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Sampling and preservation for reference
collections
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)
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.
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Preservation
of specimens
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
(of
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)
Formalin preservation:
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).
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Fluid
preservation
of soft tissue for longterm storage (according to
A. Schlichting,
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.
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Fur and hair
Preparation of skins in the
field
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
as
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
Problem of
colour
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).
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Skeletal material, teeth
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Other items
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
faeces or
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).
In:
Loris and potto conservation database: field
methods
http://www.species.net/primates/loris. |
Last
amendment: 7 November 2002
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