Sampling and preservation for reference
Mounting, preparation of skeletal material, other
Incomplete, unrevised draft. Help for improvement would be appreciated
Carcasses or remnants of dead animals, faeces and other biological material found in the wild may be very useful for obtaining data about wild populations with little disturbance of live animals or their habitat. It can no longer be regarded as ethical to kill threatened wild animals for obtaining skins for collections; carcasses of animals found or confiscated from poachers may serve as a better source of material for reference collections. Therefore, careful consideration which parts of a specimen have to be damaged or destroyed for examination, and preservation of which parts in reference collections is more useful (one of us: A. Nekaris; see also Groves, 2002 in press)
Specimens which may be useful in reference collections:
Skins; study skins, mounted specimens of the study species (the possibility of colour changes caused by preservation should be considered)
Wet preserved specimens
In lorises and galagos, morphology of external genitalia such as penis spines may be of some taxonomic importance
Skeletal material, skulls; if no dead animals are available and killing of specimens is supposed to be avoided, for instance very detailed casts of dentition of anesthetized specimens with dentists´ materials are possible. Owl pellets may be an interesting source of bones (one of us: C. Groves)
Hair samples; reference hair collection of sympatric species
Reference specimens of food items
Preservation of entire animals
Types of collection specimens of an entire animal:
For reference collections, mammals can be prepared as a variety of specimens. The condition of the specimen may determine possible ways to preserve it; if for instance decomposition of the skin has loosened the hair of a carcass so much that it can easily be pulled out or removed by rubbing (“slipping” fur), it will be very difficult or impossible to produce a study skin or mounted specimen.
The most usual types of specimens (based on Nagorsen and Peterson, 1980) are:
1) entire fluid-preserved animals (for studying anatomy and histology; fluid preservation may change the fur colour)
2) study skins with accompanying skulls / partial skeletons (some bones remain in the skin), for studying pelage colour, hair quality and moulting patterns,
3) mounted skins with accompanying partial or entire skeleton (some bones may remain in the skin, dependant on the method of preservation) or freeze-dried specimens,
4) entire skeletons, for instance for studying anatomy, geographic variation or for age determination (entire skeletons are poorly represented in collections, so Nagorsen and Peterson (1980) recommend preparation of at least one male and one female skeleton per species.
in the field
For preserving taxonomic material such as museum study specimens, different preservation methods should be considered. In the field, there may be limited access to materials and equipment necessary, so preliminary preservation with more simple methods may be necessary before final preparation as a permanent collection specimen.
Examples: procedures for preliminary preservation of a whole animal
Short-term strorage without preservation
freshly dead animals needed for mounting or skin preparation)
In a cold to moderate climate without refrigeration small animals may be stored in the shade for 4-5 hours.
After this period, in warmer climate sooner, the viscera will begin to decompose (Hangay, Dinkley 1985)
After weighing and measuring the animal and attaching an adequate label (see labelling), very small specimens (up to 100 g) can be fixed whole by submerging them in 10 % buffered formalin (tissue - formalin solution ratio of at least 1 : 12). the body cavity can be filled with formalin solution by injection until it is turgid and firm; some formalin may also be injected under the skin, into the body cavity, larger muscles and organs. If hypodermic needles are not available, the body cavity can be opened ventrally by making a slit instead, allowing the formalin to enter. Keeping the mouth open with a piece of wood or cotton may later allow examination of teeth. Then the whole body can be immersed in formalin, in the posture in which it is supposed to stay permanently because it will harden. The ratio of formalin to carcass must be at least 12 to 1 to assure a good fixation. Tissues can be left in buffered neutralized formalin for several months, but formalin hardens specimens; therefore, after fixation, longterm storage in alcohol may be better. After preservation the carcass should therefore be washed in water and transferred into ethanol for permanent storage, see below: longterm liquid preservation (Nagorsen, Peterson, 1980; Munson, 2000; Rabinowitz et al., 2000).
Equipment necessary: formalin, buffer, water, scalpel and / or hypodermic syringes, material for permanent labels, containers (not metal containers unless they are acid-proof lined, because corrosion of the metal would discolour the specimen) (Nagorsen, Peterson 1980).Formalin, however, has some disadvantages; for instance it discolours the fur, after a longish immersion, softens the bones (one of us: C. Groves) and prevents further examination for microbiology.
Preservation in alcohol:
After weighing, a whole animal can be preserved in a container of alcohol (70-90%). Removal of the intestine prior to storage of the animal in alcohol is recommended (Rabinowitz et al., 2000).
Preservation by cooling or freezing:
Removal of the skin with insulating fur before cooling or freezing may help to cool the carcass down more quickly (Schoon, lecture manuscript).
Freezing is not recommended if histological examination is planned (Wobeser and Spraker, 1980).
of soft tissue for longterm storage (according to
Ruhr-University, pers. comm.)
(see also below: problem of changes of fur colour which may be important in specimens preserved for taxonomic purposes).:
Formalin increasingly hardens tissue and may soften bones when its pH value is too low (under influence of light formic acid may accumulate from oxidation of formalin, so samples in formalin should best be stored in the dark, maybe in a refrigerator). So for longterm storage (museum specimens), after fixation a transfer into alcohol may be best. Storage in alcohol, however, may lead to shrinking and some hardening due to dehydration. For permanent liquid storage of specimens in alcohol, after fixation in 10% buffered formalin solution the specimen must be washed by keeping it in slowly flowing water for 24 hours (for instance in a box closed with gauze) for removal of formalin remnants. Then the specimen should be kept in distilled water for about 30 minutes (exchanging the water twice would be best). When the formalin is completely removed, the specimen can be transferred into 50 % alcohol for 30 minutes, then into 70% alcohol for some time. For longterm storage in a collection, a final transfer into 80% alcohol is recommended.
Fur and hair
Preparation of skins in the
After removing the skin from the animal, as much flesh as possible should be removed, but without damaging the skin with hair roots. Then the skin can be dried in the sun, or if necessary, high over a fire, either hung on a line or stretched between pegs. Salting the skin will speed the drying process and temporarily preserve the skin. Areas that still have flesh or fat should be salted thoroughly. Powdered borax can be put on the skin to further preserve it - alternatively cold ashes from a fire can be used. When the skin is nearly dry, it should be folded with the hairy sides together (Rabinowitz et al., 2000). But see below: possible colour changes of hair.
Laboratory preparation of skins earlier dried in the field
described by Downing (1945):
1) Relaxation of the dry skin by soaking it in lukewarm tap water, usually over night.
2) Brief washing of the relaxed skin with soap and water.
3) Rinsing of the skin in a degreasing agent such as varsol or carbon-tetrachloride; if greasy, the skins were allowed to stand in it for half an hour or so.
4) Drying of the skin in sawdust, using compressed air to assist the drying and to blow the sawdust out of the hair.
Colour changes caused by this method, particularly by soaking, see below
changes of hair during preservation of fur
Hair colour of live animals may differ from the colour of preserved specimen for several reasons. Some dyes like plant juice may cause a reddish hair colour, algae may cause a greenish hair colour in certain arboreal species, for instance in the sloth Bradypus or a bright green dorsal colour in Galagoides demidoff, which fades rapidly after the death of the animal (Sanderson 1940).
In addition, colour differences in series of mammal skins may be caused by preservation and storage methods rather than showing the occurrence of different colour types (red and grey varieties, random erythrism) in one species (Sanderson, 1940).
Use of chemicals may lead to colour changes. Formalin discolours the fur (one of us: C. Groves). Long immersion in solutions of alcohol, salt, alum or similar preservatives also alters colour (Downing, 1945). Sumner (1927) who cleaned fur with benzine or other chemical agents for better comparability mentions colour changes.
Drying methods in the field have some influence. Sanderson (1940) found that series of skins dried in bright sun in the field tended to turn reddish. Experiments with a maroon-coloured rat, Malacomys longipes, showed that furs became grey, dark brown or reddish-brown, according to whether they were dried in a closed container, in shade or bright sunlight. Series of Praomys, including examples of bright reddish and olive-grey varieties, could be evenly dried to a corresponding dull siena when submitted to reciprocal treatment. Drying over a fire by smoke may also change colours (Sanderson 1940).
Soaking of dried skins has a considerable effect. Downing, Cross and Prince at the Royal Ontario Museum of Zoology found a marked dichromatism in collectons of squirrel skins after different preservation: tawny olive skins had been made up in the field; skins which showed a dark tone and reddish coloration had been dried in the field and later relaxed by soaking in water and made up in the museum laboratory. Some tests with pieces of skin of a freshly killed squirrel showed that treatment with several dry preservatives (arsenic, alum, borax, salt) and subsequent drying in an electric oven at 60° Centigrade for 24 hours did not lead to colour changes, but soaking of further fresh and dried samples, with and without preserving chemicals added, in warm water led to color changes. After soaking originally tawny samples for one hour, a borax treated and salt treated sample had become hazel, an arsenic treated sample between tawny and russet, an alum treated sample between russet and hazel, and even a distilled water treated sample showed a perceptible change. Samples that had been soaked for longer periods showed further darkening and deepening of color. Arsenic treated and distilled water treated samples showed the least change, borax treated the most change; alum treated and salt treated samples were intermediate. Although the above preservatives increased the amount of change which took place, even soaking the skin in distilled water caused a marked alteration of the normal hair color. Examination of the museum collections of other species showed similar changes after soaking, although less severe than in sciuridae, besides reddening the yellow and buffy colors had become much deeper in tone and exhibited a cinnamon, pink or reddish cast. Changes were not evenly distributed over the body; certain parts changed more than others. Downing concludes that such changes in color may render specimens almost useless for taxonomic studies in which color is an essential character, that more adequate methods for cleaning and relaxing skins are necessary, possibly with minimizing of the time in which the skin is moist, and that such possibly colour-changing treatment must be noted on labels (Downing, 1945).
Later changes of colour of hair in museum specimens may occur due to fading because of exposal to light, old age, proximity of certain chemicals, radiators and other influences (Sanderson 1940).
Hairs, hair reference collections
Hair may have microscopically visible features allowing taxonomic identification. Hair may not only be collected from live animals or carcasses, but also for instance with hair tubes ( for smaller mammals: tubes of a width slightly larger than the study species with double-sided sticky tape stuck to the inside) or hair catchers (facilities with wire-brush-like structures; animals are encouraged to squeeze through), baited ore just attached where animals are likely to pass, left in the field for 1-2 weeks.
Comparison with taxonomically identified hair samples of sympatric species (either from an own hair reference collection or in museum collections) may allow identification of food / prey of the study species, of remnants of the study species in carnivore faeces or owl pellets or hairs for instance collected from nests or with hair catchers.
Hair samples for DNA analysis
Hair must be plucked (not cut) to include follicle cells. A minimum of 10-20 hairs should be obtained (AZA Prosimian Taxon Advisory Group, 2002). Bearder et al. (1996) recommend to collect especially the long guard hairs plucked from between the shoulder blades and hairs from scent glands. Loose hairs which can easily be removed by plugging may already be dead with little follicle cells with DNA left (one of us: Ch. Roos). To prevent contamination from human skin, use of clean gloves or an instrument for plugging is recommended. Samples should be stored in paper (not plastic) envelopes; no special preservation is necessary (AZA Prosimian Taxon Advisory Group, 2002).
Skeletal material, teeth
Samples for food analysis
If a carcass is found in the wild, collection of the content of the entire alimentary tract for food analysis may be useful. Examination of the stomach content alone may not be sufficient for this purpose; in galagos and pottos, after gum-eating it seems that gums are retained in the stomach only for few minutes, so usually no trace of gum is found there, but gum may be found in the caecum (Hladik 1979, partly quoting Charles-Dominique 1971 and 1974). Contents of the digestive tract can be preserved in 5% formalin or 30-40% alcohol (Rabinowitz et al., 2000), or the whole alimentary tract may be preserved in formalin, after injection of formalin into the stomach, for later analysis (one of us: A. Nekaris).
Reference material, slides for nutritional analysis from
contents of the gastro-intestinal tract
Preparation of slides for microscopic comparison with reference slides may be time-consuming, so preservation of the material in the field and later analysis may be more convenient (Nagorsen, Peterson, quoting Drodz 1975 and Williams 1962).
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In: Loris and potto conservation database: field methods
Last amendment: 7 November 2002
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