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

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14 engage in effective damage control. Proper initial specimen preparation can avoid many problems later on. Initial drying Prior to dry storage, all specimens should first be washed in fresh water to remove dirt, soften mucus and dissolve salts, which are hardened and/or precipitated by ethanol. It is very important to remove any salt because NaCl 2 is hydroscopic. To remove excess water, to dehydrate animal remains within the shells and to speed up drying, the material may additionally be washed with 80–100% ethanol. If the dry specimens or total samples were not previously washed, soak the material thoroughly so that mucous, dirt and salt crystals dissolve. Specimens can then be air dried on absorbent paper. Insufficient drying will negatively affect long-term storage of the specimens. Specimens washed with 100% ethanol dry fastest and with the least possibility of retaining any liquid. In diluted ethanol, the ethanol will evaporate faster and may leave water behind. This water may soften gelatine capsules, may smudge labels and may contribute to mould growth. In dry climates, air drying is adequate but in more humid environments, a drying chamber set to 40–50°C may be advantageous. Simple drying chambers can be constructed using incandescent light bulbs in a simple box with some openings to allow for air circulation. Blow-dry systems are unsuitable because the specimens are too easily blown away once dry. Handling Dry specimens may be kept in glass vials, gelatine capsules, or cardboard slides, with or without cushioning cotton. Specimens prepared for scanning electron microscopy (SEM) may either be stored mounted on the SEM stub (see below for storage conditions), or may be dismounted (see below) and placed in a standard container. To properly view specimens, they usually need to be removed from their glass vial or gelatine capsule, whereas the glass window of a cardboard slide usually allows adequate viewing. Specimens are best placed onto a metal, glass, or paper surface of contrasting colour. Avoid plastic as the static charge can make it difficult to move dry specimens. To move individual specimens into separate containers, a moist (but not wet) artist’s brush is safest, but forceps can also be used with care (see Tools section above). Dip the brush into clean or, preferably, distilled water or ethanol and squeeze the bristles between two fingers or touch a piece of paper. Some people use saliva but, apart from hygiene issues, it can leave residues on the shell that are obvious under the SEM. The specimens will adhere to the tip of the moist brush. Deposit the specimen in the new container on the inner lip or wall of the container; rotating the axis of the brush while keeping the specimen touching the container helps to transfer the specimen from the brush to the container. When transferring specimens into gelatine capsules, make sure that the brush contains as little moisture as possible (gelatine is soluble in water) as specimens may GEIGER ET AL. (2007) MOLLUSCAN RESEARCH, VOL. 27 become glued to the wall of the capsule. A dry, soft artist’s brush may also be used for the most fragile specimens, particularly if the brush is somewhat frayed. Specimens can be gently ‘speared’ so that they are held between the bristles of the brush. Multiple specimens can be poured into the storage container from the sorting tray. If a square dish is used, the corner may be suitable to concentrate and spout the specimens into a container. Alternatively, pour or brush the specimens onto a tightly folded piece of paper, then pour them into the container using the angulation as a guide. Gently tapping the paper may help move stubborn specimens. Other techniques include making a funnel out of paper or using a small glass or metal funnel (avoid plastic). Storage Specimen containers First, the specimens, even those collected as empty shells, must be properly cleaned and free of salt (see above). This can be done by soaking them in clean (preferably distilled) water for a few hours and then drying thoroughly. The dried specimens are best kept in small containers: ideally gelatine capsules within larger high sodium glass vials, or cardboard mounts made from acid free, archival quality board. None of the storage media is superior for all storage conditions; each has its advantages and disadvantages. The following discussion of the materials assumes that the specimens are in direct contact with that material. In many collections, specimens are contained in gelatine capsules or glass microvials and placed within glass vials that hold the label. Glass vials are the most durable solution, but are also costly. Gelatine capsules do not degrade shells, but should only be used in conditions with < 60% average relative humidity. Shorter periods of 60–80% relative humidity do not seem to have a negative impact. Gelatine is hygroscopic, thus, in conditions of high humidity, and with insufficiently dried specimens, the gelatine softens and may glue the specimens to the wall of the capsules. Usually, a gentle push with the stiff bristle of a fine artist’s brush will free the specimen. Otherwise, the gelatine can be fully dissolved in water. Specimens do not seem to be damaged by sticking to gelatine. Insects can also eat the gelatine capsules if they are left loose, leaving holes for shells to fall out. Cardboard slides, also called geology micromounts or microfossil slides, are very space efficient. However, they are usually made from acidic paper and may negatively affect specimen preservation, particularly in humid areas. They release nitric acid from the celluloid, which is known to dissolve foraminiferans (Barbero and Toffoletto 1996). Cardboard slides may be custom made using archival quality material (see below). Another drawback is that sliding the glass window usually generates a static charge and specimens become stuck to the glass. Fragile specimens may become damaged when the glass window is pulled through the slit in the cardboard. For sources and types of microfossil slides see http://www.pangeauk.com and http://
TECHNIQUES FOR STUDYING SMALL MOLLUSCAN SPECIMENS 15 www.ukge.co.uk. Chlorine bleaching (Clapp 1987) of natural cotton wool can release acid and may negatively affect the specimens, therefore, artificial cotton is more suitable. Some of us use medical grade cotton wool with glass tubes and have not observed any shell degradation. Avoid any polyvinylchloride (PVC) products, as they release hydrochloric acid from the plasticiser. Most micromolluscs do not have sufficient mass to damage one another and cushioning is not necessary. Small plastic boxes are sometimes used and some consider them superior to gelatine capsules (Kurtz 2005). We are cautious about plastics because of the plasticisers used in the material and different types of plastic are not easily distinguishable. Transparent Polystyrene boxes seem acceptable. Mylar is the only clear plastic film known to us to be of fully archival quality—it is also widely used in the fine arts business. Plastic foam has been and, to a certain extent, still is today, popular with shell collectors to hold specimens in place within a plastic box. Plastic foam is the worst possible storage for any shells and is particularly insidious for micromolluscs. All foam disintegrates in one way or another over a relatively short period of time (10–20 years). At best foam crumbles and specimens have to picked out of the material but usually it partially liquefies and becomes sticky. It can be rubbed off larger shells, but micromolluscs can usually not be adequately cleaned. The plasticisers in the foam or the foam matrix itself can also be acidic and cause an efflorescence akin to Byne’s or glass disease. Some shell collectors use commercial plastic clays, often referred to as mineral micromount, or tackless picture mountants. These are supposed to not release any grease, although they eventually do, leaving wet-looking patches on shells. Their long-term stability is quite variable. Over a 15 year period in a collection of approximately 2000 lots, some 80% of the material seemed to be unaffected by the plastic clay, although long-term contact left residues in the grooves and lamellae of finely sculptured specimens: 10% of the material became tacky and stringy and some 10% crumbled. The cause for the material’s decay is uncertain; oiled shells, and those containing animal remains, seem to be more frequently associated with the tacky or crumbled clay (Geiger 2004). As many micromolluscs are finely sculptured and as the clay may change dramatically over relatively short periods of time, this mounting medium is unsuitable for microspecimen storage (Geiger 2004). Specimens are sometimes stored on SEM stubs, though most shells can be removed from them without problems (see below). Usually, recommended storage of SEM stubs is in sealed containers with silica gel to remove moisture and, if possible, in an evacuated bell jar. These measures ensure that the specimens will not outgas when placed in the high vacuum of the SEM chamber, particularly those with field emission and LaB 6 guns, which require a vacuum at least an order of magnitude greater than for tungsten filaments. Hence, these storage requirements rather address operational issues of the SEM and do not relate to specimen preservation. Whether such storage conditions actually make a difference will depend on local environmental conditions and on the particular SEM used. The re-emergence of tungsten guns, especially on variable pressure SEMs, makes the above storage requirements unnecessary; protection from dust and storage in a normal collection environment is sufficient. Labels Ideally, the full-data specimen label should not be in direct contact with the specimens, but contained in a secondary container (usually a glass vial) housing the gelatine capsule or glass microvial housing the specimen(s) or on the backside of a geology micromount (as in the LACM). In some collections, a tiny label showing only the registration number is added to the specimen vial. Such labels offer added protection against switching data between lots. However, even small paper labels in direct contact with microspecimens should be made from acid-free paper. We are unaware of any issues relating to writing medium; pencil, pigment ink and laser writer labels; all seem to work well. There is anecdotal evidence that photocopied labels are more durable than that of laser writers because the temperature at which the toner is fused with the paper is higher in photocopiers than in laser writers (H. Chaney, pers. comm.). Heating labels in a direct heat oven improves the durability of laser print; the toner changes from a powder-like appearance to a shiny surface when examined under a microscope; ‘cooking’ labels with microwaves does not affect the durability of laser labels (Zala et al. 2005). Elevated humidity has an adverse effect on laser printing, because the temperature is lowered by the residual water in the paper. As a safety measure, the catalogue number can be written in pencil on the back of the label. Paper as a material should be carefully considered in all applications: labels, geology mounts, cardboard boxes. The term ‘archival’ can be misleading, as a number of paper products of variable long-term stability are issued with such a descriptor (Clapp 1987; Turner 1998). Acid free paper products are made from 100% cotton rags, not from wood pulp, are usually not bleached and will not release any acids. Buffered or pH-balanced products are made from wood pulp and contain a pH-buffering substance, usually calcium carbonate powder. The wood pulp will continuously release acids, which at first is neutralised by the calcium carbonate in the paper, but eventually the buffer capacity is exhausted and acids will be released from the paper product. Synthetic/ plastic papers have not been available for a sufficiently long period of time in order to assess their suitability as data labels. A good start for internet searches is http:// www.universityproducts.com, http://www.archivalsuppliers. com, http://www.archivescanada.ca/english/index.html and http://library.amnh.org/conservation/suppliers.html. The strongest paper we have found is ‘laundry tag manila’. Coated papers in particular are unsuitable because of the chemical coatings and filler materials, which will eventually deteriorate.
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