Molluscan Research: Techniques for collecting, handling, preparing ...

24
pictures can still be obtained. Then cut the body, particularly
as much of the columellar muscle as possible, using a needle.
The lower body can then be pushed out either directly, or
indirectly by inserting small pieces of wet tissue paper or
cotton wool through the hole with fine forceps or a needle.
Some of the visceral coil will be probably be left in the shell,
but the head-foot is usually rather easily removed.
Species with a very tightly coiled shell may be difficult
to process without severe damage to the shell. Such
specimens can be broken in two at mid-height, soaked and
the soft parts flushed out by inserting the apical part of the
lower half into a fine pipette. Let the water drain into a fine
mesh that can be examined under the stereo-microscope if
the body is fragmented. Sometimes the radula can be
obtained even from almost completely decayed and
fragmented remains of a poorly preserved or rotten
specimen. The two remaining pieces of the shell can usually
be glued together. Dilute the glue if it dries too fast.
The easiest way to manipulate the specimens is to hold
the specimen between index finger and thumb of your hand
with less dexterity (usually left) and moisten the specimens
and the fingertips with the immersing fluid using a fine
brush; use gloves if necessary (or work with harmless
chemicals) and a stereo-microscope as needed. With your
right hand, apply the appropriate tools (pins, forceps) to
remove the body. This technique is generally superior to
manipulating the specimen fully immersed in a suitable dish
with two instruments (needles, brush, forceps). Attempts to
construct specimen cradles with pins in a wax tray are
disappointing. The finger-method works with specimens
down to less than 1 mm in size.
Opercula
Many microgastropods produce opercula of a variety of
forms and sturdiness: from strong calcified ones to wafer
thin varieties. It is usually still attached to the foot and may
be retracted into the aperture of the shell. To avoid damage to
the operculum when extracting the body, try removing the
operculum by inserting under it either a micro-scalpel, a pair
of fine watchmakers forceps, or a fine needle. If the
operculum falls off at the first touch, the specimen is
probably more or less decayed; there may be little detail
available in the soft parts and the radula may need extra care.
During the process of extraction, hold the shell under the
microscope between thumb and index finger (as described
above). Opercula are easily imaged by SEM when still in
position in the aperture if the animal is not too far retracted
and very thin opercula are often better imaged in this way.
Contrast of structural details such as growth rings may be
indistinct when mounting thin corneous opercula on doublesided
carbon adhesives. If separate mounting of thin opercula
is desirable, mount them only with part of the operculum
touching the carbon adhesive, or place the moist operculum
on dry PVA glue.
Removing the shell the fast way
As an alternative to the above methods, the shell can be
decalcified in dilute hydrochloric acid (HCl: 2–5%), which
GEIGER ET AL. (2007) MOLLUSCAN RESEARCH, VOL. 27
should only take a few minutes. However, some bubbles will
form and the procedure may rupture the tissues when there
are internal deposits of carbonates. An alcoholic solution of
HCl is less damaging to the soft parts as less carbon dioxide
is generated and the bubbles are smaller due to the lower
surface tension of the ethanol. Decalcification with ethylene
diamino-tetraacetic acid (EDTA) is possible in aqueous
solutions (5–20%, pH 7.0 adjusted with 1N NaOH), and is
favoured by some for animal preparation for histology (not
further covered here), but is very slow. A mixture of formic
or acetic acid and formalin can be used to fix and decalcify in
a single step (as in Bouin’s fluid). Cracking the shell is
another simple option to get access to the animal (see section
Storage above).
Once the shell is removed from the body, several
options are available: investigation of external morphology
by light microscopy or SEM (see Tissue preparation for SEM
below); investigation of internal anatomy by dissection or
histology or radula extraction.
Tissue preparation for SEM
Once the animal has been removed from the shell, it
must be dried prior to further inspection using the SEM. In
some instances, it is advisable to dry the body inside the shell
and examine the exposed head-foot characters visible on the
relaxed and nicely extended animal. Drying of tissue from
aqueous or alcoholic solutions directly leads to severe tissue
shrinkage and makes detailed inspection of the external
morphology impossible.
Proper drying of animals can be carried out by three
methods: critical point drying (CPD: Fig. 9);
hexamethyldisilizane (HMDS); and freeze drying (Sasaki
1998). CPD and freeze drying require specialised equipment.
HMDS, on the other hand, can be used at room temperature,
although a fume hood is necessary for safe handling of the
liquid. Both CPD and HMDS often give suitable results
although there are sometimes unexplainable failures. In both
cases, the specimen has to be taken through a graded ethanol
series to pure, undiluted electron microscopy grade ethanol.
‘Pure’ ethanol used for storage of specimens is actually only
approximately 95% and is unsuitable for tissue dehydration,
and most problems arise due to insufficient dehydration.
From pure ethanol, the ethanol has to be replaced by either
CO 2 in the case of CPD, or HMDS, through several fluid
changes. For HMDS, better results are obtained if the liquid
is evaporated more slowly in a covered dish overnight, as
opposed to an open one in a few minutes. For freeze drying,
the specimen is placed in t-butyl alcohol and the freeze
drying machine automatically applies a vacuum to the cooled
specimen vessel. The advantage over CPD is that typically
the equipment is all automatic, not requiring the operator to
fill and empty the specimen reservoir with liquid CO 2 . There
are some semiautomatic CPD. The results of CPD and freeze
drying are comparable (see also Goldstein et al. 1992:
chapter 12.5.4, fig. 12.9). Specimens from historical
collections are often suitable for SEM tissue preparation
(Fig. 9).