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

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12 Fixation The intended use of the specimens should determine the fixation and storage fluid. For fixation and storage for specialised needs we recommend the following: • Molecular work—95–100% ethanol. See also ‘boiling method’ below. • Histology—formalin, bichromate or mercury-based fixatives, Bouin’s fluid or other histological fixatives. • TEM—ideally glutaraldehyde fixation. Formalin can also be used but with inferior results. Detailed and complicated descriptions and recipes for fixation and preservation are available but are mostly unnecessary for standard work. Also, most methods work well within a wide range of concentrations and often one kind of buffer can be replaced by another as long as they do not interfere. For example, recipes often specify that 3.7% formalin is to be used. That is simply because they used formalin : water, 1:9, but good fixation with formalin can be achieved as long as it is stronger than ca 2%. For more general information regarding fixation and preservation see Gohar (1937), Romeis (1948, 1989), Mahoney (1973), Presnell and Schreibman (1997) and Glauert and Lewis (1998). When fixing shelled molluscs, the fixative must have access to the tissues; a light cracking of the shell is usually needed, except in limpets, chitons and gastropods with a short, broad spire, small operculum and large aperture. To crack small specimens may be difficult without crushing them. A small pair of wire cutters for electronics is usually good; some models are made of stainless steel. Also, for larger specimens, a bench vice, locking vice pliers, or any other tool where you can control the cracking is better, to avoid crushing the shell. Power pliers with an extra joint for increased power are usually good for larger specimens with thick shells. Watchmaker’s forceps can also be used like a nut-cracker. Insert the specimen about a quarter of the length of the handle from the join, with one face of the forceps on the table, and gently press the other arm of the forceps until the specimen cracks. However, this method requires practice as it is liable to crush the shell unless carefully controlled. More drastic measures (e.g., a small hammer) may break the shell into many pieces and reduce the animal to pulp. Drilling a hole in the back of the shell (see above) and injecting 95–99% ethanol is an alternative for larger species (>3–10 mm), but is not as safe as cracking. For most studies involving micromolluscs, the shell is one of the most important sources of taxonomic information. For this reason, if shells are cracked or removed prior to fixation, it is important to keep an undamaged specimen for reference purposes—even an empty shell will often suffice. Because micromolluscs often have little shell material, they are particularly prone to adverse effects by preservation fluids. Acidic formalin or ethanol can quickly damage or completely destroy shells. However, formalin is a good general fixative for tissue preservation and samples can be used for TEM, SEM etc. Its biggest downsides are that the GEIGER ET AL. (2007) MOLLUSCAN RESEARCH, VOL. 27 material cannot be used for molecular studies with current techniques and it is carcinogenic. Marine samples may be fixed in 5–10% formalin-seawater, which is sufficiently buffered for short time fixation (1 day) at a pH of approximately 7 (Anonymous 2006a). It is important with any fixative to have an appreciably larger volume (factor of at least 5–10) of fixative than the specimen. For formalin fixation, a quick approach is to fill a container with molluscs, add 1/10 of the jar volume in 40% formalin and top off with water (seawater is preferred as it provides a more nearly isotonic solution). Mix the solution well by inverting the jar several times until there are no more streaks in the fluid. The bodies of the animals will provide the remainder of the water to make an approximately 5% formalin solution. To properly buffer formalin, 1 g of borax per litre seawater formalin gives a pH of 7.5–8.5 and is good for several years. However, borax may clear tissue during prolonged storage (>10 years: Anonymous 2006a) and may be considered unsuitable, although some of us have not noticed any detrimental effects. Excess sodium bicarbonate mixed with formalin (allow it to settle for several hours) made up with fresh or seawater gives a pH of approximately 8. If instead sodium carbonate is used, the pH becomes approximately 10, which is much too basic, it becomes histolytic and the skin peels off within a few years. Other buffering agents include powdered aragonite, which is more soluble than calcite, but may recrystallise and interfere with shell material, and household ammonia, which reacts strongly exothermically with formalin to form hexamine (= hexamethylenetetramine, methenamine) to pH 8.2 (Clark 1998). Hexamine decays and has to be adjusted after one and six months, and then every two years (Hemleben et al. 1988). This labour-intensive procedure will be prohibitive for larger collections (See Appendix 1 for safety notes). Recently several ‘formalin free’ fixatives and preservatives have appeared on the market. Some are based on phenoxetol, which does not replace fixation for histology or SEM purposes. Storage The most commonly used storage medium is 70–80% ethanol. Borax, powdered aragonite, or powdered calcite/ shells may be added to ethanol solutions to safeguard against shell damage. Precise amounts have not been specified, though some have expressed concern that borax and aragonite may pose problems with recrystallisation; this area needs further investigation. To reduce the problem with dissolution of shells in water-ethanol mixtures, the concentration of the alcohol can be increased. For histology, tissue shrinkage is of concern and it is customarily advised to use a graduated series (30%, 50%, 60%, 70% ethanol) when transferring specimens from aqueous to alcoholic solutions to minimise shrinkage. Glauert and Lewis (1998) question this approach, because
TECHNIQUES FOR STUDYING SMALL MOLLUSCAN SPECIMENS 13 The alcohol concentration can be kept stable in wellsealed containers. Specimens in tubes with polyethylene (never polycarbonate, which will disintegrate) closures or cotton plugs should be immersed closed end down in ethanol in larger, secondary containers of proven durability. Because ethanol vapours consist of approximately 95.5% ethanol, evaporation reduces the alcohol concentration in the medium and evaporated liquid should be replaced with 95.5% ethanol. Calcium carbonate is quite soluble in water, hence shells may easily be dissolved in an ethanol-water mixture. For example, scissurellids can become fully decalcified in as little as 18 months in ethanol at less than 80% concentration (DLG, pers. observ.). As it is impractical to monitor alcohol concentrations in small vials every six months, it is advisable to store some shells dry as vouchers. Storage of micromollusc samples with large amounts of organic material should be avoided. Ideally use high quality ethanol free of impurities, although this is much more expensive. Long-term storage in formalin is generally avoided since formalin is on the list of suspected carcinogens and is a well-known allergene. However, it has been successfully achieved in at least one major collection (AMS), where 5% seawater formalin buffered with NaHCO 3 is used. Problems with ethanol include evaporation and flammability, with collections requiring regular maintenance and special fireproof housing. For any preservative, the pH needs to be kept below pH 8.5 to avoid tissue dissolution and above pH 7 to prevent shells and other exoskeleton parts from dissolving. Such narrow tolerances require regular testing and adjustments. The pH of ethanol-water solutions is difficult to measure, though some specialty pH electrodes are available. For field storage, unbreakable plastic containers, ideally with screw tops, should be used. Eppendorf tubes (1.5 ml) and Falcon tubes (10 or 50 ml) seal fairly well, as long as the seal is free of dirt. Heat-sealed bags can leak when filled with wet sediment samples, but are useful as secondary containers. Although cheap, scintillation vials should be avoided, because ethanol evaporates quickly from them. Scintillation tubes are also potentially dangerous during air transportation, since they do not close well and are not intended for such use. Switching storage media When switching specimens from one solution (e.g., 5% formalin) to another (e.g., 70–80% ethanol), the tissue volume and other water filled spaces need to be taken into account, as water they contain will dilute the preservative. As an example, a jar half-filled with specimens and filled up with 95% ethanol will eventually result in about a 50–75% ethanol solution. Hence, for samples intended for molecular work, it is important to replace the solution with 95–100% ethanol within the first two days. Boiling method Dr H. Fukuda has kindly provided details of a method that he has successfully employed for microgastropods which is a modification of a method used by a number of Japanese malacologists for large species and is known as ‘niku-nuki’ (e.g., Habe and Kosuge 1967). It has proved to be particularly useful for instances where only one or two individuals are available and intact shells and animals are required. A living individual is placed in a small beaker in enough water (seawater if a marine species) to enable it to extend and crawl. Add hot (70–100ºC) water, which will immediately kill the animal with the head-foot extended. After a few seconds (1–2 for minute species) in the hot water, move the specimen to a smaller dish of cool water under a stereomicroscope. The animal can be carefully removed from the shell by gently pulling on the head-foot with forceps, holding the shell with a second pair of forceps and rotating in opposite directions. The animal removal can be facilitated by squirting water into the aperture using a fine syringe. The water temperature and the length of immersion in the hot water vary according to the size of the specimen and the thickness of the shell, with larger, thick-shelled species requiring hotter water and longer times. The visceral mass (digestive gland and gonad) becomes hard and loses flexibility in high temperature and sometimes cannot be removed from the upper whorls of the shell. Thus, the water needs to be hot enough to separate the columella muscle from the shell and cool enough to keep the visceral coil pliable. More details on this method will be provided elsewhere (Fukuda, Haga and Tatara in prep.). As DNA is not broken down in 80 to 100ºC, tissue can be placed in 99– 100% ethanol for molecular work (Ueshima 2002). Dry micromollusc shells Two ‘diseases’, or more correctly, chemical processes, that ultimately result in the destruction of shells, have affected many type specimens in museum collections including AMS, BMNH, NMNZ, NSMT, NHMB, USNM and ZMO among others (Fig. 5C). Some collections are more affected than others, with no clear pattern emerging. The collections in GNM, SMNH and ZMUC have largely escaped it, presumably by using different types of glass. Typical instances are illustrated by Higo et al. (2001) in micromolluscs such as triphorids and turrids. Two different kinds of ‘disease’ are recognised, Byne’s and glass, but it is sometimes difficult to decide which particular ‘disease’ is responsible (e.g., Kilburn 1996). The manifestation of both diseases is identical in that they first produce white efflorescence on the shell, which eventually crumbles to dust, but they differ in the cause. Species described prior to 1960, and many thereafter, were illustrated without the benefit of the SEM. Much needed detail for species level identification (e.g., protoconch microsculpture) cannot be observed using light microscopy. Therefore, many species cannot be positively identified from the original descriptions or illustrations and type material is the sole recourse to settle uncertainties. Accordingly, it is a high curatorial priority to upgrade storage systems of micromollusc material, particularly types, and to
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