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There doesn't seem to be common to keep a fly collection in alcohol. Is there someone who does this and can tell us a bit about the equipment and best practices? I wonder what container is used for this. The reason I ask is I got a few nice small plastic micro tubes (2ml) from a collector to send beetles by mail (but not in alcohol). Not sure what it is named in english (in swedish mikror?r/kryor?r). I suspect that insects kept in alcohol is stored in small glass tubes and not plastic ones!?

I'm also interested in how storing other insects in alcohol is done.

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it is enough to use ethanol 70%. Plastic vials are ok. I never had any problems to keep up flies inside plastic vials! Alas, the glass can broken... :S I would advice plastic vials tight and using ethanol 70%. If you want to send the fly for DNA analysis use ethanol 95% (or 90%). NEVER use formaldehyde because it alters the structures of the specimens.

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I can share with you my negative experience :@
After Paul and Kahis advises in:
http://www.diptera.info/forum/viewthread.php?forum_id=21&thread_id=6523
I did try to collect in alcohol and pin afterwards.
My test:
1. Pinning from alcohol requires much more job and time than pinning of fresh flies.
2. Pinning fresh flies in sito permits to get much better order: legs, genitalia and whatever we need to examen on this fly.

I go on pinning fly the same day I collect its!
Nikita

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hi crex,

i think for that small plastic vials the name is Eppedorf tube (at least in romanian... a direct translation... where is the case).

my oppinion is that this are very good for keeping. but one advice... not keep beetle or anything else without alcohol for to much time. otherwise there will develop a lor of mold (i'm telling this from my experience).

just collect and trow the fly/beetle/other insect (except butterflies) inside.

hope this helps you,
cosmln

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I do the same, Nikita! :)

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I would echo Nikita's comments. Pinning from fresh-caught and freshly killed specimens is always the most preferable way, in my opinion. Collecting into alcohol is only done when it is absolutely necessary: 1. where the specimen was trapped into alcohol (eg. Malaise or Pan Trap); 2. a specimen is being sent in the post by someone who doesn't have pins etc; 3. or other 'exceptional circumstances'.

Under #3 I list my trip to French Guyana where I decided to catch mainly large bees & wasps and the best way to kill them was to drop them into alcohol - but even then I still got stung once! :( We were moving around the country too much to allow me to pin specimens each evening and the humidity out there would have prevented them drying anyway. So, alcohol was a good killing agent and it helped preserve & protect the specimens and store them in a relatively small space. :) But most of the resulting specimens were very hard to set because they had contracted all their muscles. Luckily for me I didn't mind that too much and I side-pinned almost everything anyway. :D

Chris R.

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Hehe, I never said that you should collect in alcohol and pin them afterwards. I keep them in alcohol.

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Thanks for the response. I meant really to store the insects in alcohol, not just temporarily keep them during trips etc. I see now that no one seems to do that, i.e. keep (store) his/hers collection entirely in alcohol without pinning the specimens.

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Good tip on Eppendorf tubes ... didn't see Pauls post. Perhaps he could expound on his views if he keeps his collection in alcohol!?

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eppendorf tubes are too small... I never saw eppendorf tubes bigger than 4 cm? I don?t recall well, but they usually are pretty small. But they are very tight what is good. ;)

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I do store all my specimens in alcohol now.
In 1.5ml Eppendorf tubes the smaller ones and in bigger tubes the rest.
Take care with the caps, they does not fit perfectly sometimes, and the alcohol evaporates. The tubes can be put in hermetic jars filled with alcohol.
Even it is good that you have sufficient room in your freezer for your jars.

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Why do you keep the collection in the freezer? :o

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I keep most of my specimens in alcohol, partly because of the number - it would take too much time and space to pin or card-mount them all. And partly because dry specimens tend to go mouldy eventually in this part of the world. Entomologists here who keep their houses much warmer than I do still have problems with mould, mites and Psocoptera.

I find there is no problem with extracting the genitalia from alcohol- stored specimens. In fact it is often easier than from the fresh specimen. The problems are that colours are often very different when the fly is in alcohol, and if the bodies have leaked grease into the alcohol, the specimen will be thinly coated with it even after you have dried it out. That makes it hard to see dusting.

I use 70% industrial meths and 30% water, and it is best to use distilled water, especially if you are in a calcareous district, because calcium compounds tend to precipitate out. It doesn't have to be perfectly distilled water. I use the contents of a dehumidifier.

One drawback is when you want to compare your catch with reference specimens. It is time-consuming picking them all out of their tubes, and because each specimen doesn't have a label attached like it would if it was pinned, you need to work out a system for remembering which came out of which tube.

John Bratton

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As a summary (or my opinion that is):

Pros:
* Pinnging: Ease of use - good for reference collections, if you live in a climate where a dry collection is easy to keep.
* Alcohol: good for mass storage of large samples, and for soft insects (most nematocera).

Cons:
* Pinning is slow. Soft insects will not keep their shape. Pinned specimens are easily damaged by careless handling or pests - also mold in some climates.
* Wet specimens are difficult to compare with dry flies. Some details can be difficult to see. Colour will change with time. Collection needs regular maintainance.

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I don't think I'll be able to keep a pinned collection, so small alcohol tubes could be an alternative for a tiny collection. Do all use Eppendorf tubes as containers? The few tiny tubes I've got are not pointed in lower end of the tube. I also have seen in lab equipment shops that there are nice little boxes to store these tubes. How about plastic vs glass? The ones that I think is glass seems to be transperent which maybe means that you wouldn't even need to take it out of the tube to study the bugs ... but if you keep many bugs in each tube then that probably would not be as good.

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I use tubes with screw caps and a rubber lining in the cap. For long-term storage it is advisable to keep these separate tubes in a larger jar with alcohol or do an annual check to see if alcohol levels are acceptable. The rubber lining may start to decay, so the caps should be checked as well to see if the need to be replaced.
Alternatively, you can use glass vials with cotton wool to close them and keep those in a larger jar with alcohol. Disadvantage of cotton wool may be that smaller flies get stuck in the cotton wool.
In general, I'd prefer glass over plastic even though glass, on average, will be more expensive. Plastic may degrade over time (burst) and glass may give enough visibility to examine specimens without having to take them out. Problems with the number of specimens only arise when there are really many of them in relation to tube size and when too much alcohol is in the tube. I store one species per tube, unless the specimens will not be identified (for example, all Sciaridae will go into one tube, all Cecidomyiidae, all Chloropidae, all Calyptrates, etc.).

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Thanks Paul. Do you use 2ml tubes or even smaller ones?

If anyone got photos to show the vials or Eppendorf tubes and the way you store them it would be interesting to see. I found these nice 2ml Larvengl?ser with boxes to store it.

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I have 5 ml tubes (50*13 mm).

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Ohh, large ones. Any hints on the systems for labeling the collection. Inside the tubes, sticker on outside of tubes or just some number on the lid and notes on the side?

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Inside ONLY vegetal paper and written by a pencil. :) Outside I put vegetal paper and use a transparent gum tape, so label cannot be lost. ;) (really, I use two, one for the stopper, and another one, on the vial.)
I put in that label, species name, date, location, and sometimes male/female symbols. ;) I put a code as well.

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I print the labels with a cheap laser printer in acid free paper and put them inside the Eppendorfs. Country, province, municipality, habitat, UTM coordinate, altitude, day and leg figure in the label.

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I do use also glass vials of several sizes.
I am not an expert, may be someone can give you (and me) better ideas.

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Thank you Carnota for the illustrations. I like the vials with flat bottom best. I have been browsing online Lab Equipment catalogues. The size of the tubes (in ml) seems to be pretty standardized. Best if I find a company in Sweden where I can buy it. I found some, but not all sells to private persons. I'm looking for the transparent, flat-bottomed tubes with twist-off caps.

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I have been using alcohol for about 30 years (for storing Diptera) and offer a few comments.
Firstly, change the alcohol a week or two after collecting the specimen as the original fluid will become diluted. After that, its probably OK.
I always use screw capped plastic vials (don't like Eppendorf's as the caps can pop off). These vials are sold by various manufacturers in different sizes as cryopreservation tubes and are highly inert and stable. Adding glycerol at 5% will minimize risks if it drys out.
Keep specimens in the dark and preferably cool/cold and colour losses are reduced.
If you want to subsequently pin the specimen, hold it in ethyl acetate until the ethanol has exchanged and then dry it out on filter paper (you will find that oily flies like therevids loose their fat and become much better than if you pinned them immediately).
Normal 'india' ink seems to be stable for at least 30 years with no sign of deterioration but make sure the ink is absolutely dry before you put the label in the tube. I have noticed no deterioation with computer-printed labels over about 10 years and suspect that they are OK too.
A problem with wet preserved material is that dusting is hard to see . However it is usually possible with a bit of experience and changing the lighting and background carefully. I find that alcohol preserves morphology (apart from dusting) much better than does dry-mounting but colour can be a problem. Experience with a particular key usually solves this problem but not always, especially as wet material tends to fade.
One usuful tip; if your flies have important features that are hidden when the specimen shrivels on drying (eg. legs that close up hiding setae, genitalia that shrink and distort etc). Collect then into Gaults solution or similar (even dilute salt solution will do):- all the legs straighten out and everscible structures expand; you can then wah them with water before putting them in alcohol which will 'fix' the structures in their expanded state:- if you do this you may find yourself performing far fewer genutalia dissections!
Lastly; if you are in the business of describing species, please state if its a dry or wet specimen that is being described. e.g its common to find a black thorax with yellow dusting and the thorax could equally be described as black or yellow if described from dry (so many keys fall into this error and even famous dipterists do!). Wet material is more likely to be described as having a black thorax with yellow dusting and hence would be more accurate..........

One question I do have:- what to do with genitalia once they have become separated:- if they are put back with the specimen they might get lost but it would be dreadful practice to put them in another tube from the original specimen.
I'd appreciate any thoughts on this

hope it helps
cheers
Adrian

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If the specimen is going to be stored long-term in alcohol (and there are arguments for extracting and pinning or slide-mounting all specimens that are to be kept) then you can put disected genitalia in a glass or plastic microvial and store that in the tube with the specimen.

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Could anyone suggest a suitable supplier for micro-vials for this type of storage? I have found a couple of companies in America and Australia which supply them, but none closer!

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If you just want small quantities then I think Watkins & Doncaster sell a variety of little stoppered glass bottles: http://www.watdon..._home.html

You could also try Henshaws - or any supplier of microscopy equipment perhaps.

I will ask around to see if I can get some pointers from my lab contacts. :)

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Show us some image of those vials. :)

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If one puts a microvial in with the same tube as the specimen it would have to be secured in some way otherwise it would be like putting the specimen in a bead mill:- it would soon be utterly destroyed.
So presumably the microvial has to be secured somehow inside the specimen tube, in a way where it will not move, there are no dead spaces into which the specimen can move, and in which both it and the specimen can easily be removed for examination.
How is this done?
cheers
Adrian

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It can be held in place with a rolled-up piece of tissue, at the bottom of the tube. To examine it, you remove the fly from the tube, then remove the microvial, then open the microvial and get the genitalia out. (It is tedious - and is one of the reasons why long-term storage of reference material is better as pinned specimens.)

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Thanks Tony
I'll give it a try. I presume that best to use optical tissue than some paper based one (tend to break into lots of bits on long storage).
I have to take slight (but not pedantic) issue with your view that dry material is always better for long term storage. Sometimes it is, sometimes it isn't. I wouldn't keep a cecidomyiid on a pin but would keep a syrphid that way.
Generally I favour alcohol but its a case of 'horses for courses'
Dry material is very prone to damage and while I am sure that, like me, you can think of 150 year old types which are still in excellent condition; you can probably also think of many sad damaged specimens which have not stood the test of time. Basically, if you use dry material frequently for a long period, damage is more or less inevitable. Wet material is buffered against vibration damage by the viscosity of the fluid and does seem to survive well if treated sensitively (although I'll only conclude my argument in 100 years or so!). Modern containors mean that there is no reason why a vial needs to have its alcohol topped up for decades (probably 100's of years in reality) and inert plastic technology is such that exposure to air pollution, acid leachates etc is virtually nil. There is no excuse nowardays (except being poor!) for the flat-earth technology of upturning tubes in jars of alcohol etc etc (even though many current texts still groan out the neccessity of this ancient and arcane ritual.
However, my main reason for generally favouring wet preservation is that morphology is better maintained. Many dry specimens distort as they dry and I have many times, when examining a type, marvelled at the flight of fancy (or perhaps better:- educated guesswork) that enabled the original author to imagine what the thing actually looks like in life rather than what they had before them!
Just one specific example of this:- the empidid genus Monodromia Collin was characterised by having distinctly upturned antennae. They aren't actually, when seen in life or in wet material:- its just that Collin (who was an amazingly competent taxonomist) had a shrivelled up type to deal with and never saw its true morphology. I contend that were Collin have been using wet material he never would have made this bizarre error
Incidentally, I often find that if I must dry a specimen from alcohol, it can be better than dry-mounting it directly. One has to be careful that the wings etc remain open on drying. Material critical point dried from alcohol can be supurb (if brittle) but CPD equipment is expensive and not available to most.
Just some thoughts. I think we should keep open minds about preservation techniques and use which combination best serves our purpose.
Thanks again for the feedback
cheers
Adrian

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So long as there is only one fly in the tube, the genitalia shouldn't get lost. I make a note in the record book and put an extra label in the tube to remind me that the genitalia are loose in the tube.

Another option is to mount the genitalia on card so they can be pinned, although they are then separate from the rest of the specimen. There is a substance called Dimethyl Hydantoin Formaldehyde or DMHF that the coleopterists use for mounting beetle genitalia. You buy it as a white crystal and dissolve it 50/50 in water to produce a clear treacle. With a pinhead you put a tiny spot on a piece of card and then place the genitalia in. The genitalia need to be wet so that they don't contain air bubbles and float, but not too wet so that they don't dilute the DMHF. I take them out the alcohol, just touch them on to tissue paper so that 90% of the liquid is sucked off by capillary action, then put the genitalia into the DMHF. You then have about a minute or two to manoeuvre the genitalia with pins into the position you want them, after which the DMHF gets too stiff. It soon sets to give a clear hard protective capsule over the genitalia, with excellent optical properties. If in the future the DMHF gets cracked or scratched or you really need a different view, the DMHF can be redissolved in alcohol and the genitalia extracted. Or if you expose DMHF to fumes of ethyl acetate, it just vanishes into the air leaving the genitalia exposed on the card.

The biggest problem with DMHF is how to get some. Apparently it is a component in hair spray. You could buy it by the kilo but that is very expensive when 5 mls would be enough to last most amateurs a lifetime. But it seems to be very difficult to find for sale in any quantity these days, so if any Diptera.info members find a source, please let us know. However, this recent mailing on the British beetles e-mail group may offer an alternative:

?Entomologist (and retired chemist) Juan De Ferrer who lives in Algeciras, Spain, has recently been selling small (c. 20 ml) bottles of "improved PVP" solution to EU countries, except the UK, for about 4 Euros each. PVP is very slightly amber in colour but is otherwise identical to DMHF (The Coleopterist 14: 29-35}.?

Unfortunately, no contact details for Juan de Ferrer were given. Perhaps one of our Spanish correspondents knows him? I don?t know what PVP stands for.

John Bratton

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here John Bratton :)

Juan de Ferrer Andreu.

Algeciras , C?DIZ

jdeferrer (at) wanadoo.es

Cole?pteros, Histeridae 4 euros per each? :o

Saludos, from Spain. :)

Cumprimentos, from Portugal. :D

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polyvinyl pyridine

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I think the main argument against long-term storage in alcohol is that without adequate fixing, and carefully controlled concentrations, the material will eventually disarticulate. I agree completely with what you say about damage to dry specimens (not to mention the dreaded Anthrenus! :() but I do find that a dry reference collection is much easier to work with - I can line up half a dozen specimens of closely related species and compare them directly - they are never separated from their data or determination labels, so never go back in the wrong place - not so easy to accomplish with specimens out of tubes! I wouldn't pin a cecidomyid, or a sciarid, or a phorid, but I would slide-mount them for examination and long-term storage. But then I need to decide what mountant to use (but that's another thread! :))

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As a matter of fact, I think Cecidomyid should never pinned... they are so fragile! I put always these flies in ethanol 70% (i need to get some glicerin as well... I use this thumb's rule: 1 drop of glicerin for small, very small flies; 2 drops for medium flies and 3-4 drops for the big ones in a small vial (1 cm diameter, about 4 cm lenght.)

Hopefully this weekend I will show the vials I use. The big problem is that it is not easy to see the specimens through the plastic surface... but the vials have good quality. And we can buy 1000 vials for 45 euros....

I use vegetal paper inside the vial with data (written in pencil, of course) and it works very well! The data are well visible in spite of being a plastic surface... :) I put the label outside the vial on the cap (?) and surrounding the plastic surface at least 1/4 of the vial.

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Thanks John Tony & Jorge
Unfortunately, I mostly work with smallish species, invariably 5mm or less and often less than 2mm in length:- the genit are correspondingly small and could easily get lost in the tube.
I do often mount up the genit prep for dried material (usually in euparal between 2X6mm coverslips inserted in card with a similar hole in it and then mounted on a pin) but doing this for a wet specimen and thus separating it from its genitalia is not an option.
There are of course many problems with any solid mounted approach but a major one for me is that the genit can't be rotated to see its 3-dimensional structure.
I think Tony's point about ease of examining dry material is a good one although a bit of practice, a fine brush and suitable watch glasses or similar, the difference diminishes greatly.
The potential to separate specimens from their labels is very real however and needs a considered & methodical approach.

I really don't think that there is a single and simple solution to what is the 'best' way to preserve Diptera. Each method has merits and demerits depending on objectives, personal preferences, facilities and resources. We just need to be aware of these advantages and limitations and be alive to possible ways to improve things.

A (slightly) related topic which perhaps should be a separate thread:-
I have recently been experimenting with collecting bulk samples into lactic acid which partially clears and fixes the specimens. Colour is changed of course but the partial clearing seems to not only make the genitalia more examinable, but also enables many other external features to be seen more clearly. Specimens can be transferred to alcohol or slide-mounted subsequently.
It might be a technique with potential for groups where morphology is more important than colour perhaps?
cheers
Adrian

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Thanks again for all input on this subject. Specially to Adrian.

Adrian, what size of microtubes (vials) do you use and do you also (as Paul) keep only one speciemen per vial?

About the transparent vials. Is it possible to examine the specimen through the "glass" or do you need to take it out of the vial?

Where?

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I mostly use 1.5ml tubes but larger ones are available (5 and 10 ml I think).
I have a large stock of these tubes I got a long time ago. They are only semi-transparent and you have to take the specimen out of the tube to see it well enough to do anything with it although the labels are clearly visible (from time to time I have been sent clearer vials so they are available somewhere but I still think that they arn't clear enough to get a good view of whats inside
I usually keep just 1-5 specimens inside each tube but sometimes as many as 50:- it can be a good way to keep bulk material in a collection.

cheers
Adrian

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I have big problem with glued-up wings, when I am getting my flies off the 70% ethanol. I want to pin the specimen then, so do you have any tips how to do that? How to dry? When I use some paper (vegetal), the wings are still distorted or glued on body or glued-up with each other.:( When I give the specimen back into the ethanol, wings are nicely straight.

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When you take them out the alcohol, try to arrange them on the tissue paper with the wings stuck to each other above the flies back. It helps if you pick them out by their feet so that the wings are the last part to leave the alcohol. Then when it has dried, maybe 30 minutes later, take a pin, insert it between the wing bases, and gently move it towards the wing tips so that it breaks the seal between the wings. With luck, they will spring apart but stay straight.

John Bratton

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In Lisbon. The firm calls Vreis, They don?t have a website! :( But there is just this: http://www.guiane...file/vreis
They sell ONLY if we go there! You can try to write to them (email in the link I've sent). Maybe they can help with your request. I'm not sure if they have an international service.

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or maybe ???? .... ;)
you can help us???
;) :)

cosmln

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yes... cosmin put in my paypal account 90 euros and I will help you right now. :D:D

Well, for those who are interested please email me. (it will take a while, though)

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hihi, thanks Jorge, for now no need.
jus wait for my parcel to come from Germany (i hope he will come faster :) ).

thanks anyway,
cosmln

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Agreed. Wings can be a problem with some flies. I dry mine by washing first in 100% ethanol and then 24 h in ethyl acetate. I then pick up the .wet. fly from the EtAc using very fine forceps and place it carefully on fclean fine grade filter paper (using ventilation as the EtAc isn't good for you). As I place it on the paper, I make sure that the tip of the lower wing contacts first after which the rest of the wing usually snaps flat onto the paper (surface tension effect). I then release the insect so that it falls onto the paper. Usually the other wing will remain reasonably flat but if it does not, a very fine pin can be used to insert into any folds to gently lever them apart.
You will always have problems if you dry directly from alcohol:- firstly, they are not really dry as ethanol is not a good dehydrating agent. Secondly, it evapourates too slowly and surface tension of the liquid is much higher than with EtAc and will ensure that any fold with liquid trapped in between will stay folded. The evapouration rate of EtAc is so rapid that there is a slight gassing out effect as it dries:- tending to force apart folds in the wing membrane.
If you have the kit, critical point drying tends to leave wings fairly well extended. Its quite good (but expensive) but I only use is for drying bulk samples where I havn't the time to spend preparing every specimen with extreme care.
Hope this helps
cheers
Adrian

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Thank you, Adrian and John, unfortunately the fastest method isn't very effective (you need luck:]) and the better method is expensive and takes time :[

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A comment on BugGuide states "I've had no trouble with this cheap way of mimicing "critical point drying"! Works well with flies and some other softer insects that tend to shrivel when dried".

I haven't tried it, perhaps someone here will and let us know if it works:D

SEE HERE

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When examining the specimens stored in alcohol I assume you don't need to dry them first!? I don't have a collection in alcohol, but I've seen flies drowned in water and they look like a mess. Maybe one just has to get used to seeing them that way ... B)

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Practically all flies *can* be identified without drying them, but many keys rely on characters best seen on fresh or dry specimens: dusting can be very difficult to see in wet specimens. For a beginner dry material is easier, but with experience the difference is slight.

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Just a question to Adrian: could you please estimate an average consumption of the Absolute per 100 4-5 mm Fannia/Tephritis-sized flies to dry them before EthAc?

We have got 2-3 thousans of ulidiids from M. von T. and T. van H. in glass tubes, and are doing nothing with that stuff but thinking of technique to pin-and-dry them in the best and fastest way...

And does anybody have an idea how to prevent specimens pinned from alcohol from slidin'turnin'n'jumping on minutien. Live pinned bugs are glued to pins with their juices, but alcohol is not so sticky. :(

I normally use a translucent vinyl glue to fix movable flies on pins (and to repair broken wings, too), but so far saw that glue only in Belgium. They (ADER is the company) produce it for schools; it's more liquid than the common "white" PolyVinAcetate and crystal transparent and is perfect for wing breaches repairing.

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Contracting muscles is exactly the reason why, whenever I store caught insects in alcohol (when travelling indeed, for instance when there's no place for boxes in my luggage), I always kill them with ethyl-acetate (forgot the english word for it) first and let them get a little 'soft'. Admitted, it's more timeconsuming than dropping them in the ethanol directly and you need extra tubes/bottles and administration not to mix up the data.
Also, back home, I try to pin them asap. The longer in ethanol, the more bodily fluids are exchanged with the ethanol, the more loss of color and sometimes of structure.
My experience covers Syrphidae mostly, I must add ;)

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Just a question to Adrian: could you please estimate an average consumption of the Absolute per 100 4-5 mm Fannia/Tephritis-sized flies to dry them before EthAc?

I reckon a 100:1 ratio of alcohol to fly (by volume). This will strip most of the water out but the remaining 1% approx will be 'boundary' water stuck to proteins etc by van der Walls interaction and you can't get rid of this no matter how much alcohol you use or how long you leave it to equillibrate. (a 1% water content is probably dryer than if the fly was conventionally pinned from fresh when one might expect 3-5% perhaps?) In practice, once a specimen has equillibrated at the 100:1 ratio you should replace with fresh alcohol and you could then get away with a lower ratio for longer storage.
Similarly, a 100:1 ratio of Et-Ac/fly would remove c99% of the free alcohol from the specimen which is enough for it to dry well
Can't really help with the revolving on the pin situation as I normally card point flies dried from alcohol. My guess is that the alcohol and EtAc dissolve out the fats which would normally help bind the specimen on the pin if pinned from fresh. I suppose glue is the solution.
Hope this helps
Adrian

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This is a very interesting forum. I would like to add my experiences. Having to deal recently with a large collection of Anthomyiidae collected in the Dolomites in Malaise traps (at high altitude and therefore very valuable) I researched methods of extracting the flies from alcohol, and mounting them dry. This was necessary because there were a number of undescribed species present, and many species previously little known.
I found on an American Museum website that acetone was mentioned as a water-extracting and hardening agent. This worked extremely well. The method I used was:

1. Remove a number of flies from the alcohol and drain on filter paper, then drop into a small container containing acetone.
2. Leave for approx 3 hours.
3. Remove one at a time and pin with a micropin sideways (below the wing base). Drain acetone off with a piece of filter paper.
4. Pin onto a small block of plastozote and under the microscope pull out the legs, and genitalia, if a male. Unnfold the wings if necessary with a pin. Generally the membrane will be stiffenned enough, only leaving the hind margin a bit rough. Luckily the wing venation is not important in anthomyiida!
5. Blow on the fly gently to remove remaining acetone.
6. Thats it! Before staging it may be necessary to put a small amount of glue on the pin below the specimen as they are not stuck to the pin (no natuural juices); they can spin round the pin.
It is important to deal with one specimen at a time, as the drying iis fairly quick.
I have dealt with over 300 specimens this way, in batches of 10.
You will be amazed with the results, only slight cuticle shrinking in some specimens. Material prepared in this way is generally superior to most museum material, which in the past was not collected by dipterists. It is not so good as material collected and freshly pinned by oneself of course. Another advantage is that acetone can be purchased (in England) from any chemist

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That's pretty much what I do, with one exception: I pin the fly before the acetone bath.

Keeping wings open is difficult especially with small, soft flies. I have found two ways to keep at least one wing from folding.

Method one is slower and labour-intensive, but gives the best results: Remove the flies one by one from acetone and immediately bring one wing (or both wings, depending on position) in contact with a flat piece of paper. The wet wing(s) will stick to the paper surface and will neatly unfold when the fly is flightly moved on the paper. Let the acetore evaporate until the wing separates from the paper. Do not use filter paper - it will drain the acedone before the wings have time to unfold. Normal office paper works, slightly thicker paper is even better especially for smaller flies, where the size of the acetone drop carried with the fly is small. With this style it is usually possible to 'rescue' both wings.

Method two is not as good, but it is faster and can be used for larger number of flies. I use it regularly for agromyzids and drosophilids: Micropin each fly asymmetrically. Push each pin with a fly into a flat piece of plastic foam until one wing rests again the foam surface. Let the foam piece float in acetone (flies downwards) for a few hours. Lift the foam out and dry. Remove the flies when the foam surface is dry but before the flies have dried through. Some foams will melt in actone - test your setup before trying it with flies!

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Flies in acetone

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Opening wings agains a slip of paper

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Style 2: micropinned agromyzids waiting for their bath

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Results: Sapromyza (Lauxaniidae). The wings are open and undamaged, but slightly 'wavy', a typical result for style 1.

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I do prefer fresh pinned flies. But life is life and I have to deal with material from alcohol too.
An additional problem I discovered is:
most of bristles on Muscidae head and scutum are broken.
What tubes and way of its storage may be recomended for storing flies in field, during trip and posting, please?

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Hw do you control the atmosphere using the acetone? Not the most pleasant thing to be inhaling for longer times at an end.

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1. I use small containers with only enough acetone to cover the flies
2. As I don't pin the flies before putting in acetone I can get 10 flies in a very small amount
3. Keep lids on except when transferring flies
4. Hold my breath as far as possible whilst transferring flies

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Some photos.

EDIT Paul Beuk: screenshot from ealier attached pdf added.

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Can someone tell me why my photo attachments appear only as the file name, and don't open on the thread?

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pdf files are not really images and they are treated differently depending on the browser. The forum don't even try to format them as images, which is only sensible. Can't really tell why the first one failed since I cannot see the input you send.

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Thanks Paul and Kahis

I'll try another reduced to 50% size. If this does not work I will try other formats

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I have converted them to JPEG and reduced the size. If you don't succeed at first, try again!!

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Success at last!

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I would recommend not to use acetone, and in its place use hexamethyldisilazane (HMDS) http://www.phorid.net/phoridae/dry.htm
Specimens after HMDS not as fragile as acetone after applying

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... and where can I buy HMDS for 5?/l? ;)

Also, I understand rhat HMDS is significantly more dangerous than acetone. For me, 'acetonised' flies are good enough, unless there are large differences in the long-term preservation between the methods.

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Now that I found a company that sells micro tubes in Sweden I need some 70% alcohol, but where do one find that? I know there are different laws and rules for selling this so maybe it's no good asking ... I have asked at the pharmacy (drugstore) and they have it, but you need a prescription from a doctor to buy it :o:(

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Come to Poland please B)
you can buy absolut without prescription ;)
Andrzej

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From Estonia :)

Or you could contact a natural history museum and ask if they could donate a small amount for insect storage.

Using the various methylated spirits sold for alcohol stoves (T-sprit/T-r?d in swedish) is not recommended, but if you can not find anything else, go for it. Just dilute it with water to 70% before using. Try to find a brand with little/to colouring in it.

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When diluting the alcohol with water, it is best to use distilled water. It doesn't have to be pefectly distilled: water out of a dehumidifier or the ice that builds up in a freezer will do. But tapwater often gives a precipitate when mixed with alcohol, calcareous I guess.

John

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Thanks for the suggestions. Destilled water (for car batteries) I guess one can buy at the regular petrol filling station.

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Hi together,

sorry for necroing this thread.

jorgemotalmeida wrote: ..."NEVER use formaldehyde because it alters the structures of the specimens"...

Some Dipterists working with e.g. sciarids recommend the use of 1% formaldehyd-solution, because it saves structures by immediatly hardening the objects. This seems to be a contradiction.

At present I use this fluid in Malaise trap and everything looks quite good. The only disadvantage seems to be that everything gets very stiff.

What do you think about this?

Regards
Ralph

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crex, here in Portugal for 5 euros we can buy easily in supermarket, a bunch of them!!! I have 20 bottles of ethanol 70% in my house. :D lol

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Kahis, is it necessary really to mix distilled water with ehtanol 70%? I used to use only ethanol 70% for my specimens.

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Maybe in infinitesimal quantities it doesn't harm the specimens. But if you only use Formaldehyde it is sure that it will damage the structures of the specimens.

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Hi Jorge,

sorry, I must dig deeper. What exactly happens with what structures? I?m sure, that there is no problem with sciarids, but I also want to avoid serious damages at other diptera-groups:(.

My procedure is as follows:

>1% formaldehyd-solution in Malaise-trap for one week, additional some drops of "Mirasol", a surfactant from my darkroom.

>sieving through a tea strainer, careful rinsing with aqua demin.

>transfer in 70% Ethanol for long time storage

How would you do this with Malaise trap? Alcohol problematically evaporates (and accumulates water) during one week. What should I use better? ethyleneglycol?

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just a question.
right now i use alcohol (ethanol) of 87 grades. this mean is of 87%?
i have forget all chemistry i have learned ;)

thanks in advance,
cosmln

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"The Preservation of Natural History Specimens. I. Invertebrates. R. Wagstaff and J. H. Fidler. 1961."
I have found this to be a most authoratative text.
pp. 171-172: "Formalin - this is the most useful preservative. It is slightly acid and may damage specimens containing calcareous matter. it may be neutralized by being shaken with powdered chalk (5 gm/litre) and then filtered. Borax may also be used for the same purpose.
If formalin is neutralized the colours of specimens are generally better preserved than when in alcohol.
Use at 3-10% for storage.
As a preservative formalin is to be recommended for the more delicate soft-bodied animals as if used in the correct proportions it does not cause shrinkage"

I (TT) used to use formalin to 'fix' animal tissue prior to section cutting on a microtome. Tissue and cells were perfectly preserved; not sure what Jorge means by "it will damage the structures of the specimens"

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maybe, I must rewrite the sentence. I want to say: it can alter the shape of the specimen. Not necessarily damaging them.

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Interesting discussion:- lots of pro's & cons for wet and dry preservation.
A couple of points /tips to consider with alcohol preservation
1. If you kill a fly in Gaults solution or a dilute aqueous salt solution (even urine works well if you are really stuck for supplies out in the wilds!), all everscible structures evert, abdomens become fully expanded and the legs adopt an outstretched position. Just wash out the salts with water or low strength alcohol and then store or dry as per normal. I find that if subsequently dried properly, the material is much better to work with than the shrivelled distorted and sometimes mouldy material that has been impaled on a pin in the field
2. Some flies have a high oil content and get greasy on storage with setae matting etc (e.g. therevids seem to be prone to this). When drying from alcohol, you have to transfer toan apolat solvent like ethyly acetate to dry them out. Such solvents also remove the offending oils and the dry specimen never gets greasy. On the down side, when the oils have been removed, a specimen will be less strogly bound to a pin soit is probably better to card mount stuff dried from alcohol.
cheers
Adrian