool a place as possible. Store in a non-crushable box, such as plastic Tupperware or similar fare.
2) Since the color of some species fades soon after capture (e.g. some Aeshna), soak them in acetone as soon as is practical.
From Dragonflies of the Florida Peninsula ..., pp. 13-14:
1) The most important aspect of collecting is recording detailed information on the day of collection. Include the state, county, nearest town, nearest water body, nearest highway, date, time of day, weather, name of collector, habitat notes, and behavior notes. [See Herman (1986) for a useful method of scientific note-taking - TM] Of particular scientific interest are pairs associated in tandem or in wheel (be sure to record this fact), and adults associated with the larval skin from which they emerged. In the latter case, let the adult harden for a day in a brown paper bag before preserving.
2) Choice of net: It should be large, strong, and as light-weight as possible. Sid uses a modified fish-landing net with ca. 50cm hoop, 2m handle, and a net bag with mesh size a few mm across for reduced air resistance. Dark material is probably best for the net, and it should be tough enough to resist thorns.
3) Approaching dragonflies: Approach slowly, then swing the net as fast as possible from the insect's rear. Don't make a shallow swing: follow through until the dragonfly is deep in the bag, then twist the handle to close the net entrance.
[For dragonflies which rest on the ground, e.g. some Sympetrum, I approach slowly from behind, sit down perhaps 4' behind the dragonfly, and slowly edge the net over the ground (at ground level) toward it. When the net is 1-2" behind the 'fly, I rest it on the ground for a few moments, so the dragonfly will get over any nervousness caused by my approach and the movement of the net. When I think the dragonfly has relaxed its guard, I quickly swoop the net a few inches up and over the dragonfly, almost always catching it. When I miss, it's usually due to impatience: either I tried to pounce when the net was too far away from the dragonfly or without giving it adequate time to settle down before striking - TM]
4) Removing the dragonfly from the net: Hold the wings together over its back and keep the jaws from clamping onto the net (a dragonfly will often hold the net with its jaws until the head is pulled off). Place the dragonfly in an envelope with its wings over its back and the abdomen straight (if it can curl its abdomen to its mouth, it will chew the end of the abdomen off). Place the envelope in a crush-proof container and protect if from excessive heat until you get home. Killing it in the field is not advisable, since it will begin to lose color before you can commence preservation.
DPe mentions an alternative to netting which an entomologist suggested to his entomology club: Shoot them with soapy water from a squeeze bottle or plant sprayer. [Any comments from people who have tried this are invited- TM]
If he doesn't have a net, SC occasionally catches small- to medium-sized dragonflies perched near water by dumping a handful of water on the dragonfly. He estimates it works 5-10% of the time when you hit the dragonfly (it's easy to miss).
DPa used to collect odonates with a "big homemade flyswatter" consisting of a thin bamboo pole topped by a rectangle of window screen: "... very useful for the ones that persist on sitting flat on vegetation on the water surface (many damselflies, also gomphids on shore), but you had to develop some finesse to keep from squashing them too badly." (See Needham and Westfall, 1955, p. 36-7 for details)
Harking back to SB's remark about badminton as useful training for aerial netting, DPa has used a badminton racket to knock crepuscular aeshnids out of the air. He has also caught dragonflies at lights, particularly near water, and has used mist nets to catch dragonflies, particularly aeshnids.
He mentions catching damselflies by hand in the tropics, where many occur in tight places where it is impossible to swing a net. The technique is to approach very slowly from the rear and use the index and middle fingers as a forceps to grab the damselfly. The technique can even be used on dragonflies, if you are skillful. Targets under ideal conditions are the middle of the wings for dragonflies and the abdomen of damselflies.
About hand catching, SC says he occasionally catches dragonflies this way. His method is to approach the dragonfly very slowly from the front, watching the dragonfly's head and legs for signs of disturbance. If the dragonfly tenses, freeze until it appears to relax.
When you get within about 5 feet of the dragonfly, slowly extend your arm out to the side and slowly rotate the arm in the vertical plane like a clock-hand while continuing to slowly approach the dragonfly. As you get progressively closer, reduce the radius of the circle your arm is making (by angling your arm forward from your body), so that your hand, pointer finger extended, slowly spirals in toward the dragonfly. The dragonfly should track the motion of your hand by rotating its head. If your movements are slow and steady, it may be mesmerized and not take alarm and fly away. When your fingers are within a couple inches of the dragonfly, quickly grasp it by the wings ( or whatever else you can get ahold of) between thumb and forefinger or between fore- and middle-finger.
SC states that this method has worked for him with libellulids, especially some Sympetrum species, but hasn't worked with the aeshnids he has tried it on. It is from Korea.
Harris and McCafferty (1977) tested "sticky traps" as a method of catching aquatic insects flying over streams. They found that equal numbers of males and females were caught flying downstream, but that the majority of insects trapped flying upstream (9:1) were females. The only insects which were adept at avoiding the traps were aeshnid dragonflies.
Along these lines, Evans (1993: 63-4) writes: "The nineteenth century naturalist A. R. Wallace reported that natives of the island of Lombok send their children out into the rice paddies with long poles smeared at the top with a sticky substance. When dragonflies alight on these poles, they cannot escape and are served fried with onions."
Hatto (1994) reports on a traditional Japanese boy's method of catching large dragonflies (aeshnids and cordulegastrids), called "buri" or "toriko." Similar to fly-fishing, it takes advantage of the predatory propensities of these dragonflies. The game, for that is what it is, involves tying a weight to either end of a 60-80 cm long silk cord. The apparatus is tossed up in front of the dragonfly which, mistaking the weights for prey, swoops to catch them and is entangled by the string. The weights pull the dragonfly to the ground, alive and unhurt. See Hatto's article for details.
Soberingly, he concludes, "The buri or toriko was a favourite and fascinating game for boys in the summer from at least 160 years ago up to the 1960s. However, with the decreasing number of dragonflies in Japan due to environmental destruction, water pollution and concrete blocking of river banks and lake and pond shores, the sight of young boys using this traditional method is becoming extremely rare. It is now on the verge of extinction. Therefore, we are making efforts to preserve and protect this rather artistic and romantic method of catching dragonflies in Japan." [And the dragonflies, as well, we presume - TM]
Pemberton (1995) quotes Sid Dunkle as reporting a roughly 5% success rate of this technique at a demonstration in Japan in 1993.
Needham (1899) suggests that odonates can often be picked by hand from low bushes near the water following daytime rainshowers. At dusk, some of the larger species may similarly be picked from their nighttime perches, but most settle in too high to reach.
Imagoes may be collected in small, coarse-textured paper bags as they emerge from their nymphal skins or are drying their wings following emergence. Collect the shed skin with the dragonfly.
Needham and Westfall (1955, pp. 40-41) suggest several alternative techniques:
-"As a last resort, some collectors use a gun loaded with dust shot to bring down the biggest ones. They obtain a fair proportion of usable specimens."
-"Wary males may be lured within reach of a net by tethering a female to a stick, holding her out over the water with one hand and swinging a light net with the other." (The "stool-pigeon" method)
-"Perching species will often come to a reed tip set just offshore, if it offer (sic) a good point of outlook over the water. A better place to perch may be a sufficient lure. The collector waiting beside it must have patience to keep stock-still until time for swinging his net."
-"Good specimens may sometimes be taken from spider webs, though only the stoutest species will be found there in fit condition for preservation." (see SC below)
-"Sometimes the big darners are downed in a storm and may be found in the drift line on a lake shore."
SC mentions using spider silk for catching odonates. I paraphrase him at length:
Instead of a metal ring and white net, you have a metal wire and web. Go outside in the morning and find a newly built, fairly large spider web (larger than the diameter of your net-ring). Place your insect net (i.e., the metal ring) on the center of the web, push it slightly so that the web is stuck to the wire, then rotate the ring so that more of the web sticks to it (thus effectively increasing the number of layers you can get from a single web). Multiple layers are necessary for middle- to large-sized dragonflies. For large dragonflies, you may need to use two or more webs, but Soowon has never needed more than three. "Usually," he says, "two fresh webs are enough." You can often return to the same site the next day, where the spider will likely have constructed another web, but he recommends having three or four locations you collect from, so as not to drive the spider away or interfere too much with its feeding. Wet webs don't work well, so collect on a sunny day after the dew has evaporated. Avoid older webs with lots of insects stuck on them or with obvious holes in them. This method is *not* recommended for butterflies, because detachable scales may make it easier for them to escape, and you will lose some of the butterfly's wing pattern to the web. Spider web is reasonably strong, so you can swing the "net" to catch the dragonfly, or use it like the "homemade flyswatter" DPa describes above. The wings stick to the spider web, immobilizing the dragonfly.
Pemberton (1995) mentions several traditional Asian methods in addition to those described above:
1) To catch dragonflies which perch on low vegetation, apply the sticky latex of the jackfruit tree, Artocarpus heterophyllus, to the end of a slender stick or bamboo shoot. Stalk up to the dragonfly and slowly lower the sticky tip of the wand until it is just above the back of the dragonfly. Quickly tap the dragonfly with the sticky wand, and it is caught.
2) Attach a stiff wire or reed to the end of a half meter long stick. Mold a spherical globule of latex on the free tip of the wire or reed. Find a place where dragonflies are flying or perching and swing the latex globule around in a circle. The dragonflies are caught when they attack the globule, apparently mistaking it for prey.
3) Find a place where dragonflies are perching, and hold the latex-laden tip of your pole in front of and higher than the dragonfly's current perch. It will fly to the higher location and be trapped.
4) The most successful technique Pemberton observed in Bali involved stalking up to a perched dragonfly and swiftly sweeping it from its perch with a cupped hand, capturing it with the sweeping hand.
5) In Taiwan, a decoy is made by tying together the seed heads of grasses (no details given, but you can probably figure it out). Wave the decoy at a dragonfly. When it attacks the decoy, grab the dragonfly with your free hand.
Voigts (1973) describes a trap which can be placed in marshes to catch emerging odonates.
BA soaks the dragonfly in acetone, injecting the thorax and abdomen of larger specimens with acetone prior to immersion. Regardless of size, he places specimens in individual paper triangles and soaks the dragonflies for 8-12 hours (depending on size) and damselflies for 4 hours. Due to the volatility and explosiveness of acetone, he stores it in a large jar with hinge-and-clamp top.
[A note on paper triangles: it is my impression, based on two seasons of collecting, that glassine triangles or envelopes seem less likely to damage the specimen's eyes than those made from paper, particularly if you are keeping the insect alive for a period while its digestive tract empties (see PF, below). However, the glassine is less permeable to ethyl acetate than paper, so you have to squirt the poison inside the triangle or envelope to kill the dragonfly expeditiously. Comments? - TM]
SB recommends immersion in acetone for 8-12 hours, changing the acetone regularly between specimens because it hydrates quickly. Colors will be preserved reasonably well, though paler than in life, and the eyes will turn white. The specimens go brittle after treatment, so be sure they are set in the position you want them in prior to immersion. SB mentions that a colleague considers butanol superior to acetone, but he hasn't tried it himself.
RC recommends acetone. Details below.
SD recommends acetone. Details below.
EC states that he and Steve Valley have experimented with different methods of preserving odonates, and finds that injecting the dragonfly with acetone followed by placing the specimen in a glassine envelope or triangle in a jar of acetone, and storing the jar with specimens in a freezer (ca. 0 degrees C) for a week seems to work well for blue dragonflies, such as Aeshna species, but didn't improve preservation of reds.
PF uses 100% acetone for 2-7 days, the specimens placed in glassine envelopes, data in pencil on the envelope, and a paperclip placed vertically alongside the wings (which are placed in standard position, together above the back) to keep them straight. [I have found this very useful, as well - TM].
He maintains the dragonflies alive in a paper envelope for 2-3 days to allow its digestive tract to clear of waste before you kill and preserve the speciment. SC and DPa agree that this is desirable, though DPa considers it optional if you already have too many other things to worry about in the field, a usual occurrence. He recommends that you check them often, since they may not live the full three days and you want to get them into acetone as soon after death as possible. DPa kills his with acetone at the end of a collecting day; specimens from the morning, at least, have had time to defecate.
RO recommends acetone, and sent me a handout on collecting and preservating dragonflies prepared by Sid Dunkle. Details below.
Since I have extended comments on technique from RC and SD (via his own comments, in a handout by him sent to me by RO, and from his Dragonflies of the Florida Peninsula ...), I summarize them in detail here:
a) Keep the acetone in a wide-mouth glass jar or other container inert to it. The wider the mouth, the better. Lid must be leak-proof. Purchase acetone at hardware stores.
b) Kill the dragonfly before inserting the envelope into the jar by placing a few drops of ethyl acetate in the envelope. As soon as the insect is dead, straighten the abdomen and immerse the envelope with dragonfly in the acetone. For larger specimens, inject the thorax and abdomen ventrally with acetone for better infusion. After 24 hours, suck any excess out with a hypodermic needle through the same holes. (If no poison is available, place the envelope and dragonfly in the acetone for killing. After about a minute, remove it from the jar, straighten the abdomen, and replace it in the acetone). If processing more than one dragonfly, stack them upright side-by-side in the soaking jar.
c) Leave the dragonflies in acetone for at least 24 hours. Remove the envelopes for drying when the next day's catch is ready to go into the jar(s). Replace the acetone after four or five uses, when it becomes yellowish.
d) Drain acetone out of the envelopes and dry them and the specimens in a well-ventilated place.
e) When the envelopes are dry, store them upright (like index cards) in tupperware or cardboard boxes that will withstand crushing.
f) If mailing specimens, enclose the box in a larger box, protected on all sides by 3-4" (7-10cm) of packing material.
g) Acetone may not be allowed on commercial airline flights. You may have to purchase it after arriving at your destination. Bring your jars along, however, since suitable ones may be hard to locate in some places
4) If using acetone is impossible, dry the specimens as rapidly as possible after they have been killed. Placing the boxes containing specimen envelopes at close range over or under electric lights is helpful. The faster they dry, the better the color preservation. In humid locales, acetone is a necessity, since air-drying will be difficult.
5) An alternative method is to store the specimens in 70-80% ethanol, either in or without envelopes. If envelopes are not used, specimens can later be removed from the alcohol and dried in envelopes in the correct position.
Both these alternatives (#4 and #5) are less desirable than treatment in acetone.
6) Once the specimens arrive at a museum, take them out of the glassine envelopes and store them permanently in clear cellophane envelopes. Identification and collection data are typed on 3"x5" cards, which are inserted in the cellophane envelopes behind the specimen. The cellophanes are then stored upright in a cabinet like cards in a cardfile.
From a handout courtesy of RO and Dragonflies of the Florida Peninsula ..., pp. 13-14:
1) Collect specimens and keep them alive in separate envelopes or paper triangles until preservation. [For instructions on how to fold a paper triangle. See Knudsen 1972, pp. 147-148 - TM]. Be sure to write collection information on the envelopes in pencil (acetone dissolves many inks). Envelopes containing pairs collected in tandem or wheel position should be marked or associated in some way. Specimens can be kept alive for a day or two in a refrigerator, but some species, especially blue ones, lose their colors until warmed up again.
2) Killing methods: Dropping the dragonfly in a wide-mouthed bottle filled with acetone works in a few seconds. Freezing also works, but blue and green colors will darken, and the specimen will decompose rapidly when thawed. In a pinch (e.g. when acetone isn't available), dragonflies may be killed on the hot dashboard of a car, quickly dried in porous paper, labeled, and protected from insects and breakage.
3) Preservation: After killing, arrange each specimen with wings above the back and abdomen straight in paper triangle or envelope. Write collection data on the envelope in pencil (if you are killing them in the field and haven't done this already, as in step 1), then place the entire envelope in a jar of acetone. Beware of acetone's flammability! Don't breathe the fumes! Soak for 24 hours. Discard the acetone when it becomes yellow like weak urine due to dissolved fats.
Remove the specimens, still in their envelopes, from the acetone and dry them in the air for about a day. It is better to place the specimens in a bag or box with a hair dryer blowing on them for an hour or two. The hair dryer should be set on no heat or the lowest heat and placed about 2 feet from the specimens. Placing the specimens in the breeze from an air conditioner also works well. The specimens are dry when the abdomen and legs do not move with gentle finger pressure, i.e. they have become brittle. Hot air from a furnace or oven melts the fat in the specimens. This is not desirable.
Some fading of colors will occur, and eye color will not be preserved, so make notes on coloration before preserving. [Smithe, 1975 is useful for standardizing color descriptions. - TM]
4) Storage: Store specimens in paper or glassine envelopes or, better, in transparent cellophane envelopes containing a 3"x5" data card. Store envelopes on edge like cards in a cardfile in tightly closing plastic containers, with a few naphthalene moth balls (paradichlorobenzene dissolves some kinds of plastic) for pest control. Do not pin dragonflies because the head and abdomen are prone to breakage if not supported.
5) Mailing specimens: Place the specimens in their envelopes in a small box with padding to prevent them from sliding against each other. Nest this box in a large box with padding, and label the outer box "Fragile" and "Contents: Dead Insects, No Commercial Value."
6) Final disposition: Be sure to state in your will who gets your collection when you go die, so your specimens don't go to waste. [Note: Most countries, and even the states of the United States, have different regulations concerning collecting. Be certain you have any necessary collecting and importing permits. Museums and other repositories may be reluctant to accept your specimens if you can't prove that they were collected in accord with applicable regulations. - TM]
In 1995, I put a follow-up question to Entomo-l: "In looking at older collections of pinned dragonflies, I notice that color seems to be better preserved in many of them than in the specimens I treat in acetone. Is acetoning really the best method of preservation?"
Dennis Paulson replied:
"I suspect the dryness of the surrounding air is critical in air-drying odonates. Anything placed in a rather rigorous desiccating atmosphere probably dries quite well, and some of my air-dried odonates look pretty good. Most of them, however, haven't held their color as well as acetone-preserved specimens. Nor are they as strong (structural integrity), which I consider an equally valid reason for using acetone. That's even if you discount the (somewhat?) lower likelihood of acetone-dried specimens to be subject to pest attack.
Acetone supposedly also takes some of the fat out of specimens, and fat definitely causes deterioration. This may be a problem only in migratory species. I've had Pantala and Tramea and some others ooze grease until they discolored much of the index card on which they were stored. It wasn't until I had been replacing these cards for some years that it dawned on me that they represented some sort of index to fat content, which probably related to migratory status! When we prepare birds we always record the amount of fat, which is highly correlated with migratory status. With acetoning, we're now probably losing that information.
It may be that pinned specimens experience a lower humidity than those stored in glassine envelopes, which of course is the way I've always collected mine. Thus pinned specimens on the average may look better than those originally stored in envelopes.
I think there is a great variation in the way acetoned specimens turn out, depending certainly on the amount of use the acetone has already had, but perhaps even on the acetone itself. A friend of mine bought some in a hardware store this summer, and I was amazed that his specimens didn't look all that good. When he got a new supply, all of a sudden they were fine.
I have had some acetoned specimens in which the eye color preserved moderately well, in most the eyes turn dark, in still others they become very pale. Perhaps again either age or type of acetone. I am pretty consistent in how long I leave specimens in it, ca 24 hours, although I've found I can leave them in for only 12 and they still preserve all right. Might be worth experimenting more with this."
And Bill Mauffray wrote:
"The whole key is drying the Odonate specimen as quickly as possible. Before Acetone, I used to place my newly collected dragonflies in plastic cases in the direct sunlight on my dash board. I would remove from the sunlight before they got crispy. Then I would allow them to air dry in PDB fumes for a few days.
The air drying method worked for most of the greens, browns and yellows as well as powdery pruinose colors like bleus and whites. It did not work at all for the blues in Argia and other zygopts.
The key here is being able to dry the specimens rapidly. In the humid parts of the US and the tropics, the moist air works against this method. You have to use acetone to preserve color patterns here in Florida and in the tropics.
It is true that Acetone destroys the eye color, and pruinose colors (if the acetone is not clean). I still hold out the pruinose specimens for air drying.
It would be nice if we could find another substance that would allow for quick drying, color preservation in both eyes and body, and be readily available no matter where you go (Acetone is hard to get in some countries)."
***Disclaimer: I report the following methods for completeness, but am not vouching for their safety. Do them only in a laboratory where suitable safety precautions can be observed (e.g. working in a fume hood), and know what you are doing. Consult material safety data sheets for any chemicals you use.***
(Recommended for species with metallic colors, e.g. Somatochlora)
Set the specimens on small cards "using cotton" (?). Place the specimens in a tightly closing jar with ethyl acetate. Leave the specimens in the jar over night. After removing the specimens from the jar, leave it exposed to the air for awhile before storing them.
Make a slit beneath the fourth abdominal segment with sharp scissors or a scalpel. Be careful not to damage the external genitalia on segments 2-3 and 8-9. Grasp the intestines with forceps and pull them out. Place the specimen on a plate or tray and gently heat it over any available heat source, being careful not to scorch the specimen. It should not take long to dry by this method. Alternatively, dry it in an oven no hotter than 110 degrees F.
(Requires that you have access to a vacuum desiccator)
a) Room temperature (after Moore 1951):
Puncture the specimens at the membranes separating the abdominal sternites to prevent them from contorting during drying. More than one specimen may be dried at a time, but don't mix dragonflies and damselflies. Fully evacuate the drying chamber for dragonflies, partially evacuate for damselflies (experience will tell you when to stop evacuating for different sized specimens). Use an ample supply of desiccant (in this method, phosphorus pentoxide). After about 24 hours, the specimens should be ready (longer for large batches). The eyes dry last, so when they turn opaque, the specimens are fully dried.
Repressurize the chamber slowly, so the specimens are not damaged by a sudden in-rush of air. If the abdomens of any specimens appear "marred by grease" (female aeshnids are susceptible to this), detach the abdomen and soak it in ethyl acetate for a few hours, then let the abdomen dry and reattach it using gum arabic [Note: I am not personally recommending detaching the abdomen - TM]. Drying should begin as soon after death as possible. Kill red, yellow, and black and yellow species using sulphur dioxide made from "equal small parts of powdered potassium metabisulphite and powdered citric acid, in the bottom of a glass tube, covered with a wad of blotting paper kept moist by a few drops of water." Kill blue and green species with ethyl acetate.
b) Low-temperature drying (after Davies 1954):
This method is notable because it is said to preserve the living color of the eyes, which other methods do not do well. It requires both a glass vacuum desiccator and a freezer. Concentrated sulphuric acid is the drying agent (handle with care). Keep specimens alive long enough to clear their intestines. Kill them (cyanide is recommended) and set them on small cork blocks. Immediately cool the specimens to 2-3 degrees Celsius by placing them in a -10 degree C freezer. Put the blocks with specimens into the desiccator and depressurize to less than 0.5 mm Hg. After five hours, slowly release the vacuum and remove the specimens, which are ready for storage.
When I (TM) process dragonfly specimens, I take standard measurements and do a detailed color description before putting the specimen in acetone. Thus there may be a 1-2 hour delay between the time I remove the specimen from the killing jar and the time I put it in acetone.
To ensure that the preserved specimen will be useful and pleasing to look at, immediately after removing the dragonfly from the killing jar, I lay it out on a paper pad or pinning board and arrange all the parts the way I want them to be in the preserved specimen (e.g. abdomen straight, legs clearly visible and not blocking any important structures, wings aligned above the thorax, penis unsheathed, etc.), using insect pins to brace and hold in position the various body parts. Note: The pins are inserted in the pad or pinning board, *not* through any part of the specimen.
Leaving the pin-braced specimen in position for 10-15 minutes before taking it up to measure and describe makes it much easier to get it back the way you want it before immersing it in acetone later.
I find it much harder to get a pleasing setting if I don't do a preliminary layout before processing. Of course, if there is only a brief delay between killing and submerging in acetone, this is not so critical. Even then, a brief period of air drying in the desired position might make it more likely that the specimen will stay in the desired position when you submerge it.
Acetone dissolves lipids, so if your specimens might be analyzed for lipid content (e.g. in energetics studies), preserve in alcohol. (May, 1992)
Acetone may remove the pruinosity from pruinose dragonflies (e.g. mature male Libellula forensis and L. lydia). See comments by BM in the previous section.
KS, HR, and BM have suggested that acetone may destroy or damage DNA, rendering specimens preserved in acetone unsuitable for molecular biological research (KS was the first to raise the issue). However, TC (based on work done with his colleague, James Purdue) reports:
"We have found no problems for PCR amplifications up to 550 base pairs. No tests have been performed for segments greater than 550 base pairs have been attempted. Specimens treated with formalin present some special problems. The literature contains techniques for dealing with these, but we have not tested them."
BM reports that he receives requests from molecular systematists for specimens which have not been preserved in acetone, but that people doing other sorts of taxonomic research want acetone-treated specimens. He suggests that collectors taking a series of specimens set aside a few to be preserved without acetone, and be sure to label them to that effect. The Internet is also an ideal way for researchers to request freshly collected specimens custom-preserved to meet their needs.
In the meantime, preservation in acetone seems inadvisable if you are doing molecular analyses. KS brings her specimens back from the field alive and freezes them until she needs them (colors may not be preserved, as mentioned under SD in the previous section). Alternatively, she recommends preservation in 95% alcohol. She also implores collectors to keep accurate records of post-capture treatment of their specimens, so molecular researchers will know what chemicals they have been treated with.
"The best way to ensure a specimen's freshness is to store it in liquid nitrogen in the field and then store at about -80C. This is essential for allozyme analysis. A decent yield of DNA can be abstracted from material in 95-100% Ethanol, once again cold temperatures are preferable for long term storage. I suggest injection with alcohol for large specimens since the alcohol may not penetrate the body fast enough to avoid spoilage. If none of this is available or if you are interested in samples already in collections, rapid drying will preserve the DNA to a lesser extent. I recently extracted DNA from a pinned specimen of Apis mellifera that was almost 100 years old. This should only be considered a last-ditch option."
Corbet, Philip S., Cynthia Longfield, and N.W. Moore. 1960. Dragonflies. London: Collins.
Davies, D.A.L. 1954. On the Preservation of Insects by Drying in Vacuo at Low Temperature. Entomologist 87: 34.
Dunkle, Sid. 1989. Dragonflies of the Florida Peninsula, Bermuda, and the Bahamas. Gainesville, FL: Scientific Publishers.
Evans, Howard Ensign. 1993. Life on a Little Known Planet. NY: Lyons & Burford.
Harris, T.L., and W.P. McCafferty. 1977. "Assessing Aquatic Insect Flight Behavior with Sticky Traps." The Great Lakes Entomologist 10(4): 233-239.
Hatto, Y. 1994. 'Buri' or 'Toriko', a Traditional Japanese Method of Catching Dragonflies. Odonatologica 23(3): 283-289.
Herman, Steven G. 1986. The Naturalist's Field Journal: a Manual of Instruction Based on a System Established by Joseph Grinnell. Vermillion, SD: Buteo Books. (ISBN 0-931130-13-1)
Knudsen, Jens W. 1972. Collecting and Preserving Plants and Animals. NY: Harper & Row.
May, Michael L. 1992. Migrating Dragonflies in North America. Argia: The News Journal of the Dragonfly Society of the Americas, July 1992, pp. 4-8.
Moore, B.P. 1951. On Preserving the Colours of Dragonflies and Other Insects. Proc. S. Lond. Ent. Nat. Hist. Soc. 1951: 179.
Needham, James G. 1899. Directions for Collecting and Rearing Dragon
Flies, Stone Flies, and May Flies. Washington,
D.C. Govt. Printing Office. (Bulletin of the United States National
Museum, no. 39)
Needham, James G., and Minter J. Westfall, Jr. 1955. A Manual of the Dragonflies of North America (Anisoptera). Berkeley: University of California Press.
Pemberton, Robert W. 1995. Catching and Eating Dragonflies and Bali and Elsewhere in Asia.. American Entomologist 41(2): 97-99.
Smithe, Frank B. 1975. The Naturalist's Color Guide. NY: American Museum of Natural History. (Order from Special Publications Dept., AMNH, Central Park West at 79th St., New York, NY 10024, USA; ISBN 0-913424-03-X; ca. $20 US)
Tennessen, Ken. 1994. A Collecting Tip for Gomphids. Argia: The News Journal of the Dragonfly Society of the Americas 6(3) : 12.
Voigts, David K. 1973. An Odonate Emergence Trap for Use in Marshes. Proc. Iowa Acad. Sci. 80(2): 67-68.
Question: Are there any good photographic field guides to North American dragonflies which could be used for identifying distinctive species without the necessity of catching and killing individuals?
Everyone recommends Sid Dunkle's guides to dragonflies and damselflies of Florida. While they focus on Florida, Bermuda, and the Bahamas, they cover about 1/3 of the species present in other parts of North America. This is more than any other photographic guides I have come across. They are available from the International Odonata Research Center store.
Dragonflies of the Florida Peninsula ... ISBN 0-94541-723-3
Damselflies of the Florida Peninsula ... ISBN 0-94547-785-3
The Audubon Society Field Guide to North American Insects and Spiders, by Lorus and Margery Milne (New York: Knopf, 1980; ISBN 0-394-50763-0) covers 24 species of dragonflies, 12 of which are not in Dunkle, and 10 species of damselflies. The photos are well-done*, though much smaller than those in SD's books. It is a useful supplement to Dunkle.
*[Since I wrote this in 1994, several professional entomologists have denigrated this guide to me, commenting (among other things) that the photos are not that great. In particular, they said that the colors are often not entomologically correct. Nonetheless, I find this guide useful for general insect identification in conjunction with other field guides and more technical materials. Other general North American insect guides show few odonate species. - TM]
Sharon J. Collman, WSU Cooperative Extension Urban IPM Resource Center, University of Washington, for urging me to contact DPa.
Dan Hilburn, for passing my request on to both Sid Dunkle and Richard Orr.
Alvaro Jaramillo, Vancouver, BC, Canada, for sharing his experiences in and enthusiasm for odonate natural history with me.
David McCorkle, Western Oregon State College, for a suggested contact close to home.
Special thanks to Dr. Dennis Paulson of the Slater Museum of Natural History at the University of Puget Sound, Tacoma, Washington, USA, for answering numerous beginners questions about odonates and odonatology for me.