[Nhcoll-l] long-term storage of amphibian larvae in formalin

[A.J.van_Dam at lumc.nl]

DMDMH (trade names Dekafald/Glydant) could just be the best of both worlds (formalin-acidification versus ethanol-shrinkage). It is a stable aldehyde compound with neutral pH (6.5-7.5) in aqueous solutions and seem to have less impact on degrading DNA. It is sure worth giving it a try. Preferred concentration: 5-10% of saturated stock solution (55%) with addition of 5-10% glycerol.

Regards,

Dries

Andries J. van Dam, conservator

Museum of Anatomy
Leiden University Medical Center
Building 3 (V3-32)
P.O.Box 9600
2300 RC Leiden
The Netherlands
tel: +31 (0)71 52 68356
E-mail: A.J.van_Dam at lumc.nl
Visiting address: Hippocratespad 21
Scientific associate Natural History Museum London
http://www.nhm.ac.uk<http://www.nhm.ac.uk/>
Directory Board member ICOM-CC
http://www.icom-cc.org<http://www.icom-cc.org/>

Director Alcomon Company
http://www.alcomon.com<http://www.alcomon.com/>

<http://www.alcomon.com/>
________________________________
Van: nhcoll-l-bounces at mailman.yale.edu [nhcoll-l-bounces at mailman.yale.edu] namens Watkins-Colwell, Gregory [gregory.watkins-colwell at yale.edu]
Verzonden: woensdag 2 juli 2014 17:36
Aan: Carola Haas; nhcoll-l at mailman.yale.edu
Onderwerp: Re: [Nhcoll-l] long-term storage of amphibian larvae in formalin

A lot of amphibian larvae are housed in 10% buffered formalin long-term because it helps prevent the softer body parts from shrinking in ethanol.  I have, however, found it difficult to maintain the pH properly and even the best of formalin solutions can result in some specimen clearing long-term.

I do not keep reptile eggs in formalin.  I fix them in formalin and then transfer them to ethanol.  Long-term exposure to formalin can damage the eggshell and cause issues with histology.  I treat reptile eggs as I would a whole reptile specimen and transfer them to 70% ethanol for long-term storage.

Amphibian eggs and egg masses stay in 10% buffered formalin because they do shrink in ethanol to the point that they are essentially useless.

But, as for amphibian larvae, we’ve started transferring them to 70% ethanol for long-term storage.  This also makes them easier to work with from a health and safety perspective, especially with a lot of student workers.  Honestly the vast majority of our amphibian larvae were field preserved in 70% ethanol to begin with and never had formalin used.  The added bonus is that DNA can be more easily extracted from them, but the down-side is that the more delicate features are difficult to discern.

I think that no matter what you do, there will be a cost/benefit.  Understand your institutional priorities and weigh those against the rarity of the specimen. This also might be a good reason to photograph examples of each taxon/developmental stage PRIOR to changing storage fluid.  Even if you only change to new formalin, you should document.  There’s a chance that your formalin isn’t buffered the same way as what was used in the past.  That difference can cause some issues with the specimens.  So, really, whatever you do there is a risk.

Good luck

Greg


--------------------------------------
Gregory J. Watkins-Colwell
Collection Manager, Herpetology and Ichthyology
Division of Vertebrate Zoology
Yale Peabody Museum of Natural History
170 Whitney Avenue, Box 208118
New Haven, CT  06520
203/432-3791  or    fax: 203/432-9277
-----------------------------------

From: nhcoll-l-bounces at mailman.yale.edu [mailto:nhcoll-l-bounces at mailman.yale.edu] On Behalf Of Carola Haas
Sent: Thursday, June 19, 2014 3:44 PM
To: nhcoll-l at mailman.yale.edu
Subject: [Nhcoll-l] long-term storage of amphibian larvae in formalin

I received such great help for my previous request, and hope folks won't mind my sending another one so soon.  (I'm just a field biologist who has been tasked with a cleanout and reorganization of our collection.)

We have a number of larval amphibians (tadpoles and salamander larvae) preserved in 10% buffered formalin.  Most of our fish, amphibian, and reptile specimens were fixed in formalin but then transferred over to ethanol for long-term storage. I have read that formalin is more appropriate for long-term storage of reptile eggs and larval amphibians, but I wanted to check and make sure that is still the current practice?

I would like to improve the safety of our collections by switching to ethanol if that is acceptable, but obviously not if it will degrade the specimens.

Thank you!

Carola A. Haas
Professor, Wildlife Ecology
Dept. of Fish & Wildlife Conservation
112 Cheatham Hall
MC 0321 Virginia Tech
Blacksburg, VA 24061
cahaas at vt.edu<mailto:cahaas at vt.edu>
540-231-9269
http://www.fishwild.vt.edu/faculty/haas.htm




-------------- next part --------------
An HTML attachment was scrubbed...
URL: http://mailman.yale.edu/pipermail/nhcoll-l/attachments/20140702/75c28e71/attachment.html