A guide to methods of preserving
animal specimens in liquid preservatives.
Originally prepared for the Zoo Hons ‘94 field course
by Kelvin K. P. Lim & N. Sivasothi
I. OBJECTIVES OF PRESERVATION
In this guideline, we are mainly concerned with the taxonomic reasons for preservation. The scientific description of an animal species requires the detailed examination and description of a representative type specimen and a series of specimens which are subsequently deposited, catalogued and maintained in a museum or zoological collection. This remains a reference for other workers to consult in future.
Specimens from any field collection should be deposited in a reference collection in an institutional for the long-term maintenance and access for the future. The animals should therefore be preserved in the best possible condition and where possible, ensure that the natural colour is retained, their external appendages (e.g. fins) are erected and stomach contents intact.
Care should be taken to ensure that specimens are undamaged. Features important in the taxonomic study of fish, for example, are easily damaged with contact even after preservation. Live crabs before preservation should be kept individually as some species will damage each other and other animals, especially fish even when they are being directly preserved.
Specimens collected during an expedition are to be killed immediately on site. Photography, if required shoud be conducted on the spot. Do not crowd living animals in small containers - this will result in damage to their surfaces or appendages. Do not keep animals for preservation “later” as it may die and pollute a container, killing others, even leading to a distortion of morphological features and other damage. This reduces their value as scientific specimens which is the objective of colelction in the first place. A well-preserved specimen will generate more accurate information and is ultimately more humane.
II. THE PRESERVATIVES
Note: A specimen immersed in a liquid preservative may not be totally infused with the liquid. This may be noticeable when the specimen floats. In the long term, such a specimen will rot, despite being surrounded by a presevative. In large volume animals, injection of the specimen with the preservative is required to reach the inner tissue.
1. Formalin
Usage: We do not use formalin for preserving arthropods, only vertebrates. Specimens are not kept permanently in formalin since it is acidic and difficult to handle.
Although generally avoided for long-term storage, formalin may be used in instances where colour is important since alcohol dissolves most colours almost immediately.
i. Formalin - 1 week (fix soft tissue)
ii. Water - 1 day (leach out the formalin)
iii. Alcohol - long term storage
These are aproximations based on fish with lengths of up to 30cm. The length of time for each step may have to be increased with increasing size of specimens.
NOTE: Conc. (100%) formalin = water saturated with 40% formaldehyde.
We use 10% formalin (= 4% formaldehyde) in preservation. Whenever we mention concentrations of this preservatve, we refer to formalin and not formaldehyde.
Thus we carry concentrated formalin and dilute it at sampling sites: Mix one part concentrated formalin to nine parts water. In bottles meant for killing and fixing fishes, fill about two-thirds the bottle’s volume with 10% formalin. As formalin is acidic, it should be buffered by adding a pinch or two of sodium bicarbonate.
Warning: Inhalation of formalin fumes causes extreme discomfort and could be harmful. Contact with the fluid causes severe irritation to the skin; contact with sore or raw spots results in extreme pain. It is also said to be a carcinogen. In the event of spillage, always rinse hands, the outside of receptacles and other affected areas thoroughly with water.
Storage: Concentrated or any other forms of formalin should be kept in safe, water-tight, spill-proof bottles, e.g. pep-bottles, and should always be clearly labelled. Plastic bags containing formalin should be leak-proof and maintained in an upright position at all times, and the opening should be rolled or folded down and sealed with tape or clips.
Securely cover all receptacles for formalin fumes can be very offensive smelling, and can cause extreme discomfort to the nose and eyes.
2. (Industrial) Alcohol
Usage: Alcohol is usually not used for killing and fixing vertebrates, but is used for most arthropods. Insects, crustaceans and arachnids can be simply dropped into alcohol for immediately preservation. Note that the colour of a specimen is lost almost immediately once immersed in alcohol.
Alcohol usually comes in the 95% concentrated form. For long-term preservation, it is usually diluted with water to 70-75% strength. This is the lowest concentration at which preservation will be maintained. During field collections, ensure that solution you use is not diluted by the water which comes with the samples.
Warning: Alcohol is usually safe to handle, but can cause irritation to the skin in cases of prolonged contact. Always rinse hands thoroughly with water after working with alcohol. Industrial alcohol is toxic and should never be drunk. Receptacles containing alcohol should always be properly and clearly labelled.
Alcohol is highly flammable. Never work with this fluid in the vicinity of open flames. Alcohol is prone to rapid evaporation, and receptacles holding it should be securely covered at all times, and not be opened unnecessarily.
III. THE RECEPTACLES
There are various types of containers which one can use to fix specimens:
1. Plastic water-tight bottles. Where possible, the bottle-type containers should be as tall as the length of the largest specimens, such that the specimens can be set straight when the bottle is placed on its side. It can however, be difficult to handle in the field where one has to screw and unscrew the bottle-cap numerous times in a row.
2. Plastic bags. Not as desirable, but with careful usage, and with knowledge of its limitations, these can be successfully utilised in the absence of plastic containers. Its disadvantage lies mainly in that its leak-proofness is not guaranteed, and also it can easily be punctured by sharp spines. When plastic bags are used, go for those with thicker material. Plastic bags are very useful for fixing the larger specimens, as well as for containing mud and leaf-litter samples.
3. Shallow trays. May be of metal or plastic. These are mainly used for fixing and setting specimens of reptiles and amphibians. The animals are set half-submerged in formalin in the tray which is then placed within a plastic bag, or covered with a plank. They stay like this for one or two days until they are hardened before being placed totally submerged in preservative for further fixing.
IV. PROCEDURE
1. Data labels: Always make sure to mark or insert a data sheet into the bottle for every site collected. This should preferably be done immediately before leaving the site for another. A data sheet is extremely important for it records the place where the specimens were collected, and the date of collection. Specimens without such data are virtually worthless. Data sheets should not disintegrate in liquid and should be labelled with waterproof ink or soft pencil.
2. Bottle labels: Always label bottles with preservatives clearly, so that the contents will not be misused, or worse, drunk accidentally.
3. Positioning fish: When using bottles, always place fishes with strong spines on their fins into the bottles backwards. This will enable easy extraction of the specimens later, even if the spines become erected, without having to damage the spines.
4. Injecting large specimens: Large fishes above 10 cm, crustaceans and herptiles should preferably be injected with a small amount of 10% formalin after they have expired in the preservative. Fishes larger than 6 cm and all crustaceans and herptiles, when procured dead, should always be injected before they are fixed in formalin. This prevents the innards from rotting, a condition which leads to distortion of the specimens.
When injecting specimens, find a place with running water, so that hands can immediately be rinsed in the event of spillage of formalin. Inject 10% formalin into the body cavity of the specimens through the vent until the body bloats (only slightly) and formalin starts to drip from the mouth. For larger fishes and herptiles, an incision could be made on the belly to enable formalin to penetrate the body cavity.
5. Long specimens: When dealing with long specimens, always place bottles on their sides, such that the specimens could be preserved straight, and not curved.
6. Handling: Always make sure that the cap of the collecting bottle is securely screwed, so that the preservative fluid does not spill, and specimens do not get lost. This would also ensure that the animals in their death throes do not jump out, and gets lost or damaged; and it cuts down evaporation of the preservative. Furthermore, struggling fish often spalsh formalin into the eyes of ‘concerned individuals’.
V. THE ANIMAL GROUPS
1. Fishes
Whenever possible, all fish specimens are to be immediately placed in 10% formalin upon capture. This is undoubtedly extremely painful for the animal, but death is usually fairly quick, and not only that, the fish usually dies with its finnature well spread-out, and the body straight and well-stretched. Examination and counting of fin rays and scales would be relatively easy on such well-preserved material.
Specimens are usually left in formalin to be fixed (ie. become thoroughly hardened) for a week or slightly more if larger than 10 cm, before they are soaked for a day or two in clean freshwater before being transferred to 70-75% alcohol for long-term preservation. It is possible to preserve fish specimens for a long time in formalin, but if the preservative is not buffered, the high acidity is likely to render the specimens brittle and transparent.
2. Herptiles
We usually do not kill herptiles by throwing them into formalin. They are usually kept individually in plastic collecting tubes or plastic bags until they could be killed either by freezing or chloroform.
It is also important to set the specimens carefully so they do not take up too much space. Shallow trays are useful for setting herptile specimens. Snakes should be coiled up, and frogs are to be placed on their bellies with their limbs set at right angles to their body. All digits should be stretched out flat and not left to curl. A long pair of forceps is useful for such procedures. Depending on the size of the specimens, they are to be set half-submerged in formalin for a day or two, then transferred to full-submergence in formalin for further fixing. After a week or more, they can be soaked for two or three days in water before transfer to 75% alcohol for long-term preservation.
3. Arthropods
These are the easiest to process in that they can be killed immediately and stored in alcohol. Crabs and prawns may also be killed in formalin, but this renders their joints hard and brittle, and their limbs cannot be easily manipulated. The larger arthropods (especially those with hard exoskeletons) sometimes need to be injected with 10% formalin to prevent their innards from rotting.
4. Molluscs
It is important to relax the animal before preservation. Apparently the most satisfactory way to do that is to drown the animal in cool boiled (deoxygenated) water. The mollusc could be enclosed within a (as small as possible) plastic vial of water, making sure that no air bubbles are trapped within to speed up the drowning process. This may take from one to about three days. Frequent periodic checks need to be made on the specimens for they can rot rather fast once they expired. Before injecting the animal with a little formalin, make doubly sure that it is limb and lifeless. Try pinching the eyestalks (note that even this is not foolproof). Upon the slightest contraction, repeat the drowning process quickly.
It is important to preserve relaxed molluscs to gain an idea of their appearance in life. Newly killed molluscs could then be fixed in buffered 10% formalin for two or three days, then transferred to 75% alcohol after soaking for a few hours in water. Molluscs can also be relaxed in a solution of magnesium chloride, but this does not work very well with land molluscs.
The meaning of ‘relaxed’: An animal, when it becomes relaxed, is limb with appendages fully stretched out, and muscles loosened. A snail when not relaxed, will be contracted within the shell, and its foot and eyestalks would not be externally visible. A frog or snake, when not relaxed prior to preservation would often become contorted into awkward poses. It is usually not needed to relax fishes or arthropods meant for liquid preservation.
5. Warning
Do not mix large fish with minute specimens as the latter may get accidentally swallowed. Do not to mix crabs and prawns with fish as the former may do considerable damage to the latter.
Never mix specimens from two different localities in the same bottle. Always use a fresh bottle for each site.