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

[John E Simmons]

On Wed, Jul 2, 2014 at 11:36 AM, Watkins-Colwell, Gregory <
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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