Hi, Hugues Benoit and others,
I suggest you might try a marking technique that we have recently developed
Analyzing the growth and form of molluscs shell layers, in situ, by
cathodoluminescence microscopy and Raman spectroscopy Journal of Shellfish
Research 15: 659-666, 1996.
This paper discusses the use of manganese to mark the shell of abalone,
Haliotis rubra. The manganese mark in the shell, which is in the form of
manganese carbonate, is detected, in situ, by bombarding the area of
interest with electrons using cathodoluminescence spectrocopy. (This works
similar to a television tube)
Why I suggest our technique? The area of interest in our abalone is within
the spire or internal part of the shell where minimal growth occurs, as
compared to the outer growing margin. The amount of shell abalone lay down
over a few days is minimal. In some cases the marks are very very fine (<1
um)but very distinct, so much so they even map crystal growth of shell
layers. A side issue to your study is that manganese also visualises
microstructure by colour (see pap). Furthermore, repeat staining in the
order of two or three days has shown distinct marks, which may be suitable
for your otolith work.
The issue you need to consider is what concentration is toxic to your
flounder. We have found that 200 mg/L of MnCl2 in seawater bath is fine
for our abalone. However this may be quite different for vertebrates and
We have trialled flourochromes (Day et al., 1995 Mar. Freshw. res.,
46:599-605, but were less effective than manganese.
Post graduate studies
Abalone ageing and shell microstructure
University of Melbourne
At 04:23 PM 5/7/97 -0230, you wrote:
>I'm about to commence a study on the effects of growth on the timing
>(age/size) of early life history transitions (hatching/metamorphosis) in
>marine fishes. My"lab rat" is Yellowtail flounder, which will be reared
>from eggs to past metamorphosis, at different temperatures (2-12 degrees
>In a perfect world, during the course of this study, I would like to stain
>the otoliths of the larvae repeatedly at set intervals. This way, at the
>end of the study, I can approximate the size of larvae at previous time
>intervals. The only problem is that yellowtail flounder are relatively
>slow growers, and I predict that growth will range from 0.05 mm/day to 0.2
>mm/d. Litterature reports indicate that in cases of such slow growth, a
>large period of time (weeks to months) is required between subsequent
>stainings to maintain separate clear bands.
>My questions to the list are:
>1. What stains do you recommend using, which minimize mortality, show
>clear patterns, and which create narrow bands on otoliths (to reduce the
>amount of time between subsequent stainings)? Literature results are
>inconclusive for the more popular stains. Also, is immersion the best way
>of staining larval otolith? Might incorporating the stain in the food for
>the larvae create narrower bands?
>2. In your experience, is it feasible to repeatedly stain such slow
>growers? If yes, how often can this be done given that yellowtail hatch
>at about 2 mm and metamorphose at 14mm (growth rates of 0.05-0.2 mm/d)?
>I appreciate any advice that you could provide. Unpublished personal
>experience is often of great value, and in this case will help to set my
>experimental protocol. Please respond to me directly at the adress below.
>I will compile the responses for the list at a later date.
>Ocean Sciences Center
>Memorial University of Newfoundland
>St-John's, Newfoundland, Canada
>e-mail: [log in to unmask]