Preservation techniques for benthic invertebrate samples

[Buz Wilson_Australian Museum]

At 13:31 13/4/95 -0800, Fraser R. Sime wrote:
>The Northern District of the Calif. Dept. of Water Resources has an
>extensive benthic invertebrate sampling program in the north state. I am
>heavily involved in the coordination of this program. For many years, we
>have used formaldehyde as our main preservative for benthic samples
>collected in the field. These are archived, sometimes for many months,
>before being id'd and enumerated, etc..
>
>Recent changes in safety regulations, disposal problems and concerns for
>the environmental health of workers has forced us to look at other
>alternatives for preserving our benthic samples. We do use ethanol for
>short-term work, but have found it to be less than ideal over extended
>time.

Long experience with both shallow water and deep-sea benthic programmes has
demonstrated that to obtain easily identifiable invertebrates also useful
for taxonomy, one must fix the samples with buffered seawater/formaldehye.
We, however, do not leave the specimens in the fixative for long, usually
12-24 hours is sufficient.  Then the samples should be washed with fresh
water to remove the salts, and put into 70-80% ethanol (some folks use the
cheaper isopropanol, but it is also more poisonous). I draw a distinction
between fixation (cross linking  proteins to "toughen up" the specimens) and
preservation (prevention of little buggers from eating your sample).  Some
types of taxonomy (i.e. that practised by meiofauna workers, particularly
nematode specialists) require that the specimens must not be put into
ethanol, as this will degrade their surface. Moreover, if you want accurate
wet weights, ethanol will leach some of the organic material from the
specimens. In these cases, placing the material in a reduced concentration
of buffered formaldehyde is sufficient for preservation, but it is essential
to insure that the sample does not go acidic (a problem with long storage of
formaldehyde solutions). I do not recommend putting marine benthic samples
directly into ethanol because this will cause an insoluable precipitate in
the sample and render many specimens difficult to identify. Freezing is ok
as long as you are not interested in identifying anything, i.e. such as
doing biomass and C/N ratios etc (rather uninteresting in my book - but
then, I'm a taxonomist). Most soft invertebrates turn into unidentifiable
mush after freezing and thawing.

A great deal of hysteria surrounds the use of formaldehyde. Nevertheless, I
regard it as a useful tool, albeit a potentially damaging one like many of
the tools that we use.  We can no longer use it in a cavalier fashion, and
one should *never* breathe the vapours or get formalin on one's skin. If
your sampling occurs during protracted periods at sea, you can wash the
samples into ethanol on board, discarding the waste formaldehyde over the
side. Protective masks can be obtained that will effectively remove any
formaldehyde from inspired air. If the washing occurs on land, then measures
for the safe disposal of the waste solutions must be taken. Spend the money
and use formaldehyde safely - it gives good results. If you're interested,
contact me directly and I can dig out a reference or two on handling benthic
samples.

Cheers,

Buz Wilson
Australian Museum