Plastic bags and buckets, always with enough seawater to cover the specimens, are convenient ways to collect. Change the water from time to time to avoid temperature increases. If possible, photograph the specimen before collecting it and ensure that you have a reliable system to match the specimen to the photograph (because after you have collected it that ascidian will never look the same again). The colours of living specimens
Figure 27.6 A12, Diplosoma simile larva. A, immature larva with the tail curved around the ventrum of trunk, anteromedian adhesive organs, the rastrum (a T-shaped outgrowth from the larval haemocoel dorsal to the base of the tail at the posterior end of the trunk) differentiated into hairs; A2, tip of the right horn of a mature larval rastrum (indicated in A1) showing Prochloron cells entangled in hairs differentiated from the thick larval test covering the rastrum. B , Didemnid CaCO3 (aragonite) spicules: Didemnum albopunctatum (globular spicules with rod-shaped rays), Didemnum membrana-ceum (stellate spicules with conical rays and some giant spicules). (Figure: light micrographs, Kott 1982; SE micrographs, Kott 2001.)
are conspicuous, especially in colonies where the surface is not obscured by epibionts and sand. However, there is a great deal of intraspecific variation in colour and usually it changes when specimens are removed from their substrate. In the field, ascidians can often be confused with equally brightly coloured sponges although, unlike sponges, the apertures of the ascidian close tightly when it is disturbed.
Field notes should be made in situ before the animal is disturbed and any changes resulting from disturbance, including its removal from the substrate, should be noted together with colour, size, shape and details of the zooid arrangement in the colonies and notes on the habitat, location, collecting method and depth. Make the notes in pencil on good quality waterproof paper and put them into a plastic bag with the specimen; or in a field notebook, carefully cross-referencing each entry to the number that you keep with the specimen. It is a good idea to take a colour chart with you when you collect ascidians. (Make a replicate of any reliable standard by matching its colours with colour chips from the paint charts you can pick up at any hardware store; then paste up the matched chips on a few sheets of cardboard, waterproof or seal them in plastic and carry them with you in a black plastic bag to avoid fading.)
Dislodge specimens carefully from their substrate using a sharp fishing knife. Sample the colonies too large to collect (it is not necessary to take the whole specimen). However, draw the whole specimen in a field notebook or on the back of the label, note its general dimensions and shade or indicate in some way the portion of it that you have sampled. Take the sample so that it is representative of the colony, that is, cut a wedge from the centre of the colony to its outer margin, or from a large common cloacal aperture to the outer margin, or take a whole lobe or branch. Always cut vertically, from surface to the base, parallel to the zooids. Never cut an ascidian colony horizontally or you are likely to cut zooids in half. If you cut into a didemnid colony, do it underwater as damage causes cells to generate acid that can dissolve calcareous spicules. The sea water will help to neutralise the acid quickly.
Narcotise specimens with menthol crystals for up to three hours (colonies) to five or more hours (large solitary specimens) by using just enough water to cover the specimen and closing the bag or vessel as closely as possible. The specimen is narcotised if the siphons do not close when stimulated. As menthol is a bacterio-cide, the specimen remains in good condition despite the long time taken for narcotisation. Also, menthol is expensive so save the menthol crystals when narcotisation is complete. Do not on any account use an alcoholic solution of menthol or you will merely kill the specimen before it is narcotised.
Fix the specimen by quickly adding to it one part of 40% formaldehyde to the nine parts represented by the specimen, plus enough seawater to cover it. Do not add formalin already made up to 4%; if the specimen is the same volume as the preservative you will end up with only 2% formalin. In other words think of the specimen as water (which it mostly is). Keep the preservative neutral with a little calcium carbonate in the water. (Note. Formalin is generally used to describe a dilution of 40% formaldehyde, so that 10% formalin is actually 4% formaldehyde.)
Preserve the formalin-fixed specimens after at least three months by transferring them to 70% ethanol with a dash of glycerol.
Permits are required for collecting ascidians, see Chapter 12 for details.
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