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

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8 after it has been immersed in corrosive chemicals. Needles, pins or razor blade fragments can be glued or otherwise attached to tooth picks or other wooden sticks. Heated needles can be pushed into thin wooden strips or perspex/ plexiglass rods, but the heating makes the metal more sensitive to corrosion. Surgical needle holders are useful for holding small needles, pieces of a razor blade (or anything else thin or flat). There are also special ‘blade holders’ available for this purpose. Pins and needles Needles are probably the most important piece of equipment for radular and many other micromollusc applications. The finest and most expensive needles are made from electrolytically etched tungsten wire (USD/Euro 5–10 each) with a 1 µm point (Fig. 4C). These can also be made using tungsten wire and suitable equipment (Hubel 1957). Their points can easily be deformed, which is sometimes an advantage for certain types of manipulation. Micro-pins used to pin small insects come in black or stainless steel and in diameters from 0.1 to 0.2 mm. The stainless steel needles are less prone to rust and the thicker ones are stiffer but less pointed, with much variation in the shape and quality of the point. The black steel pins are sensitive to rust (Fig. 4B) although their life is prolonged by rinsing and drying after use. Household pins are much blunter and thicker but are often made of chromium plated brass and are less sensitive to chemicals. Sewing needles are made of chromium plated steel, available in many sizes and can be useful for work on larger specimens (Fig. 4A). Most of the ready-made needles for surgical use are inferior to micro-pins and much more expensive. All metal needles or pins can be bent by holding the very tip with a pair of watchmaker’s forceps and bending the outermost fraction of a mm to a suitable angle. This tool is useful for moving radulae from one rinse to the next (see below). Such needles, including those with a minute hooklike end, are also valuable for dissecting. A needle with its point bent at 45° is excellent for picking small single valves of small bivalves; turn the shell so the concavity is up and ‘hook’ the valve under the hinge. Needles with a 60° bent point are good for pulling the animal out of coiled shells. The needle point is inserted along the wall of the shell, then the point rotated to hook the animal. Hairs For cleaning dust particles from mounted radulae it is often better to use hairs (rather than a pin) glued to a small handle of wood or inserted into a holder. Eyebrow hairs (if straight) and eyelashes are commonly used but some animal hairs such as the pointed and stiff whiskers of a cat make very good tools. The stiffness of hairs decreases with increasing length of the hair. Brushes One centimetre wide brushes are useful for manipulating dry samples while the finest brushes can be GEIGER ET AL. (2007) MOLLUSCAN RESEARCH, VOL. 27 used for manipulating individual specimens. The diameters of brushes are graded, with ‘0000’ being the finest, and may be made of either synthetic fibre or natural hair. Synthetic fibre is chemically more resistant, can be used with bleach and hydroxides used in radular extraction and are usually a little stiffer. Natural hairs are often better for picking up small shells but are more expensive. It may be necessary to shape the tip, or to increase the stiffness of the brush by shortening the hairs. Many fine paintbrushes have one or a few much thicker and stiffer hairs to act as a support for the others. These can be cut off to avoid the risk of specimen damage if the brush is used for cleaning. Pliers The smallest sizes of regular tool pliers are useful for cracking and opening small shells. Locking vice grip pliers will prevent the specimens being crushed. For most microshells, dissolving the shell is a better method (see below). Wire cutters for electronic use come with a variety of cutting edges, pointed, blunt, straight, angled, etc. and some are made of stainless steel. These are good for opening medium-sized (>5 mm) shells from the aperture, by breaking the outer lip. For cutting steel needles, use wire cutters with tungsten carbide edges. Watchmakers forceps can be used for cracking very small shells (see below for details). Drills Some power tools resembling a dentist’s drill have a flex-shaft attachment and a variety of rotary tool bits, engraving cutters and high-speed cutters that can be used to open shells with minimal damage. Drill bits are available down to about 0.7 mm diameter and can be used for grinding a hole of >0.7 mm diameter in the back of the shell. A hole can be made in thin-shelled species simply by scratching the shell with a needle. Tool sharpening A few different types of very small files for jewellery work are useful, both for keeping other tools in shape and for filing holes in 3–5 mm (or larger) shells. Files rust easily so, if in contact with seawater, they need to be rinsed with hot fresh-water and wiped dry. For the rough shaping of coarser tools, a bench grinder with as fine a wheel as possible can be used. For more detailed work use fine sand- or carborundum paper or sharpening stones. For the final sharpening of forceps and needles, use a fine-grained stone such as ‘Arkansas Stone’ (see also http://www.antiquetools.com/sharp/sharphistory. html). With some practice, a good point can be achieved for watchmakers and some other forceps. Start with (if necessary) roughly bending the tips so they are parallel, then grind them off to the same length and start sharpening on a fine carborundum stone. The final grinding is done on an Arkansas stone, by moving it back and forth in the direction of the points. The finishing should be done under a stereomicroscope for better control.
TECHNIQUES FOR STUDYING SMALL MOLLUSCAN SPECIMENS 9 Bowls There are two types of bowls that we find particularly useful for working on microscopic animals. Square, solid glass bowls (‘embryo-bowls’), ca 40 x 40 x 15–17 mm are ideal. Use a square piece of glass cut to the same size as a lid. The sides of the ‘bowl’ slant at an angle, so nothing is concealed by a meniscus. They are easy to handle since the outer sides are straight and allow a good grip, as opposed to watch glasses. The lid usually stops evaporation, but some can have irregularities that prevent a tight closure. Depression slides (concavity slides) are available with different sized depressions. Those with a depression about 18 mm diameter and 2.5 mm depth are good for cleaning smaller radulae. As a lid, another depression slide with a larger diameter depression can be used upside down. If a regular flat glass slide or coverslip is used as a lid when heating KOH in radular preparation, condensation will form on the glass of the lid above the fluid, which will finally connect with the fluid in the depression and may draw it and the radula into the capillary space between the slides. The larger ‘dome’ above solves this problem. Larger dishes useful for sorting are discussed below. Collecting Field collection of micromolluscs requires some specialised techniques. Most micromolluscs are difficult to see with the unaided eye and usually cannot be identified in the field without magnification, making targeted collecting for a particular species difficult. Thus the likely habitats of the target organisms, or a range of microhabitats in the case of surveys, usually need to be sampled. The collecting methods covered below are simple techniques that require minimal equipment and can be undertaken by hand in intertidal and other shallow-water aquatic systems. The process of obtaining small specimens is not limited to intertidal and SCUBA as the same or similar methods can be used on a larger scale as, for example, with samples collected by various remote-sampling devices such as dredges, trawls, epibenthic sledges or grabs. In such cases, equipment (such as sieves and containers) needs to be scaled-up. In order to obtain a representative collection, a range of techniques should be employed. The choice of the final volume of the sample, and the number of samples, depends on the question to be answered. Small samples of 50–100 ml will reveal the dominant species whereas samples of 10 litres and more may still miss rare species (rarefaction effect). All collecting, domestic or foreign, should be conducted under appropriate and applicable permits. However, because microscopic species cannot usually be collected in a targeted fashion and substrate sampling is essential, this needs to be appropriately covered. Some authorities do not provide the option for substrate collection and require a priori list of species and numbers of specimens to be collected. These issues are best dealt with on a case by case basis. Hand collecting methods Shell grit. Sediments are usually distributed nonuniformly. Some areas accumulate organic material and biogenic carbonates. These shell-rich portions of the sediment are often referred to as shell grit or shell sand. They can simply be scooped up and processed like any other sediment samples. They usually have mainly empty shells and sometimes can be the only source of certain species. The fine sand can be removed from these samples in situ by washing it in a sieve or moderately fine mesh bag (Fig. 4F). While the bag method works well for the larger micromolluscs, it will lose many of the smaller species. Algal samples. Algae are a habitat of many molluscs, those with large fronds usually having fewer (but often different) individuals and taxa than the heavily branched or foliose species, such as many of the turfing algae. Kelp holdfasts can also harbour different species. Certain molluscs are found only on particular algae; for instance, sacoglossans are generally found on green algae (Chlorophyta). In (ant-) arctic waters, larger algal species harbour many micromolluscs, particularly in holdfasts. Large algae such as Laminaria sp. may lose their blades seasonally, thus at most a single season of micromolluscs can be encountered on the blade, while the holdfast may contain several seasons’ worth of fauna. Algae can be processed on site or collected for later processing; larger algal species can be placed in a bucket, smaller species may fit into a ‘zip-lock’ bag. The most durable freezer zip lock bags with slider closure mechanism are also suitable for SCUBA collecting. One litre of algal volume for turfing species usually produces a representative sample. The method of extraction of the molluscs from algal samples removed from the habitat depends in part on the intended use of the specimens. They can be extracted alive (using a binocular microscope) for observations on living material or to extract specimens for special fixation. For bulk collection, the simplest technique is to vigorously wash algae on the shore in a bucket or bowl filled with seawater. The algal material is then removed and the sample allowed to settle briefly. The water can then be gently decanted, being run through a sieve to catch any floating molluscs (e.g., opisthobranchs). Such samples are ideal for collecting living specimens for later examination. More thorough washing can be carried out by vigorously shaking algae in a 0.5–1 litre jar half filled with water from the environment and with a tightly fitting lid. Samples may also be pre-treated with an irritant or narcotic (see below) to ensure that tenacious specimens are released from their substrate. Leaves and other litter. Rich organic material provides molluscan habitats particularly in mangrove and other upper littoral and supralittoral habitats as well as terrestrial habitats. Mangrove litter can be washed as for algae and is ideal for collecting e.g., small ellobiids, truncatellids and assimineids. Rock washing. Smooth rocks may be hand-washed with bare hands in a bucket. Byssally attached bivalves, limpets, chitons and opisthobranch may be tenacious and
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