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

[A.J.van_Dam at lumc.nl]

Just to add to Rob's comments, of course change of fluid should be decided on pH monitoring, not on the collection example I mentioned where indeed several variables can be of influence (temp./specimen/container/seal/buffer molarity etc.). Still I would recommend when acidity has significantly lowered (more than one pH, coming close to pH 5) to change the fluid.

Regards,

Dries.

Andries J. van Dam, conservator

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________________________________
Van: Robert Waller [rw at protectheritage.com]
Verzonden: donderdag 3 juli 2014 9:05
Aan: Dam, A.J. van (DOO); simmons.johne at gmail.com; gregory.watkins-colwell at yale.edu
CC: cahaas at vt.edu; nhcoll-l at mailman.yale.edu
Onderwerp: RE: [Nhcoll-l] long-term storage of amphibian larvae in formalin

Just to clarify, formalin is stable with respect to spontaneous decomposition but not stable with respect to oxidation.  A buffer system, by definition, will slow but not eliminate drift in pH levels. Different containers and closures will lead to different rates of pH decline. I would hesitate to adopt a fluid change schedule based on a different collection unless that collection uses the same containers and the same buffer formulation.
Robert Waller

From: nhcoll-l-bounces at mailman.yale.edu [mailto:nhcoll-l-bounces at mailman.yale.edu] On Behalf Of A.J.van_Dam at lumc.nl
Sent: July-03-14 6:46 AM
To: simmons.johne at gmail.com; gregory.watkins-colwell at yale.edu
Cc: cahaas at vt.edu; nhcoll-l at mailman.yale.edu
Subject: Re: [Nhcoll-l] long-term storage of amphibian larvae in formalin

Formalin is not stable! We noticed in our fetus collection stored on phosphate buffered formalin (pH 7.3) that after ten years pH is around 6, after 20 years around 5, and after 30 years around 4 (close to unbuffered formalin). Therefore, when using buffered formalin as a preservation fluid, I recommend to change the fluid every 10 years.

Regards,

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<mailto: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/>

________________________________
Van: nhcoll-l-bounces at mailman.yale.edu<mailto:nhcoll-l-bounces at mailman.yale.edu> [nhcoll-l-bounces at mailman.yale.edu] namens John E Simmons [simmons.johne at gmail.com]
Verzonden: donderdag 3 juli 2014 5:50
Aan: Watkins-Colwell, Gregory
CC: Carola Haas; nhcoll-l at mailman.yale.edu<mailto:nhcoll-l at mailman.yale.edu>
Onderwerp: Re: [Nhcoll-l] long-term storage of amphibian larvae in formalin
On Wed, Jul 2, 2014 at 11:36 AM, Watkins-Colwell, Gregory <gregory.watkins-colwell at yale.edu<mailto:gregory.watkins-colwell at yale.edu>> wrote:
...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.
If buffer the solution with 4 g monohydrated acid sodium phosphate + 6.5 g anhydrous disodium phosphate per liter of one part commercial formaldehyde with nine parts deionized or distilled water, that should be a very stable buffered system. The sources of error that can produce clearing include failure to rinse out field buffers, using tap water to dilute the formaldehyde, and not measuring carefully. You can purchase pre-buffered formaldehyde but personally, I would not trust it for use with scientific specimens. I have never seen clearing when this buffer system is used properly.

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.

If the formaldehyde is properly buffered, it will not damage reptile eggs. However, I would not bother fixing reptile eggs in formaldehyde unless it is necessary in the field. Better to preserve them directly in 70% ethyl alcohol.
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.
Because formaldehyde solutions are essentially water ("10% buffered formaldehyde" is really about 96% water), amphibian larvae in 1:9 formaldehyde and water solutions can be safely transferred to deionized or distilled water when people use them, then returned to the buffered formaldehyde for storage. Unless you rinse the specimens very thoroughly when you transfer them to alcohol, you are still going to have trace amounts of formaldehyde in the alcohol solution that can pose a safety issue. Wear neoprene (nitrile) gloves when handling specimens in any case.

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.

I agree with you here, Greg. No solution is perfect for everyone, and documentation of what is done to specimens is critical. However, you can greatly reduce risks to specimens and to workers by controlling which chemicals are used, how they are mixed, and how they are used with a set of enforced SOPs (Standard Operating Procedures).
--John
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