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From: Lucy Helvenston <[log in to unmask]>
Reply-To:Scientific forum on fish and fisheries <[log in to unmask]>
Date:Mon, 3 Jan 2000 08:15:54 -0800

text/plain (182 lines)

Thanks for all the replies to my otolith preservation question. I've
received some very useful information which I hope I'll be able to put to
use if my project funding is extended! :)

Here's all the replies for those of you interested in otolith preservation
in ethanol.  Thanks again.


   I don't have an answer for you, but this is an issue I'm also very
interested in.  My preservation technique for larval fishes (6-20 mm SL) is
field preservation in 10% formalin for 1-4 days and then switching the
samples to 95% EtOH for long term storage.  I have done very few otoliths
so far, but I have not yet had any problems.  Since your technique is
undoubtedly more gentle than mine, and you are having some problems, I'm
concerned.  If someone suggests a better way to preserve fish for otolith
work, would you please forward the posting to me?  Thank you very much,

Michael Cooperman
Dept. Fisheries and Wildlife
Oregon State University
[log in to unmask]

Otolith preservation, especially for long-term storage, is problematic.
Because otoliths are composed of arangonite, they dissolve very quickly if
not well-preserved. My own work on leptocephalus otoliths ( 250 um in
diameter) indicated that at least 95% EtOH should always be used.  It is
best if the EtOH is buffered (I used Tris) to 7.0 pH.  Do not use calcium
carbonate as a buffer, as it dissolves at a lower pH than arangonite.  Also,
it is equally important to uses lots of preservative and/or change the
preservative at least a week after collection; earlier for stout bodied
fishes. After initial preservation, un-buffered, or weakly-buffered EtOH
quickly becomes acidic if not completely changed. Remember, it's not the pH
of the overall solution that matters, just the  pH of the preservative in
the labyrinth  of the inner-ear.

Formalin, isopropyl, and/ or denatured alcohol shouldn't be used unless the
otoliths are removed promptly, because of thier low initial pH.

By far the best method is fresh removal ,embedding in some sort of resin (I
used thermoplastic glue) and storing them dry.

Hope that helps a bit. I can give you a reference on otolith preservation if
you wish...

I think that pH is a big problem with some samples and not such a big
problem with others.  For larval fishes, acidic conditions will "eat away"
the otoliths, but this problem decreases with size of fish.  I use buffered
90-95% ETOH (I buffer with ground up marble chips) for larval and juvenile
fishes.  For adults or large-size juveniles 10%  formalin may be OK
initially, but I wouldn't store the fish in formalin too long.
 I think your pH changes reflect the destruction of the otoliths.  You had
acidic conditions intially, over time this was "buffered" by the otoliths
(or other bone).

I would be interested to hear what others say


Darran L. Crabtree
State University of New York
College of Environmental Science and Forestry
243 Illick Hall
One Forestry Drive
Syracuse, New York 13210
[log in to unmask]

I hope I can be of some assistance.  Ideally, you would want to remove the
otoliths as soon as possible without having to preserve the fish.  But because
this is not always practical, adult and juvenile fish can be frozen for later
analysis.  You can also preserve the fish in ethanol or formalin, but you can
run into problems with the otoliths dissolving because of the pH of the
sample.  Formalin can be used if it is buffered.  If you preserve otoliths in
ethanol, use 80% or greater and change the ethanol within 24 hour because it
becomes dilute from the water in the fish's tissues. You may also need to
buffer the ethanol.  If you do preserve otoliths in formalin or ethanol for
any length of time, the pH should be checked regularly.

I hope this is information is of some use.  If you would like some more
information, there is a very good chapter on otolith collection and
preservation in Otolith Microstructure and Analysis edited by D.K. Stevenson
and S.E. Campana (1992).

Andrew Munro

Montana State University
Biology Department
Bozeman, MT 59717

e-mail: [log in to unmask]

I, too had problems with preservation of fish larvae and otoliths and
learnt by
bitter experience.  I recommend 95% ethanol to preserve specimens.  Any lower
concentration and my experience is that the otoliths are either unreadable or
not there anymore!  If the otoliths are really important, then I attempt to
preserve in as high a concentration as possible and then pull the fish out and
preserve individually or extract otoliths as soon as possible.  My
experience is
that samples become acidic quite quickly and will dissolve larval otoliths
within a few days.

Cheers, Paul.
Sorry, I can't say for sure about the unbuffered 70% EtOH, but 95% did not
have the
same effect on mine.  I do know that I chose 95% because I read somewhere that
70% could cause problems.  Unfortunately that was over 2 years ago and I can't
remember the problem or the reference off the top of my head.  Perhaps the
additional water content causes a chalky coating (pure speculation)?

I have kept some of my juvenile yellow perch otoliths in 95% ethanol for
more than a year with no
deleterious effects.  They were already removed from the fish, but I'm not
sure that would make
a large difference.  The otoliths of individuals in which I injected a
formalin based preservative
into their body cavities and then froze to preserve gut contents did not
fare so well though ;-) .

What is the exact problem?  Are they just generally cloudy/dark or did they
a chalky coating?  I found that I had to actively clean my perch otoliths
very well (unlike
some other species I looked at), otherwise I would get tissue adhering to
them, and
obscuring an otherwise clear otolith.  As Secor, Dean, and Laban state in
Manual for Otolith Removal and Preparation for Microstructural Examination
"Fibrous tissue, composed of the macula and otolithic membrane, is commonly
observed adhering to otoliths".

Hope things turn out for the better, reading good ones is tricky enough
having bad ones to worry about.  Of course there are always bad ones ;-)

Alec Dale
Dept. Biol. Sci.
University of Windsor
Windsor, ON, Can
N9B 3P4
I have also had some problems with ethanol preservation of my silver hake
otoliths. I preserve them in 95% unbuffered ethanol. I think the problems
arise when the ethanol is not changed about 24 hours after preservation.
The water from the sample dilutes the ethanol and from what I have read
ethanol below about 80% can in fact be acidic to larval otoliths. I would
think that juvenile otoliths would be stronger and less susceptible to
pitting, etc. but perhaps this is your problem. I suggest you take a look
at the following document if you can:
Stevenson, D. K. and S. E. Campana.  1992. Otolith Microstructure
examination and analysis. Canadian Special Publication of Fisheries and
Aquatic Sciences 117.

Good luck. Sincerely, Jennifer Jeffrey
Jennifer Jeffrey
Oceanography Department
Dalhousie University
Halifax  NS  CANADA  B3H 4J1
Email: [log in to unmask]

Lucy Helvenston
Department of Biology, SDSU                                     SDSU Office Phone:  (619)594-0994
Southwest Fisheries Science Center, NMFS                        NMFS Office Phone:  (858)546-5645
8604 La Jolla Shores Dr.                                        NMFS Fax:           (858)546-5656
La Jolla, CA 92038                                              email:  [log in to unmask]

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