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

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28 slowly rotate the column bottom while carefully pulling the filter from underneath the retention ring from one segment after the other. Wash the radula in water and mount. Retrieval GEIGER ET AL. (2007) MOLLUSCAN RESEARCH, VOL. 27 rate is approximately 80–90%. Proteinase K, though, does not work well on formalin fixed material, in which proteins of the tissue have been cross-linked (Holznagel 1998). FIGURE 11. Recovery of the radula from a spin column used for DNA extraction. A. Cut the column above the retainer ring for the filter. B. Oblique view of the cut column. The arrow highlights the radula. C. Enlargement of B with the radula visible near the edge of the filter retainer ring. D. The radula has been removed with fine forceps and is placed into a gelatine capsule for temporary storage. Images DLG. Cleaning the radula To find the radula after maceration, examine the container under the stereo-microscope, using the substage illuminator. Incident light may make particles shine and hide the radula. Horizontal illumination through the sides of the glass vessel may be employed if a substage illuminator is not available (pseudo dark field), but will only be effective if the fluid is clear. The radula has to be washed in water and then possibly in ethanol. Two methods are outlined below. In the first, fluid is exchanged, minimising handling of the radula. The fluid can be removed with a Pasteur pipette or a pipettor and discarded into a separate container in case the radula is inadvertently removed with the fluid. The remaining fluid film can be blotted with a fragment of folded paper tissue, keeping it a safe distance from the radula. A small radula that is stuck to the paper will usually be lost because it is difficult to distinguish the radula from the paper fibres. It is easiest to move the radula when dry by touching it with a moist tungsten needle, or very fine, moist insect pin. Eyelashes or other hairs are too flexible for radular manipulations. In the second method, the radula is transferred in a succession of fluids, which prevents it from drying. The radula is picked up with fine entomological forceps, a bent needle, or with a pipettor and transferred into a series of washing solutions. Do not use a paintbrush to transfer a radula, as it can easily get entangled in the bristles and lost. Minute radulae can be washed in drops of water on a glass histology slide. Clean radulae can be stored in tubes in 80% ethanol. Sometimes the maceration solution is very dirty and it may help to dilute it with distilled water or more KOH solution. Heat may also help if nothing else does. As a final resort, the contents of the container can be poured through a very fine sieve with low edges. The mesh should be a
TECHNIQUES FOR STUDYING SMALL MOLLUSCAN SPECIMENS 29 maximum of 1/10 of the estimated length of the radula (as a general guideline, the radula is approximately one tenth to one third the length of the shell/body). Rinse with hot water, which will usually dissolve the remaining grease, tissues and particles. Medium size radulae may be brushed with a very fine paint brush (#00 or #000, preferably artificial hair), while holding the end of the radula with a needle pressed against the glass. With very small radulae there are few possibilities for further cleaning; they can be moved in the water or scratched with a fine needle against the glass so it makes small vibrating jerks. Use a needle with its tip bent close to 90° and scratch with the tip at right angle to the glass. If the radula still looks dirty, try a new KOH bath. If nothing else helps, try diluted, warm bleach (rather than cold, more concentrated bleach, as less hypochlorite is carried over to the rinse). Neogastropod radulae have a sheath of thin, transparent cuticle surrounding the part not in use. This should be removed by a quick dip in commercial bleach and subsequently rinsed, or with a fine needle when it has been glued and is drying. Do this at the same time as the lateral teeth are unfolded. Do not be concerned about the radular membrane extensions that under-lie the anterior part of the radula in vetigastropods and taenioglossate caenogastropods. They may even facilitate mounting. In general, do not use an ultrasonic cleaner, unless there are spare radulae. It is a good method for cleaning many taenioglossate radulae, but in some cases teeth may fall off, get entangled, or the radula may disintegrate entirely. Not even experienced practitioners can predict the result. SEM mounting of micromollusc radulae Mounting can be achieved by a variety of techniques, the choice mostly depending upon size and the type of radula (Fig. 12). Techniques for light microscopy are not discussed here as they are detailed elsewhere (e.g., Mikkelsen 1985; Coovert and Coovert 1987; Bradner and Kay 1995). Hickman (1977) discussed the types of information to be gained from both light and electron microscopy of radulae and stressed the complimentary nature of both techniques. Large radulae (>1 mm long) can be mounted on doublesided sticky tape, or double-sided carbon tabs. Medium sized radulae can be mounted on double-sided sticky tape and manipulated in a drop of water or ethanol (Fig. 12D). Some workers also like to roll the radula onto pins or conical mounts as the information gained increases the more the radula is folded or twisted (Bradner and Kay, 1995). The slow evaporation method of Moretzsohn (2004) seems to work well for larger radulae, however it is not clear whether it would work as well if applied to the preparation of small radulae. Minute radulae can be dried directly onto a piece of coverslip (see below). Consideration of storage options and times should also be taken into account when determining the mounting method used. Orientation Identifying the orientation of the radula can be difficult. Use the highest magnification of the dissecting microscope. If the radula is too small to see individual teeth, rely upon the appearance. The anterior margin of the radula with completely formed teeth has two flaps. The teeth there are facing outwards from the curve. The posterior end of the radula is tapered and thinner than the anterior part. The upper and lower surface of the radula can be difficult to distinguish but they reflect light differently; the underside has a more glassy appearance, whereas the top is more sparkling. The long axis of the radula tends to curl with the base inside; the curling of the radula along its long axis is often impossible to see. It is easy to distinguish which way up a radula is lying by transferring a wet radula to a fragment of a cover slip on a slide and examining it under low-power using a compound microscope. The radula can then be mounted directly on the cover slip. For very small radulae, it may be difficult to judge whether it has been properly mounted. To ensure suitable results, several radulae can be mounted, or the radula can be folded in a L or V shape so that both sides are facing upwards (this method is especially useful for elongate radulae and for species with short teeth). Tilting the stub towards the light will usually show the reflection pattern better. Special techniques for small radulae Several of techniques have been successfully used to mount very small radulae. These include the use of different surfaces such as: • Double-sided sticky carbon tabs (Fig. 12C). Other sticky substances include double-sided tape and wet, blackened, photographic paper. Place a small drop—the amount of water that sticks to the head of a fine pin— on one of these surfaces mounted on a metal stub. Place the wet radula into this drop and orient it. • Radulae of species with many teeth per row (e.g., rhipidoglossate, ptenoglossate, many pulmonates) can be placed directly on a glass cover-slip glued onto a stub. Orientate the radula in a small drop of water and let the water evaporate. Excess water can be removed with great care with a fine strip of filter paper (see above). Alternatively, pull the radula out of a drop of water and let it dry in place. The radula will adhere to the glass without any need for adhesive. Some narrow, taenioglossate radulae may tend to spring off the glass and may need to be mounted on carbon tape. Radulae mounted on cover glass can, with some practice, be removed with water even after viewing in the SEM. • Glue a histology cover-slip (round, 12 mm diameter) to the mid-point of the histo-slide with a very small amount of saliva. Let it dry well in an incubator. Cover part of the cover-slip with a thin layer of
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