Probably the commonest questions I get relate
to curation, i.e., how to collect, relax, photograph, preserve,
store, rehydrate, and ship medusae. It's not all that tricky,
just different from most other taxa due to the delicate
nature of the medusa's soft body.
HOW TO CITE THIS SITE:
I have put alot of time and effort into acquiring
the knowledge herein and making it freely available via this
web site, and it is scientifically and ethically appropriate
to be acknowledged for my contribution when this information
is used. The suggested citation for this work is:
Gershwin, L. 2002. Curating Medusae. Electronic
internet document available at http://www.medusozoa.com/curating.html.
Published by the author, web page established May 2002, last
updated
30 November, 2003
.
Collecting Medusae
NOTE: In most regions, collection of medusae requires
an official collecting permit. Contact your state Fish &
Game office for more information. The fines for collecting illegally
can be severe (e.g., up to $500 per specimen in California).
The most important part of collecting medusae
is to be gentle with them! In all cases, the ideal way to collect
medusae is, if possible, to dip them out of the water in a bowl,
bucket, cup, or whatever, such that the water supports the body.
The worst way to collect them is to lift them out of the water
such that their delicate epidermis is abraded, or that their
structures are forced to bear their own weight (except in the
case of most chirodropids, but please be careful, as some of
these can be deadly to humans!).
To dip medusae out of the water, do not try to
lift the bucket up under them -- being pelagic, their tentacles
and body will simply follow the current of water right out of
the bucket -- but rather, use the flow of the water to help
you get the specimen into the bucket. Position the animal at
the water surface, and position the bucket near the animal,
then dip the edge of the bucket into the water and the specimen
will ride the current into the bucket. Lift the bucket out of
the water as soon as the specimen has been captured, to avoid
losing it in the outflowing water of a full bucket.
One of the worst enemies of medusae is change
in water temperature. Cold-water species will disintegrate into
schmoo as their water warms, and warm-water species will grow
progressively sluggish as their water cools. Also, many medusae
are unable to stay up in the water column without current, and
will eventually die if left inactive for too long. Thus, it
is essential that you keep the water at the natural temperature,
and minimize the time the medusa spends out of its natural environment.
Most types of medusae cannot be left together
in a captive situation, as they are medusivorous (they eat other
medusae) or their stings are dangerous to other species. In
general, the larger scyphozoans such as Chrysaora and
Cyanea will eat just about any other medusa they can
wrap their lips around, even specimens larger than themselves.
Anthomedusae tend to be voracious predators on leptomedusae,
except Aequorea, which will eat medusae of any type that
are unlucky enough to be nearby. Rhizostomes do well with eachother,
but they are very active and will run into eachother endlessly
and quickly deplete the oxygen in the bucket. Cubozoans get
their tentacles irreversibly tangled. Cubozoans also seem to
paralyze rhizostomes, but this effect wears off after a half
hour or so of separation.
Relaxing & Preserving
Medusae
There are a variety of relaxation methods used
for invertebrates, and I have tried only a few. In my experience,
they tend to be taxon-specific, i.e., one method works better
for some species, while another method works better for other
species. They will be covered in the later paragraphs of this
section according to taxon. For preserving, I always use approximately
5% formalin in seawater, with the concentrated formalin added
into the seawater with the animal in it, rather than the animal
added to the solution of fixative (except with Cubozoa, see
below). I think there is less distortion of the specimen if
it only reacts to the formalin, instead of to the formalin plus
the stress of being transferred. For really humongus or beefy
medusae, I use closer to 10% formalin to fix the animal, then
later replace the solution with 5% for permanent storage. To
clarify the formalin-formaldehyde confusion: Formaldehyde is
a gas; formalin is a saturated solution of water and formaldehyde
gas (100% saturation occurs at 37% formaldehyde). Thus, concentrated
formalin (= "pure formalin", = 100% formalin) actually
contains only 37% formaldehyde.
For larger specimens (bigger than those examined
under a dissecting scope), Andre Morandini showed me the best
method: first set up a slow trickle of the relaxing solution
through airline tubing, so the specimen never realizes it is
being relaxed; after it is fully immobile, replace the slow
trickle solution with formalin, until 5% is reached.
For smaller specimens, I typically relax them
by slow addition of magnesium chloride or menthol into the petri
dish, then when the animal is no longer responsive when checked
microscopically to prods and pokes, then I begin adding concentrated
formalin drop by drop, maybe 1 drop per 15-30 minutes, to allow
time for diffusion across the dish.
Scyphozoa: I typically
use magnesium chloride for relaxation. I recommend Morandini's
slow drip method for both relaxation and fixation (see above).
Cubozoa: Either magnesium
chloride or menthol work equally well, though I now superstitiously
believe that menthol is actually superior (this did not come
easily to me -- Wolfgang Zeidler had to practically whack me
over the head). Note: if you attempt to relax more than one
at a time in the same container, they will hopelessly tangle
their tentacles. The key to preserving large, tropical cubos
was worked out by Phil Alderslade: "The problem with Chironex
is that they 'sting' the side of the container/bucket. This
causes the tentacles to adhere to the side, and become hopelessly
tangled in knots as the animal continues to swim about. Chironex
will stop swimming if put into cold water - so I chill the relaxant
before placing the jellyfish in it. The relaxant is isotonic
magnesium chloride. Stock solution is 30% magnesium chloride.
For use, this is diluted 1:3 with distilled water, which is
then diluted 1:1 with sea water and chilled. Pick up the live
Chironex, with tentacles hanging untangled, and lower it into
the cold relaxant. It will not swim about. For very large animals,
leave in the relaxant for several hours. Complete relaxation
of muscle tissue with magnesium chloride is virtually irreversible,
unlike menthol, so when the animal is then placed into the formalin
fixative the tentacles should not contract. The formalin does
not need to be cold." For smaller species, I have modified
Alderslade's method a bit, and it seems to work very well: chill
the formalin solution (5% formalin in seawater), and add the
specimen directly into the very cold formalin, not the formalin
to the specimen. I have fiddled with this a bit, and have been
able to manipulate the type of specimen I want. For the tentacles
very very relaxed and long, fully relax the animal, THEN pick
the animal up out of the relaxing solution and put it (tentacles
first) into the cold formalin. The idea is to instantly stun
the muscles at the moment of death. For some odd reason, no
matter how relaxed and unresponsive the specimen is, the tentacles
will still contract in room temperature fixation. If you want
the tentacles at a normal swimming length (not overly extended,
just normal), then don't relax the animal first, just drop it
straight into chilled formalin solution while it is still swimming.
Room temperature, non-relaxed fixation tends to produce pretty
distorted specimens which curl the pedalia and tentacles up
into the body cavity.
Hydrozoa:
Anthomedusae: Most tend to do best
when relaxed with menthol, then "topped off" with
mag chloride. No, I don't know why, I just know that it is so.
If mag chloride is used alone or first, some species crumple
unreversibly. And for curious reasons, methol is typically insufficient
-- the animal seems fully relaxed until the first molecules
of formalin touch it, then it crumples. I have tried a new method
with great success on a recent expedition: chilling the animal
in its seawater, then adding chilled formalin into the seawater;
if the specimens are relaxed in menthol while they are chilling,
the results are amazing! This works especially well for pandeids
and zancleids.
NOTE for Porpita:
These can be difficult, as they almost always lose their tentacles
during handling and definitely the moment they touch the first
molecule of formalin. Jane Fromont came up with a magical result:
flawless, intact, display-quality specimens, simply by very
very slow relaxation and fixation. I typically add liquified
mag chloride (you get this when it sits exposed to air too long)
to the petri dish, maybe 1 drop every 15-30 minutes, until the
specimen is completely non-responsive, maybe a total of 5 or
6 drops over a period of a couple of hours, then do the same
with concentrated formalin. The best results will be had from
a single specimen in a single standard size petri dish. They
will drop their medusae -- they look like little tiny stars.
For handling and transferring Porpita, they can be tumbled
and even poured without losing their tentacles, but if the tentacles
come in contact with an object from above or below (e.g., sand,
a spoon, your finger), they will stick and be easily shed. In
general, keep the animal floating and your specimen will be
flawless.
Leptomedusae: Most
respond equally well to menthol or mag chloride, and preserve
very well when first fully relaxed.
Limnomedusae: I haven't
experimented much with different techniques, but they seem to
do ok treated as for the leptomedusae.
Narcomedusae: For
reasons that are completely cryptic to me, mag chloride makes
some narcomedusae explode on contact. However, menthol works
fantastically! Fixation after relaxation produces stunning specimens.
Trachymedusae: Trachys
are tricky -- I just like how that sounds :-). Actually, I just
typically treat them as for leptos.
Preserving for DNA Analysis
You will have to make a choice, whether to have
a reference specimen or a DNA sample, as the two are not yet
complimentary. Presently, it is extremely difficult to get DNA
from formalin preserved medusae. And alcohol or freezing completely
ruins the specimen as a specimen. Of course, being radially
symmetrical, if you only have one specimen and you really really
need both, you can always cut a chunk out for DNA and preserve
the rest for identification. In general, I preserve first and
foremost in formalin, because I figure the DNA is useless if
I cannot identify the species. If I have ample specimens, I
store some in liquid nitrogen or a -80C freezer and some in
more-or-less 100% EtOH. I say more or less, because the critter
itself has a high water content that must be accounted for.
The trick with EtOH is to keep water from getting back to the
tissues, so you want the alcohol content to be pretty high.
The long term prognosis for alcohol specimens
is not good; they seem to degrade quickly, so extract the DNA
as soon as possible. Freezing actually seems to enhance extraction,
and there is no indication that long term freezing has negative
results (barring power loss).
An interesting paper by Dawson et al. (1998) compared
several storage methods, and came to prefer a DMSO-NaCl solution.
This has the added benefit of not making airlines and postal
services get nervous about the flammability and explosive properties
of alcohol.
I have also tried drying specimens, but I haven't
actually extracted yet, so I cannot tell how well this works.
In theory it should work well if the specimens dry quickly (i.e.,
without rotting).
Photographing Medusae
In situ photography and aquarium photography are
covered in detail by Wrobel and Mills (1998). However, most
of the images herein were taken under a dissecting microscope
while the specimens were still alive. It helps if they are relaxed
ahead of time to the point of immobility. I prefer the versatility
of a digital camera, but I believe a film camera will work just
as well if you have a photo tube. With many digitals, you can
literally hold the lens up to the ocular, and shoot away! The
most important part of photographing medusae (the more transparent
they are, the more important this is), is good, bright, focused
lights coming in through the sides of a clear dish. I prefer
the scientific type of "cool lights" with the two
flexible antennae. You can get a good, strong, low side light
if you fiddle with it. When the light is right, even the most
transparent of medusae will catch the light and reflect it back
for great photographs. If you have any vibration at all, use
a deeper dish.
In general, the most aesthetically pleasing angle
will be semi-oblique slightly from below if the medusa is flat-ish,
or straight on laterally if the medusa is tall-ish. If you plan
to identify it from the photographs, you will need (at the very
least), the shape of the mouth and stomach; the form and number
of the radial canals; the shape, location, and length of the
gonads; any exumbrellar structures or patterns such as nematocyst
warts or nematocyst tracks; "extra" structures such
as a peduncle, cnidophores, good up close shots of the tentacles
so that you can count the number per quadrant or octant as well
as discern the form of the tentacle bulbs or lateral extensions,
if present; and especially of the number, form, and relationships
of any other marginal organs (e.g., statocysts, ocelli, cirri,
marginal warts, rudimentary tentacles).
Storing Medusae
This is a bit disputatious at the moment, so I
will try to explain all sides and then give you my opinion,
which is based mostly on bad experiences and superstition. Most
historical specimens are stored in formalin, and many are in
absolutely dreadful shape. They sort of disintegrate with time.
But I think this is also somewhat dependant on body type, handling
and labeling, and the percentage of formalin. Bigger, heavier
medusae do better, full stop. But even some smaller ones from
the early 1900's are nearly flawless, if they were stored in
a jar inside a jar, so they weren't bouncing around with the
label and bubbles, and god knows what. Southcott's types from
the 50's and 60's are also flawless, but he tended to fix and
store in about 2% formalin -- probably a good idea. I habitually
tell people 5-10%, because that's what I was always told, but
I am coming to think this is too strong.
Kay Petersen told me that the Copenhagen Museum
switched all its specimens over to alcohol years ago, and I
must say that they have one of the best collections I have worked
with. The specimens are in fantastic condition. However, it
is really time consuming to do it right. As a general rule,
change the solution 10% per day toward 70% alcohol as a final
storage solution. The idea is to very very slowly replace the
water inside the tissues with alcohol. If you just transfer
the specimen directly, you may very well permanently ruin it.
Rehydrating Dry Specimens
This method was shared with me by Daphne Fautin.
It is impossible to fully restore the specimen to it's pre-dehydrated
form; the best you can hope for is that some of the characters
will be interpretable. Use Dawn liquid dishsoap -- yes, Dawn
really does work better -- no, I don't know why. Submerge the
specimen in a pretty concentrated solution of dishsoap and tap
water, and let it sit a week or two (or shorter or longer, depending
on what it needs). You can pinch it and see if it is getting
plump. Gradually decrease the soap and increase the tap water,
until the specimen doesn't seem to be retaining any more water.
Then represerve as normal.
Shipping Medusae
Hands down the superior method for shipping medusae
was shared with me by Paul Cornelius. The specimen must be in
a rigid container (to prevent it from being crushed or crumpled)
and there must be ABSOLUTELY no air bubbles. Even the smallest
air bubbles will allow for slosh, which will damage the specimen.
Think of it in as needing an amniotic sac of sorts to cushion
it from the traumas and dramas of transport. The best way to
get "all the air" out is to seal the container underwater.
Another method was detailed by van Impe (1992),
in which the specimen is sort of congealed into agar gel. I
actually think this is very very cool, but I have not tried
it. I would worry about whether it is staying completely hydrated.
Which reminds me, Paul Cornelius also suggested shipping larger
specimens "dry" (i.e., wrapped in damp towels) in
a rigid container. This is supposed to cut down on the weight
for shipping, as large specimens of course need a large amount
of water. With all due respect, this is a terrible idea, as
can be evidenced by some of the specimens on which he used this
method. Medusae need to stay wet, fully wet, and their "skin"
does not do well with even microscopic abrasions.
Some countries and carriers can be fiddly about
formalin (all are fiddly about alcohol). I have had great luck
with TEMPORARY storage in tap water for the duration of shipping
time. This seems to appease most airlines and postal services.
Note that they are also squeamish about seawater, and prefer
to have it replaced with tap water (this was explained to me
as having something to do with the corrosive properties of seawater
if it leaks into the corners of an airplane).
