Eumorpha pandorus ruined in relaxing

[exoticimports]

My buddy found a dead pandorus in his garage; even after 5+ months there it was in great condition, albeit minus the antennae.





So I took it home and tossed it in the relaxing box with some Papilio glaucus. As always, it was simply set on the wet paper towel, and got saturated like everything else does. The fluid is mostly water, with a dash of alcohol to prohibit mold. This is what the results were:





We've discussed water staining on some butterflies such as Colias. Clearly, the water wrecked this sphingid. Note that the Papilio glaucus were fine.

Food for thought.

Chuck

[nomihoudai]

Green turns yellow when relaxed in water. In Europe we have the genus Geometra (Geometridae) that needs to be mounted from fresh samples otherwise they turn yellow.

[Deleted]

Yep….with the green ones, EA and water ruin the coloration. I carefully inject the moth and actually mount it in the field. It’s a pain to do, but absolutely necessary.

[kevinkk]

It looks like a different species now. So- apparently it's going to be very difficult for me to get a collection of green sphinx moths.

[Chris Grinter]

Direct contact with a wet paper towel is bad for hairy moths, you can see where the water crinkled up the wings and matted the hairs on the moth. I relax green leps the same way I do everything else, in a tupperware with just water. Paper towel below and a paper towel on the lid to add humidity. For big things you need to inject a little warm water and come back later in the day and squeeze the thorax to get that water moving around in there. 24hr in the relaxer and they spread very well. But yes some do turn yellow-ish and there isn't anything you can do, even if you spread fresh they sometimes yellow in your collection over time. But some stay pretty green over the years.

[vabrou]

Chuck, FYI, not sure where you heard using alcohol with water is a good thing, but that has just the opposite effects. Nor will it prevent mold or fungus, where did you hear that? You should never place relaxing specimens directly upon a wet paper towel, NEVER, NEVER, NEVER. These are common 'old entomologists tales'.

[Perhaps you might find as informative, a paper I wrote and published in 2014 titled 'How to eliminate drying in delayed processing and spreading of lepidoptera specimens' which works for most all insect orders as well. The procedures I use to spread freshly collected specimens (yesterdays trap pickups) are discussed here. My collecting chambers attached to the traps are either insulated or protected from direct sun rays (strategically placed in heavy vegetation). When specimens are retrieved from the traps the following day, they are fresh as being captured an hour ago, then placed directly into chlorocresol treated relaxing boxes. Then the relaxing boxes sit on my desk another day (now about two days after being captured) (relaxing), then specimens pinned and spread without any problems whatsoever. They sit on the spreading boards for a minimum of 48 hours, then go into a temperature controlled low temperature drying oven for 48 hours or more depending on specimen body size. Thoroughly dried specimens are then removed from boards and placed in low-humidity controlled transfer boxes, and then eventually into the master collection Cornell drawers, housed in a temperature controlled environment, dehumidifier running, annually fumigated, and permanent long term storage in total darkness (Closed cabinets). Using glass top drawers in cabinets without a closeable door is worthless, because in 20, 30, 40, 50...... years, these specimens will be garbage, devoid of color. green specimens are the first ones to turn snow white. Also NEVER, NEVER, NEVER use naphthalene around an insect collection for any purpose.  I have seen huge collections not stored in closed dark containers totally devoid of colors, only gray or white.  
Free open access link: www.academia.edu/14134587/How_to_eliminate_drying_in_delayed_processing_and_spreading_of_lepidoptera_specimens

Consider this method I have used for injecting water into dried specimens for many decades. I first wrote about this matter in 1973, as time progressed I filled jars (glass or plastic) with granular or round briquettes of paradichlorobenzene, then filled jar with tap water to top. Use this water from the jar to fill syringes to inject into thoraxes of dried specimens. Yes, I know this chemical is not soluble in water, but some tiny amount will be injected internally along with the water. Will this be beneficial, time will tell? No detrimental effects have been noted over the decades. As for molds, yeast, mildew and fungus, all of your specimens (including all of us and everything on the planet) are covered in spores, etc of these various organisms. A common promoting factor of all of these is water (humidity).

I have about 40 or so of these chlorocresol relaxing chambers around here all the time. These become dry laying around in air-conditioned environments, and I just add water to use them when that happens. Chlorocresol has embalming qualities.  In the US, the only source for this chemical is BioQuip.  They discontinued this chemical several years ago and I convinced them to bring it back.  They did.  This chemical was first documented in 1961  Literature citation: Tindale, N.B. 1961. The chlorocresol method for field collecting. Jour. Lepid. Soc. 15: 195-197 /font]

[trehopr1]

Everyone has their own take on the caveats of relaxing. As for myself, I never allow any of my specimens to come in physical contact with water or direct dampness.

I have a fine screen on which to place my specimens which is elevated about (1/8 in) above a large sponge.

Prior to putting anything in the relaxer I squeeze out the sponge, I resaturate it with hot water and then I ring out the sponge halfway; then place it in the relaxer. I have a moth ball in each corner of the relaxer and the specimens simply lay on the screen. After 24 hours most things are sufficiently relaxed.

It is the humidity that works on the specimens (best) I have found. Although, I would have to admit that Pieridae have been problematic in the past with very tiny green spots occurring on (some) specimens. So, I now spread all my Pieridae right away whilst fresh and avoiding the relaxer for these.

[exoticimports]

I wonder, was it the alcohol or the water?

[vabrou]

Regarding green geometrids and similar problematic leps, I have personally spread more than 7,000 green geos of 12 or more species I have here at my home property. I have published about a few of them in scientific literature with good quality color images. Using chlorocresol relaxing chambers allows green specimens to remain unaffected in any way. Search for solutions and past related publications, or better yet listen to proofs of these matters by well experienced entomologists. Insect collecting literature by novices and PhDs is filled with misinformation, conjecture, and unproven anecdotal conclusions. People do something a few times and come to false conclusions. One should have actual years and decades of data before having a 'one answer fits all opinion'. Over my lifetime, I personally spread about 1,000,000 lepidoptera specimens, the majority using chlorocresol in relaxing and storage containers. If someone has proof to the contrary of what I say here, please put up NOW.

Some green species have naturally occurring yellow, mignonette and brown phenotypes, and when I publish about these, I try to always show the variations, if possible. I never illustrate poor or discolored specimens. Certain adult geometrids e.g. have naturally occurring color variations of off-green phenotypes associated with emergence during fall and winter months, e.g. N. lixaria (see attached link). These same discoloration issues can and do occur on many chlorophyl colored noctuidae and notodontidae species as well. Use of chlorocresol eliminates this problem on them as well.

As Bill Garthe mentioned previously, ethyl acetate and water will quickly/instantly turn green specimens to yellow. That is well documented in past entomological literature. I have provided 7 free access links to a few species accounts I have published on some of these delicate green lepidoptera (geos and noctuids). Delicate lepidoptera are best spread within 24-48 hours of capture while held in a chlorocresol charged relaxing chamber, and never in direct contact with water or a water impregnated surface.  Use of metal screening to hold relaxing specimens away from direct contact with water is not a good idea as water droplets form on metals due to temperature fluctuations. Plastic or fiberglass screening works much better, as these don't absorb or attract water.

Warning, never ever combine in the same containers chlorocresol with paradichlorobenzene or napthalene. IF you do, you will be sorry you did not heed this warning.

In Tindale 1961 "Chlorocresol method fresh material may be held for several months or even almost indefinitely. A very sensitive test of the usefulness of the method is shown by the fact that it is usually impossible to hold Geometrid moths of delicate fugitive green and blue colors for many weeks without damage". free access Link to that publication below.


www.academia.edu/540165/Nemoria_lixaria_Guen%C3%A9e_1858_Lepidoptera_Geometridae_in_Louisiana
www.academia.edu/428173/Dyspteris_abortivaria_Herrich_Schaffer_1855_Lepidoptera_Geometridae_in_Louisiana
www.academia.edu/34755411/Dichorda_iridaria_Guenee_1857_Lepidoptera_Geometridae_in_Louisiana
www.academia.edu/232096/Phrygionis_privignaria_Guen%C3%A9e_1857_Lepidoptera_Geometridae_in_Louisiana

www.academia.edu/1021407/Leuconycta_diphteroides_Guenee_1852_and_Leuconycta_lepidula_Grote_1874_in_Louisiana
www.academia.edu/34755474/Agriopodes_fallax_Herrich_Schaffer_1854_Lepidoptera_Noctuidae_in_Louisiana
www.academia.edu/30567504/Feralia_major_J_B_Smith_in_southeast_Louisiana_So_Lepid_News

images.peabody.yale.edu/lepsoc/jls/1960s/1961/1961-15(3)195-Tindale.pdf

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[kevinkk]

coming to false conclusions is something that happens in other disciplines as well.

Vabrou- I'm wondering if you've looked at the blog section and the article/video of "how to relax a butterfly"
and what you think about that.

[vabrou]

Kevinkk, regarding the relaxing video with Madagascar moth. I see this may work for some butterflies/moths, but I don't think it is good to have any specimen externally in direct contact with liquid water. I have been pinning, spreading lepidoptera for about 66 years since a child. The majority of the approximate 1,000,000 specimens I have personally processed during my lifetime have been pinned/spread within a day or two of capturing them, none having been allowed to dry out before spreading. This makes for the highest quality specimens.  All of my life I have religiously run my traps every day, and process as many as I can, every day of the year, because tomorrow there will be hundreds of thousands or more to deal with, and the next day, and the next day.  I pick out the cream of the crop of captures and 99.999999% of the rest wind up being discarded. Moths go from trap collection chamber directly into chlorocresol relaxing containers. Once in the chlorocresol relaxing containers, they can stay there for days to more than a week or more, without any adverse effects. I do have a world collection of Eudocima moths which mostly come to me as dried papered specimens of which I spread by a special technique of cutting the wing muscles below all four wings and also often injecting water into the head/eyes which quickly relaxes the antennae and travels into the thorax, and sometimes injecting water internally at wing bases of thorax. But first the papered specimens spend a few days in the chlorocresol relaxing box. These are large moths with very strong wing muscles. I once had a world collection of Sphingidae (about 40,000 specimens) and these too were vey difficult to spread because of strong wing muscles. Back in 1973, I first published a one page note about injecting water into dried specimens using hypodermic syringes. And since that time have fine tuned this method. External wetting of lepidoptera can occur with this injection method as well. Injecting coleoptera usually works well for all dark colored specimens. Though I capture coleoptera and the majority of all beetles I collected over the past 40 years, and want to keep (live or dead) go right into jars filled with 70% isopropyl alcohol. These can remain there for days, week, months or longer. Never us 90% isopropyl or ethanol for beetles. I just place a label on the jars with date and location to process later. The alcohol wash cleans the dirt and debris besides dispatching the specimens. A side effect of me driving around on my tractor to all my traps with jars filled with beetles in alcohol is that the rough terrain bounces the jars around and the beetles are agitated pretty clean by the time I get back to the house.  Other collectors always remark, 'how do you get your beetles so clean'?

[58chevy]

Thanks, Vernon. This is the best information on relaxing I've ever come across. I wish I had known about chlorocresol when I was a young collector. Does Bioquip still carry it? I've heard they're having a tough time staying in business.

[vabrou]

Yes BioQuip has it (chlorocresol), I just received a few cans last month. Owner is retiring and considering the whackos running California into the ground, I'd get the hell out too ----long ago. Thousands amateur and professional entomologist have left that state in recent years. Wish they would stay away from Louisiana, because they bring with them the same lunacy they are fleeing from.

[kevinkk]

Nothing ever stays the same, I find change to be problematic at best. I wonder how Biouip did taking over Combined Scientific.
I guess I made it out of Cali at the right time, only to come back to my home state of Oregon-
Losing Bioquip would be unfortunate. Sometimes I buy from Carolina Bio, time will tell.

[exoticimports]

I used chlorocresol once in 1998 in Ecuador. It melted and merged with the specimens, which was a mess.

Chuck