Relaxing and preserving
ascidians for taxonomy
Gretchen Lambert glambert@fullerton.edu
The ascidians should first be relaxed, then fixed in 10% seawater formalin
buffered with sodium borate to help preserve spicules
and color. (For molecular analysis and preservation of calcium carbonate spicules, a subsample should be
placed directly into 95% ethanol.) The formula for 1 liter of fixative is: 100
ml of full strength formaldehyde (37%), 850 ml of seawater, and 50 ml of
distilled water (or reverse osmosis water, or tap water if it does not have a
lot of minerals in it; sometimes I get a precipitate with tap water). To this
is added 1 gram of sodium borate. Mix thoroughly before use. The borate is not
very soluble so it takes a while to dissolve; thus I make the solution a few
hours or even a day ahead of when I want to use it. It is necessary to use the
50 ml of distilled water instead of all sea water so that the solution will not
be hypertonic. Ethanol is definitely NOT a good preservative for taxonomy, and
ascidians should never be placed directly into ethanol (except a subsample as described above). Ethanol makes the tissues
opaque and brittle, and it removes all color.
An easy way to relax ascidians is with
menthol crystals. You can carry a small vial of crystals in the field with you,
place a few crystals in a Ziploc bag with the sample in sea water and seal
tightly. By the time you get back to the lab the animals will usually be at
least partially relaxed. If not, keep them in the bags for a bit longer, or
proceed as follows.
A second method for relaxing ascidians uses
menthol in ethanol, a technique I learned from Don Abbott. Fill a small bottle
(10-20 ml or so) with crystals of menthol. Then fill the bottle with 95%
ethanol and shake to dissolve the menthol, which is much more soluble in
ethanol than in water. Place the ascidian in a dish of seawater (or the
menthol/seawater from your Ziploc bag). Then add about 5 drops of the
menthol/ethanol and QUICKLY cover the dish tightly (I use a small sheet of
glass or plastic with a weight on top) to prevent evaporation of the menthol.
Every 10 minutes or so add another few drops of the menthol/ethanol until when
you insert a sharp probe into an open siphon there is absolutely no response.
(Use a hand lens or microscope to be sure about this!) Relaxation may be
achieved in as little as 10-15 minutes or so for some species, but may take
several hours for others. Fill another dish with fresh seawater, have a jar of
the seawater/formalin fixative ready. Lift off the crystallized menthol that
will be floating on the water surface (I put it on a paper, dry it and put it
back into the bottle for re-use). Transfer the relaxed specimens to the dish of
fresh seawater, rinse briefly to remove the extra menthol crystals (you may
have to do this twice) and then quickly transfer to the jar of fixative and
cover. If it's a large solitary, hold it upside down to let the seawater drain
out of the open siphons before immersing it into the fixative with the siphons
pointing upward so that the animal will quickly fill with the fixative.
If you won't be returning to the lab, take
the formalin, bottles, etc in the field with you. Leave the samples in the ziploc bags with menthol for several hours, then rinse in
tap water or blot on paper towels and transfer to formalin. (You can of course
fix the ascidians directly in the 10% seawater formalin without relaxation but
it makes identification especially of colonial species very difficult.)
Ethanol
fixation:
Before placing a specimen into 95% ethanol, if
you can, rinse in tap or distilled water to remove as much seawater as
possible, because the seawater will cause precipitates to form in the ethanol. At
the very least, blot on paper towels before placing in the ethanol. Do this in
the field with fresh specimens. Don’t use colonies that have been in menthol
for several hours and may be half dead by the time you get to the lab. With
colonial species, cut into small pieces before placing in the ethanol. With
solitary species, remove from the tunic, discard the tunic, and blot thoroughly
(or rinse and then blot) the body before placing in ethanol, to remove as much
liquid as possible. You can also dissect out the gonads and preserve only the
gonads in ethanol. Store in a freezer as soon as you can.
Opinions vary as to how to store ascidians
long-term for museum vouchers. I and the Monniots in
I agree with Patricia Kott
that "no power on earth will maintain the living colour
of ascidians after they are collected- so nothing can replace: (1) colour notes on living specimens before they are removed
from the substrate; and/or (2) in situ photographs." To this I would add notes on the general
appearance of the living animal or colony. For example, the atrial languets of
many aplousobranchs are highly contractile, and their
short stubby appearance in preserved zooids bears little resemblance to their
appearance in living zooids. Larval morphology is of great importance in the
identification of colonial forms; care should be taken to preserve brooded
embryos and any swimming tadpoles.