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

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require some assistance to dislodge. The upper and
undersides of rocks are very different environments; algal
films or turf usually cover the upper sides, while colonial
animals such as sponges, tunicates and bryozoans usually
live beneath the rocks along with their specialised
carnivores.
Sculptured rocks or large pieces of dead coral can be
scrubbed with a brush (e.g., 9 x 25 cm oval brushes or a
round brush about 5 cm in diameter are effective). When the
rocks are lifted out of the water, they can be brushed in a
bucket. The residue in the bucket, particularly from coral
washings, may harbour potentially dangerous animals so
care is required.
Rocks buried in sediment often have an anoxic,
blackish or rusty underside; certain molluscs occur almost
exclusively just at the oxic – anoxic border (e.g.,
phenacolepadids, some rissooideans, marine valvatoideans
and galeommatoideans).
Rock brushing while using SCUBA is best
accomplished within a cloth bag—a pillowcase is ideal—or
in a plastic laundry basket with a plankton net lining. The
rock is placed into the container and brushed within it from
above with the specimens mostly falling into the container—
although opinions differ as to how many specimens float off
rather than sink into the tub. Alternatively, the rocks can be
collected underwater and placed, with as little disturbance as
possible, into a large bucket or cloth bag attached to a buoyline.
The container can then be hauled slowly to the surface
by the boat crew and the rocks can then be scrubbed as
described above, minimising the risk of losing specimens.
Other methods
Hasegawa (2004) and Hickman and Porter (2007)
recently reported the collection of samples of Scissurellidae
using floating light traps. The use of attractants (light, bait)
may be worth exploring, particularly for micro scavengers
and predators.
Small grab samplers (e.g., Petite-Ponar, Wildco, NY,
USA: www.wildco.com), have been used for the collection
of micromolluscs (Geiger 2006a). At 14 kg weight it is
transportable as luggage on commercial airplanes and can be
deployed and recovered by hand from a small boat by a
single person. Sampling beyond normal SCUBA depth to
220 m has been achieved (B. Raines, pers. comm.) and,
unlike a dredge or benthic sledge, it requires line only as long
as the sampling depth, and recovers even the smallest species.
However, the sampling area is very small (15 x 15 cm).
Air-lift pumps can be used as a very effective way of
sampling both hard surfaces and substrate and are also a
means of obtaining, with careful and targeted use, large
quantities of living specimens (e.g., Bouchet et al. 2002).
Dredging and benthic sledge can provide significant
amounts of material and sample a larger area than either grab
or air-lift pump. The benthic sledge is advantageous as it
only skims the top surface where most micromolluscs are
found, but infaunal taxa will largely be missed.
Some taxa are commensals or parasites and their hosts
GEIGER ET AL. (2007) MOLLUSCAN RESEARCH, VOL. 27
need to be examined—for example Eulimidae on and in
echinoderms, pyramidellids on other molluscs, Epitoniidae
on Actinaria, Aeolidioidea and Solenogastres on Hydrozoa,
and Doridoidea on sponges and bryozoans.
Methods of collecting terrestrial micromolluscs include
sorting leaf litter and soil samples, beating foliage and
carefully examining specific habitats—bark, rocks, crevices,
logs etc.
Narcotisation and relaxation
The procedures and concentrations for narcotising
animals vary greatly, except for magnesium salts, where an
isotonic solution (7.5% in freshwater) must be used in order
not to disturb the osmotic balance of the animals. It is
recommended that as few narcotising agents as possible
(including water, cold and heat) be used and users should
aim to get to know them well.
There are two main reasons for narcotising animals:
• To facilitate and improve the yield of shake samples.
• To relax animals for detailed studies.
The first is somewhat simpler, as it is acceptable if the
animal retracts into the shell or curls up. An irritant such as a
small quantity of formalin, a small amount of detergent, or
some freshwater for marine and estuarine species can be
added to the sample. Many molluscs will retract into their
shells but remain alive. Limpets and chitons may not
necessarily fall off unless such a method is employed, but
non-shelled molluscs may be adversely affected. If
specimens are intended for histology, non-isosmotic
treatment is best avoided. A secondary shake in water with
the irritant after an initial shake in habitat water may produce
additional species in a sample. Note that byssally attached or
cemented bivalves (e.g., Mytilidae, oysters) and some
limpets can usually not be reliably collected other than by
physically removing them.
The whole sample may be pre-treated with magnesium
chloride to anaesthetise the animals (75 g MgCl 2 per 1 litre
of freshwater).
The algae may also be placed in a closed bag in full
sunlight so that heat stress will kill the animals, or for
tropical samples, cooling in the fridge or freezer will have
the same effect. In these cases, a single shake per subsample
will be sufficient to extract the vast majority of the
specimens.
Relaxation of animals for soft-part studies needs to be
more controlled and depends on the particular species in
question. The most common method is by gradual addition
of a 7.5% MgCl 2 solution in freshwater to the holding
container. Various molluscs respond differently to
magnesium chloride; some will hardly be affected while
others may immediately retract into the shell. Gradual
addition of the narcotic produces the most satisfactory
results. Introduce the solution away from the animal and
gently stir the water. Wait a few minutes and watch how the
animal reacts. Once the extended animal has stopped