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D f^% A A Life Sciences Miscellaneous Publications 
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Mammal Collectors' Manual 

D. W. Nagorsen and R. L. Peterson 



Digitized by the Internet Archive 

in 2012 with funding from 

Royal Ontario Museum 


d.w.nagorsen Mammal Collectors' Manual 


A Guide for Collecting, Documenting, 
and Preparing Mammal Specimens 
for Scientific Research 

Publication date: 6 June 1980 
ISBN 0-88854-255-0 
ISSN 0082-5093 







The Royal Ontario Museum publishes three series in the Life Sciences: 

life sciences contributions, a numbered series of original scientific publications including monographic 

life sciences occasional papers, a numbered series of original scientific publications, primarily short and 
usually of taxonomic significance. 

life sciences miscellaneous publications, an unnumbered series of publications of varied subject matter 
and format. 

All manuscripts considered for publication are subject to the scrutiny and editorial policies of the Life 
Sciences Editorial Board, and to review by persons outside the Museum staff who are authorities in the 
particular field involved. 


Senior Editor: R d james 
Editor: c McGOwan 
Editor: p h von bitter 

d w nagorsen is a Curatorial Assistant in the Department of Mammalogy, Royal Ontario Museum. 
R L Peterson is Curator-in-charge in the Department of Mammalogy, Royal Ontario Museum, and 
Professor in the Department of Zoology, University of Toronto. 

Cover: A hoary bat (Lasiurus cinereus) and a deer mouse (Peromyscus maniculatus) 

Canadian Cataloguing in Publication Data 

Nagorsen, David W. 

Mammal collectors' manual 

(Life sciences miscellaneous publications ISSN 

Bibliography: p. 

ISBN 0-88854-255-0 pa. 

1 . Mammals — Collection and preservation — 
Handbooks, manuals, etc. I. Peterson, Randolph 
L., 1920- II. Royal Ontario Museum. III. Title. 
IV. Series: Life sciences miscellaneous publication. 




© The Royal Ontario Museum, 1980 

100 Queen's Park, Toronto, Canada M5S 2C6 



1. Introduction 5 

2. Collecting Policy 6 

2.1 Collecting Laws 6 

2.2 Firearms 7 

2.3 Collecting Ethics 8 

3. Methods for Collecting Mammals 8 

3.1 Bats 8 

A. Equipment 8 

B. Collecting Techniques 11 

3.2 Other Small Mammals 12 

3.3 Fur-bearers and Large Mammals 16 

4. Documenting Specimens 17 

4.1 Recording Data 17 

4.2 Specimen Tags 19 

4.3 Measurements and Weights 22 

4.4 Determining the Sex of Mammals 26 

4.5 Reproductive Data 30 

A. Males 31 

B. Females 31 

4.6 Locality Descriptions 32 

4.7 Habitat Descriptions 34 

4.8 Methods of Capture 35 

4.9 Miscellaneous Field Notes 35 

4. 10 Photographic Records 35 

5. Methods for Preparing Specimens 36 

5.1 Preserving in Fluid 36 

5.2 Preparing Skins 38 

A. Study Skins 39 

B. Flat Skins 54 

C. Skins to be Tanned 57 

5.3 Preparing Skulls 60 

5.4 Preparing Skeletons 60 

6. Special Techniques 61 

6.1 Karyotyping 61 

6.2 Collecting Parasites 63 

6.3 Tissues for Biochemical Study 64 

6.4 Blood Samples 65 

6.5 Preserving Stomach Contents 66 

6.6 Preparation of Sperm Slides 66 

6.7 Fixing Tissues for Histological Study 67 

7. Shipping Specimens 68 

7. 1 Methods for Shipping. 68 

7.2 Import/Export Regulations 70 

8. Public Health Hazards 72 

9. Acknowledgements 72 

10. Selected Bibliography 73 

1 1 . Appendices 77 

11.1 Appendix 1 : Checklist of Field Equipment 77 

11.2 Appendix 2: Karyotype Kit for 120 Small Mammals 78 

Mammal Collectors' Manual 

1 . Introduction 

In 1965, one of the authors (R.L.P.) prepared a pamphlet Collecting Bat Specimens 
for Scientific Purposes, in which procedures were outlined for collecting, measuring, 
preparing, and shipping specimens. The usefulness of that leaflet prompted us to 
expand the concept into a more detailed book that includes most mammals. Our 
intention here is not to encourage indiscriminate collecting. Rather the purpose of the 
manual is to provide a guide that will serve as a set of standards for anyone collecting 
mammals for scientific specimens. By following the guidelines in this manual, 
specimens and associated data should have maximum value for research. In addition 
to museum biologists and mammalogy students, the guide should be useful to 
ecologists, biologists involved in environmental impact studies, parasitologists, 
cytogeneticists, and other laboratory scientists who may find it necessary to prepare 
voucher specimens for their research. Naturalists and wildlife biologists wanting to 
prepare museum specimens from rare or unusual mammals obtained from hunters, 
trappers, or road kills should also find the manual helpful. 

Several general guides already exist for the mammal collector. These include Hall 
(1962), Anderson (1965), Setzer (1968), Giles (1971), and DeBlase and Martin 
(1974). However, several of these publications are now out of date and do not 
describe the newer techniques available. Also, the kinds of field data that should be 
recorded may be given only cursory coverage. A number of specialized papers (see 
bibliography) dealing with collecting techniques for specific mammals or field data 
(reproductive data, locality descriptions) exist in various scientific journals. 
However, the average collector may be unaware of these publications. Therefore, we 
have attempted to produce an up-to-date guide on the methods for collecting, 
preparing, and documenting mammal specimens by combining methods derived from 
our own field experience and from the published sources listed in the bibliography. 

The recording of field data is strongly emphasized in the manual. In 1972 the 
Department of Mammalogy at the ROM initiated a computerized cataloguing system 
for storing and retrieving specimen data. With this system, it is now possible to 
process a large quantity of data for each specimen. In addition to standard 
measurements, information on weight, sex, age, date, reproductive data, habitat 
description, and a precise locality description, including latitude and longitude, are 
stored on magnetic tape. Other institutions with major mammal collections have also 
begun to use similar computer systems and it is possible that in the future most 
museum catalogue records may be stored in a central data bank. To utilize fully the 
potential of these cataloguing systems, collectors should provide the maximum 
amount of data for specimens. 

With an increasing concern for the conservation of mammalian species and the 
additional restrictions being placed on collectors, it is most important that a 
reasonable and responsible collecting policy be followed. Collecting ethics and 
collecting laws are discussed in section 2. The recent proliferation of import/export 
regulations for scientific specimens is another area of concern for collectors. 
Canadian and US import/export regulations are discussed in some detail in section 7. 
Collectors are urged to read these two sections carefully before collecting specimens. 

2. Collecting Policy 

2.1 Collecting Laws 

In recent years there has been a great increase in the number of collecting laws and 
endangered species acts that directly affect the scientific collector. These laws may be 
complex and ambiguous and obtaining the necessary permits for collecting in an area 
may involve considerable bureaucracy. Nevertheless, the collector has an obligation 
to learn and comply with these laws and regulations. It is essential that permits be 
obtained prior to any field collecting. Mammals that are protected under endangered 
species legislation should not be disturbed or collected except under special permit. 


In Canada, a Scientific Collector's Licence issued by the various provincial and 
territorial governments is needed. Fur-bearers and game species are usually regulated 
under provincial Game Acts and Regulations and special permission may be 
necessary for collecting these species. Additional permits may be needed to work in a 
provincial park. In Ontario, for example, collectors who plan to work in provincial 
parks must have their research proposal approved by the District Manager and the 
Director of the Parks Branch, Ministry of Natural Resources. Permission from the 
federal government is required for collecting in national parks. Some provinces and 
territories also require permits for salvaging dead mammals or parts thereof 
(carcasses, bones, shed antlers). For more information, consult the appropriate 
provincial or territorial governments. 

Although Canada has no federal endangered species act, Canada has signed the 
Convention on International Trade in Endangered Species (see section 7.2). Ontario 
and New Brunswick have passed provincial endangered species legislation. 

United States 

The US laws are complex and involve state and federal authorities. Generally, 
wildlife is regulated by the state governments and the collector should contact the 
appropriate state governmental agency for permits. To collect scientific specimens in 
a national refuge or in a national park, a permit issued by the Refuge Manager or the 

Superintendent of the national park is necessary. Permits to collect in national parks 
are issued only to persons officially representing reputable scientific institutions and 
annual reports are required. Although state and federal requirements for scientific 
collecting must be met, special permits are not needed for collecting in national forest 
systems. Collectors, however, are requested to contact the local Forest Service 
District Ranger before initiating any fieldwork. 

Marine mammals are covered under the Marine Mammal Protection Act. You may 
take and process marine mammals and parts thereof (bones, teeth, ivory) only under 
permit. Walruses (Qdobenus rosmarus), sea otters (Enhydra lutris), polar bears 
(Ursus maritimus), and manatees (Trichechus sp.) are under the jurisdiction of the 
Director of the US Fish and Wildlife Service. Cetaceans and all pinnipeds (except 
walruses) are under the authority of the Director of the National Marine Fisheries 

Collectors working in the US should be aware that endangered and threatened 
species are protected at both the federal and state levels. The federal Endangered 
Species Act that took effect in 1977 prohibits the taking and capture of all listed 
mammals as well as the import and export of these species. Species protected under 
the Act are listed as either "endangered" or "threatened". The various prohibitions 
of the Act apply to live or dead mammals and their parts or products. If you plan to 
salvage or utilize dead mammals listed in the Act, you must have a permit issued by 
the Director of the US Fish and Wildlife Service, or if endangered marine mammals, 
from the Director of National Marine Fisheries Service. For further information, 
contact the Federal Wildlife Permit Office, US Fish and Wildlife Service, 
Washington, DC 20240. Some states have also passed endangered species legislation 
and mammals protected under state law may or may not be the same as those listed 
under the federal Endangered Species Act. Information on these state laws can be 
obtained from the appropriate state governmental authorities such as conservation or 
wildlife departments (see McGaugh and Genoways 1976). 

Other Countries 

Many other countries have also passed endangered species legislation and regulations 
for the collecting of scientific specimens. Collectors planning to work abroad should 
contact the governmental agencies in these countries well in advance of any field trip. 
It is essential that one understand the regulations in these countries and obtain the 
necessary permits before any field collecting. Moreover, export permits may be 
required in some countries for transporting specimens out of these countries. 

The collector should also be aware of the Convention on International Trade in 
Endangered Species. Even if species listed in the Convention are not protected in the 
country where the collector is working, it may be impossible to import specimens of 
species listed in the Convention into North America without permits (see section 7). 

2.2 Firearms 

In recent years there has been a growing anti-hunting sentiment and collectors 
therefore are urged to use discretion when collecting with guns. Handguns are strictly 
controlled in Canada and permits issued by police departments are required. In some 
provinces municipal or provincial police departments may issue permits; in other 

provinces, the Yukon, and the Northwest Territories, permits are issued by the Royal 
Canadian Mounted Police (RCMP). In the US, firearms are generally regulated by the 
various state governments and collectors should consult their state or local police 
departments for information. When planning to use firearms in foreign countries, 
investigate thoroughly the various laws in these countries pertaining to firearms and 
ammunition. These laws include customs regulations covering the importation of 
guns and ammunition. 

2.3 Collecting Ethics 

The possession of a valid collecting permit does not give the collector the right to use 
irresponsible collecting methods. Specimens should be collected in the most humane 
method possible and any damage or destruction to the local biota or collecting sites 
must be prevented. Indiscriminate collecting of excessive numbers is discouraged, 
particularly in areas where large numbers may concentrate. When possible, take 
mammals alive and once the required sample has been collected, release the 
remaining mammals unharmed. Obviously with techniques such as snap trapping this 
procedure is impossible to follow. Specimens acquired for systematic collections 
should be carefully prepared and thoroughly documented using the standards outlined 
in this manual. 

Systematic research on a species requires a statistically adequate number of 
specimens from various localities. Because some species are sexually dimorphic 
(e.g., one sex may be consistently larger than the other), a representative series 
should contain samples of both sexes. For a given locality, 10 to 15 adults of each sex 
are usually an adequate number for a species. A few young animals in each sample 
may be useful for studying growth and age variation. 

To study geographic variation in a species, representative samples from various 
localities throughout the geographic range of the species are required. Distance 
between collecting sites is a function of habitat diversity in a given area and in regions 
with homogeneous biomes and habitats (e.g., the boreal forest in northern Canada or 
the tropical rain forest in the Amazon basin) localities may be 15 to 160 km (10-100 
miles) apart. But in regions that support a diversity of biomes and habitats (e.g., 
mountainous regions or river systems), collecting sites may need to be close together, 
8 km (5 miles) or less. 

3. Methods for Collecting Mammals 

3.1 Bats 

A. Equipment 

mist NETS 

Mist nets or bird-banders' nets (Bleitz Wildlife Foundation, 5334 Hollywood 
Boulevard, Hollywood, California, USA) are effective for capturing bats alive. The 


nets consist of a fine nylon mesh (50 or 70 denier) thread with mesh consisting of 
36 mm (1.5 inch) squares fitted on a string frame that divides the net into panels. 
Loops of cord at the end of each panel hold the net on supports such as long slender 
poles of bamboo cane (Fig. 1). 

In tropical regions a machete is useful to cut and trim suitable poles and to clear 
vegetation. To set the net, secure one pole in the ground. Usually the pole can be 
driven into the ground by repeated jabbing and twisting, however, you may have to 
provide additional support by piling rocks against the base of the pole or by attaching 
guys of cord to nearby trees or stakes. Place the loops of one end of the net in proper 
sequence over the pole and unfold the net, keeping it taut and off the ground. When 
the net is completely unfolded, erect the second pole and slip the loops over it. Once 
on the poles, the net is opened by spacing the panel strings at intervals along the pole. 
It is important that the net be taut enough to prevent sagging but loose enough to 
provide adequate pockets to stop bats from bouncing off the net. Remove any leaves, 
sticks, or insects that may become entangled in the net. Although the poles and net 
should be set and adjusted before dark, the net should not be opened on the poles until 
the collector intends to use it. This will prevent the capture of birds that are frequently 
active several hours before sunset. It is important to check nets frequently and 
regularly and to remove netted bats, as large bats can cause extensive damage by 
chewing themselves free. To catch species that are adept at freeing themselves 
without entanglement, it is necessary to actually stand by the net and remove bats as 
they hit the net. This is particularly true for some of the small, African free-tailed bats 
(Molossidae). Constant attention is also necessary in agricultural areas where 
domestic animals may wander into untended mist nets. 

Equipment required to attend nets includes: a headlight with spare batteries and 
bulbs, a flashlight, collecting bags, gloves, nylon fishing line to repair broken shelf 

Fig. 1 A four-panel mist net (12.5 m x 2 m; 42 ft x 7 ft) set on poles. 

strings, cord or rope for guy lines, and a knife or scissors for cutting badly entangled 
bats from the net. 

To remove a bat from the net, determine first the side from which it entered. With 
the bat held firmly in the gloved hand, use the ungloved hand to remove the bat from 
the open side of the pocket, beginning with the head. The best method is to remove 
netting from the bat's mouth first and then work back, freeing the wings and feet. 
Once removed from the net, bats can be kept alive in cloth collecting bags about 
35 cm x 20 cm (14 inches x 18 inches). Use several bags and keep the larger species 
separate from the smaller ones. If bats are not prepared immediately as specimens, 
they can be kept alive overnight in these bags. To increase the chance of survival, 
keep the bags in a cool, well-ventilated place. 

Nets should be taken down before dawn to avoid catching birds and diurnal insects. 
First, remove all leaves or sticks, then slide the string loops together on the poles to 
close the net. In areas where theft is not a problem, nets may be closed and left on the 
poles during the day. When the net is to be removed from the poles, tie a piece of 
string to the top loop of each end and pass this string through all the loops to keep 
them in proper sequence. Then remove the net from one pole and walk towards the 
other pole collecting and folding the net in your hands. The net can then be doubled or 
redoubled into a compact bundle and stored in a plastic bag. 


An insect net with a long, extensible aluminium handle (no. 324 Tropics net, 
BioQuip Products, P.O. Box 681, Santa Monica, California, USA) is useful to 
collect bats in caves, mines, and buildings. This particular net has a 4 m (12 ft) 
handle composed of six pieces, each 60 cm (2 ft) in length, which screw together. An 
ordinary insect net can be modified for bat collecting by making a long handle of 
suitable material. You can also improvise with a coat hanger or a wire of similar 
gauge bent into a hoop and laced with mosquito netting or cheesecloth. 


Although less preferable to netting because of potential specimen damage, shooting is 
a useful technique for collecting some of the sac-winged (Emballonuridae), 
vespertilionid (Vespertilionidae), and free-tailed (Molossidae) bats that are difficult 
to net because they forage above the tree canopy and for bats that roost in large caves 
with high ceilings or in tall palm trees. For minimum damage to specimens, use fine 
shot (no. 12 shot) loaded in .22 or .32 calibre rifle shells or in .410 gauge shotgun 
shells. For best results .22 rifle shells loaded with no. 12 shot should be used in a 
special smooth bore .22 gun. Auxiliary barrels ("Aux.") that slip into 20, 16, or 12 
gauge shotgun chambers are made for holding .32 gauge or .410 gauge shells loaded 
with no. 12 shot. 


Several designs for bat traps have been used but one of the most versatile is the Tuttle 
trap. This trap consists of two rectangular aluminium frames that support vertical wire 
strands. Bats collide with the wires and fall unharmed into a canvas collecting bag. 
The Tuttle trap is not available commercially; however, for a description, including 
specifications for construction, see Tuttle (1974). In arid regions with restricted 


water, a single strand of fine piano wire can be stretched across a pond or water tank 
about 20 cm (6 inches) above the water. Bats striking the wire will fall into the water. 
As bats swim to shore, they can be captured with a hand net. 

B. Collecting Techniques 

Bats are collected primarily from diurnal roosts and from foraging or drinking sites at 


Caves and Mines 

Caves and mines (especially abandoned ones) are usually productive sites for the bat 
collector. Larger caves or mines that may contain bats may be located from 
large-scale topographic maps (1:50 000), which usually indicate cave or mine sites, 
and by questioning local residents. The collector may save considerable time and 
effort by hiring a local person familiar with a particular mine or cave to act as a guide. 
The cave or mine should be entered slowly with a minimal amount of noise. Some 
species roost in dim areas near the entrance; others prefer dark areas deep within. 
Examine each hole or depression in the roof as well as side openings while listening 
for vocalizations or for sounds of flying bats. Bats roosting on the ceiling may be 
taken in a long-handled insect net or shot — if there appears to be no danger of the 
ceiling's collapsing. Bats flying about can be captured in bat traps or with small 
pieces (1 m; 2-4 ft) of old mist net strung across corridors. Mist nets set near the cave 
or mine entrance before dusk may capture large numbers of bats as they leave to 


Many bats, particularly some vespertilionids (Vespertilionidae) and free-tailed bats 
(Molossidae) roost in tile, thatched, or metal roofs, attics, and cavities between walls 
of buildings. Local inhabitants can usually provide information on buildings that 
contain bat colonies. Bats roosting in buildings may often be captured by hand (use 
gloves), with hand nets, or with long forceps (25 cm; 10 inches) that will reach into 
holes and crevices. If you cannot capture bats from buildings during the day, it may 
be possible to collect them with mist nets, hand nets, or bat traps as they leave the 
building to forage at night. By carefully observing the building in the evening, you 
can usually locate the openings that bats are using for exits. 

Other Roosts 

During the day, some bats roost in hollow trees or logs, under the bark of trees, in 
rock crevices, under large leaves such as palm or banana fronds, in culverts, under 
bridges, and even under rocks or stones. Migrating tree bats and Old World fruit bats 
may hang from trees and bushes in open, exposed areas. 


Although foraging bats can be collected with traps or by shooting, generally the most 


effective technique is mist netting. To net foraging bats efficiently, the collector 
should become familiar with the most productive areas. Many species can be netted 
near their feeding sites (orchards, wild fruit trees, flowering shrubs, trees, and over 
ponds or streams). Forest trails, the edges of forested areas, and highland passes are 
often used by bats as natural fly ways. Isolated ponds in arid regions may attract great 
numbers of bats, particularly during the dry season. Nets set over streams or forest 
trails should be positioned in narrow areas where natural obstacles funnel bats into the 
net. Factors that will reduce netting success are rain, heavy dew, and moonlight. 
Some experimenting with the height of the net above the ground, the angle of the net 
relative to a flyway and its position relative to surrounding vegetation is necessary to 
obtain satisfactory results in different localities. Many species fly just above the 
ground vegetation so that nets set with the bottom strand about 20 cm (6 inches) 
above the ground is a good starting point from which to experiment. 

3.2 Other Small Mammals 

Snap Traps 

The most successful trap for catching small rodents, marsupials, and insectivores is 
the snap trap. These traps are generally sold commercially (Victor Traps, Litiz, 
Pennsylvania, USA) in mouse-trap and rat-trap sizes. The larger, more powerful 
rat-trap is designed for killing mammals the size of rats and small squirrels. Designed 
for mammals of shrew and mouse size, the smaller mouse-trap is more effective, but 
the spring bar on the trap frequently crushes the skull of the specimen. 

Most collectors prefer the Museum Special model (Fig. 2) (Woodstream 
Corporation, Litiz, Pennsylvania, USA). This trap is intermediate in size between the 

Fig. 2 Museum special snap trap shown in the set position. Trap size is 14 cm x 7 cm (5.5 
inches x 2.7 inches). 


mouse- and rat-trap and it is designed to kill mammals weighing up to 50 g (2 oz). 
Advantages of this trap are an extremely sensitive trigger mechanism and a spring bar 
designed to break the specimen's back rather than to crush the skull. Equipped with a 
weak spring, this trap can be used to catch small mammals such as shrews without 
seriously damaging the specimen. Snap traps should be checked at least once every 
24 hours, preferably in the early morning. Dead mammals decompose rapidly, 
especially in warm weather. In tropical areas you may have to check traps more 
frequently as ants may quickly eat specimens. 

Live Traps 

Live traps are used to obtain live mammals for karyotyping, biochemical analyses, 
and parasite studies. Also, live traps permit the collector to select only those 
mammals required for specimens and release others unharmed. As some animals 
(e.g., shrews) may be reluctant to enter, live traps are generally not as effective as 
snap traps. To sample the mammalian fauna of an area accurately, you should 
supplement live trap lines with some snap traps. 

For mammals such as squirrels, hares, and foxes welded wire mesh traps are 
available in several sizes (Havahart Company, P.O. Box 551, Ossining, New York, 
USA; National Trap Corporation, P.O. Box 302, Tomahawk, Wisconsin, USA). 
These traps usually have a front and rear door and are activated by a bait pan in the 
centre of the trap. 

For small rodents and some insectivores the most popular live traps are the 
Longworth (Longworth Scientific Instrument Company Limited, Thames Street, 
Abingdon, Berkshire, England) and the Sherman trap (H. B. Sherman Company, 
Box 683, DeLand, Florida, USA). The aluminium Longworth trap has a trigger 
mechanism with a detachable nest box. The Sherman style trap, a rectangular box 
constructed from aluminium or galvanized metal, has a spring-loaded treadle which 
releases the door when depressed. An assortment of sizes and models, including 
folding and nonfolding are available. We have found a modification of the Sherman 
trap (Canadian Penitentiary Industries, Sir Wilfrid Laurier Building, 340 Laurier 
Avenue West, Ottawa, Ontario, Canada K1A 0P9) to be effective in catching 
mammals ranging in size from shrews to small squirrels. The trap is a nonfolding type 
with a screened rear door (Fig. 3). 

To prevent deaths in live traps, check them at least once daily, preferably early in 
the morning. Also place some cotton wool for nesting material inside traps during 
cool weather. 


An effective bait for small mammals that can be used in snap traps or live traps is a 
mixture of peanut butter and rolled oats. Chopped nuts, seeds, bits of chopped fruit 
(apples, raisins, bananas), or cheese can be added to this mixture. Experiment with 
different combinations of bait to determine the one most effective in a particular area. 
Plastic containers with screw-top lids or plastic squeeze tube containers are useful for 
carrying premixed bait when checking traps. 



Fig. 3 Nonfolding type of Sherman live trap with screened rear door. Trap size is 30 cm x 
8 cm x 8 cm (12 inches x 3 inches x 3 inches). 

Special Traps 

Special traps are made (Victor Traps, Litiz, Pennsylvania, USA; Z. A. MacAbee 
Gopher Trap Company, 110 Loma Alta Avenue, Los Gatos, California, USA) for 
capturing such fossorial mammals as pocket gophers (Geomyidae) and moles 
(Talpidae). Gopher traps (Fig. 4) are set in tunnels that are 20 to 900 cm (6-36 
inches) below the surface of the ground. Locate the shallow tunnels near freshly 
discharged earth, remove a section of the tunnel, and set a pair of traps in each 
direction in the runway. When excavating earth to repair the opened tunnel, the 
pocket gopher will set off the trigger mechanism of the trap. To prevent traps from 
being dragged into the burrow system, tie them with wire to a firm stake. 

Of the various mole traps sold commercially, the harpoon type (Fig. 5) appears to 
be the most efficient. Push the trap into the ground over the mole tunnel and flatten 
the raised tunnel so that the trigger pan will be set off by the mole as it moves through 
the tunnel. Consult Baker and Williams (1972) for a description of a live trap for 
gophers and Yates and Schmidly (1975) for a mole live trap. 

A simple, effective trap for shrews and certain species of rodents (e.g., jumping 
mice and microtine rodents) is a fruit juice or similar-sized can 35 cm x 18 cm (14 
inches x 7 inches) set flush with the ground. Cone-shaped pitfall traps, which have 
the advantage of being easy to carry in the field because they nest together, are sold 
commercially (Northwest Metal Products Company, P.O. Box 10, Kent, 
Washington, USA). It is not necessary to bait pitfall traps as the mammal simply 


Fig. 4 Pocket gopher trap in the set position. Trap size is 15 cm x 5 cm (6 inches x 2 

tumbles into them. About 75 mm (3 inches) of water in the trap will prevent rodents 
from jumping out. Set them adjacent to streams, in runways, or near burrows for best 
results. When sampling a habitat, set at least a few pitfall traps for they frequently 
capture species not taken in other types of traps. 

Operating A Trap Line 

To sample the small mammals of an area thoroughly, traps should be set in a variety 
of habitats (forest, open grassy area, transition zones). The recommended procedure 
is to set the traps in a "trap line" at regular intervals and roughly in a straight line. 
Trap sites are marked by tying a piece of coloured, plastic flagging tape or strip of 
cloth to a tree branch or a clump of vegetation. The total number of traps in a line 
(usually 30-100 traps), the number of traps at each site, and the spacing of traps is 
determined by experience. For most habitats, you will obtain good results by setting 
traps about 10 m (30 ft) apart with two or three traps at each site. To be certain that no 
trap sites are missed when checking a trap line, many collectors number their traps in 
sequence. Permanent numbers can be painted on live traps and numbers can be 
written with a pencil on wooden-based snap traps. Another method is to write trap site 
numbers on plastic flagging tape with a felt-tipped marker pen (waterproof ink type). 

Rather than randomly selecting trapping sites, carefully choose the most favourable 
microhabitat, for example, the base of trees or stumps, on top of logs, in conspicuous 
runways, at burrow entrances, or at the edge of streams or ponds. Many small 
mammals confine their movements to runways that appear as well trampled miniature 
trails in the vegetation. Other signs indicating the presence of small mammals are 
droppings, tracks, piles of cut grass or sedges, and seed caches. In tropical forests 


Fig. 5 Harpoon type of mole trap. 
Trap length is 40 cm (16 inches). 

some species of small mammals are arboreal and traps set in the branches of trees will 
catch species not otherwise taken. 


For mammals such as rabbits, squirrels, and small- to medium-sized carnivores, 
shooting may be more effective and humane than trapping. To prevent excessive 
damage, use a small gauge shotgun (.410 or 20 gauge) loaded with light shot instead 
of a .22 rifle. BB shot or no. 2 shot is recommended for fox-sized animals, shot no. 4, 
6 or 7'/2 is suitable for hare-sized, and no. 1 1 or 12 shot is used for smaller mammals 
(pikas, squirrels). Auxiliary barrels are discussed in section 3.1 

3.3 Fur-bearers and Large Mammals 

Steel leg-hold traps, Conibear traps, snares, and shooting are the usual methods for 
collecting these mammals but check local laws and regulations before doing so. 
Arrangements with experienced fur trappers to secure specimens taken in season 
often provide good results. For reviews of the techniques used for trapping 
fur-bearers, consult The Manitoba Trappers' Guide (Manitoba Department of 
Renewable Resources and Transportation Service 1965) or Stains (1962). 


4. Documenting Specimens 

Described in this section are basic field data that should be routinely recorded by 
anyone collecting mammals for scientific specimens (field number, date collected, 
nature of specimens, measurements, sex, reproductive data, locality descriptions, 
habitat descriptions, method of capture, and miscellaneous field notes). However, it 
cannot be over-emphasized that this information must be accurate. If you are not 
certain, do not guess! If you are uncertain of the sex, for example, then indicate so 
with a question mark on your catalogue or field notes. 

4.1 Recording Data 

Collectors usually record their field data in a notebook. DeBlase and Martin (1974) 
and Hall (1962) recommended that field notes be organized into three sections: (1) a 
journal; (2) a catalogue; (3) species accounts. At the ROM we use a slightly different 
system. Specimen data are recorded on printed catalogue sheets (Fig. 6). Similar or 
modified sheets can be designed to suit your particular needs. The following is a brief 
explanation of headings of columns shown on the catalogue sheet in Figure 6. 

The museum number is assigned to the specimen when it is accessioned by the 
museum, consequently leave this space blank. Each specimen listed in the catalogue 
must have a separate field number and this number is also written on a tag (in pencil 
or waterproof ink) that is securely tied to the corresponding specimen (see section 
4.2). A tentative identification, even a common name of the specimen, should be 
entered under the species heading. Always write the date with the month in full, that 
is, June 10, 1965 not 10/6/65. Documenting the categories for sex, measurements, 
and locality are discussed in sections 4.4, 4.3, and 4.6. The nature of the specimen 
(skin and skull, skull only, skin only, skeleton only or preserved in formalin), should 
also be listed in the remarks for each specimen. 

Such information as reproductive data, habitat, and field observations (see sections 
4.5, 4.7, and 4.9) may be entered in the remarks section; however, usually it is 
impossible to fit all of this information on the catalogue sheet. We recommend that a 
field notebook or diary be kept in conjunction with the catalogue sheets. To ensure 
that data are associated with the appropriate specimens, carefully list the field 
numbers with their corresponding data. Figure 7 illustrates a page taken from a 
typical field notebook. At the ROM we also use printed sheets for recording 
reproductive data (Fig. 8). You may wish to design similar sheets or simply record 
these data in your notebook. 

All catalogues, field notes, photographs, and maps of collecting sites are kept 
permanently by museums as documentation for researchers. As catalogue sheets and 
field notes are often the only source of information for specimens, your catalogue and 
notes should be well organized, legible, and as accurate as possible. Write in pencil 
or waterproof ink as other types of ink will run if subjected to moisture. Catalogue 
sheets or notes that have been seriously damaged from moisture, grease, or blood 
should be recopied. 








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Fig. 7 A page taken from a field notebook of a ROM collector to illustrate the data recorded. 
Specimens are the same as those listed in the catalogue sheet (Fig. 6). 

4.2 Specimen Tags 

All specimens listed in your catalogue should be labelled with their corresponding 
field numbers. DeBlase and Martin (1974) described some of the different kinds of 
tags used by field collectors. ROM collectors use printed field number tags (Fig. 9d) 
that are resistant to alcohol and formalin to label study skins, skulls, skeletons, or 








































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specimens in fluid. If these are not available, you can construct similar tags from 
vinyl-coated paper or white cardboard (Fig. 9c). When labelling fluid-preserved 
specimens, ensure that tags are resistant to preserving solutions. A recommended 
method for coding field numbers is to write your initials preceding the specimen 
number (e.g., JHW 237). This practice will prevent any possible confusion with 
specimens of another collector. 





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Fig. 9 Three types of specimen tags and method for tying. 

A Knot used for stringing study skin tag. 

B Study skin label used by many field collectors. Information recorded includes 
locality, date, sex, collector's name, measurements, reproductive data, identifica- 
tion of specimen, and field number. 

c Label constructed from vinyl-coated paper; field number consists of collector's 
initials and specimen number. 

D Paper label (alcohol and formalin resistant) with printed field number. 


For study skins and entire specimens in fluid, some collectors use a label similar to 
that illustrated in Figure 9b. Information recorded on these labels includes locality, 
date, sex, collector's name, identification of specimen, measurements, and 
reproductive data. Standard measurements (see section 4.3) are always listed in the 
following sequence: (1) total length; (2) tail vertebrae length; (3) hind foot length; (4) 
ear length; and in bats (5) tragus length, and (6) forearm length. At the ROM these 
labels are now generated by our computerized cataloguing system. As a result, we do 
not fill out these labels in the field and study skins and fluid-preserved specimens are 
labelled with field number tags (Fig. 9d). 

To prepare the tag for tying, run a thread through two holes near one end of the tag 
and tie a knot about 2.5 cm (1 inch) from the tag (Fig. 9a). You can save 
considerable time in the field by stringing your tags with thread before you take them 
into the field. 

Tie tags on study skins and specimens in fluid above the ankle with a square knot 
(Fig. 37). For flat skins, also write the field number on the cardboard stretcher with 
waterproof ink. If printed field tags are used, they should be tied directly to the 
cardboard (Fig. 41). Some collectors (Anderson, 1965) record such data as sex, 
measurements, locality, and date on the card. For "cased" or "open" skins from 
larger mammals that are to be tanned, attach the field number through the nostrils. 

If skulls or skeletons are stored in individual gauze bags for drying, tie a tag around 
the neck of the bag (Fig. 39b). If skulls or skeletons are not placed in individual bags, 
attach the tag directly to the specimen. Skull labels can be tied around the mandible or 
in larger mammals to the zygomatic arch. A suitable area for tying labels to 
articulated skeletons is the pelvis. For disarticulated skeletons, label each portion 
with a tag. If large skeletons are placed in burlap bags, you can also attach a tag to the 
outside of the bag. 

4.3 Measurements and Weights 

Mammalogists rely on weights and body measurements for aid in identifying 
specimens, determining the age of specimens, and for studying variation between 
different populations. It is essential that the collector record accurate measurements 
and weights before the specimen is prepared as a study skin or preserved in fluid. 
Study skins shrink somewhat during their preparation and reliable measurements 
cannot be made from the finished skin. Fluid-preserved specimens become stiff and 
inflexible once they have set in the fixative and are difficult to measure accurately. 
Measurements used in research papers and descriptions of species are always given in 
metric units. Linear measurements should be in millimetres and weights in grams or 
kilograms. Indicate an approximate measurement by the circa abbreviation "ca", 
e.g., ca 180 mm. Also note any aberrant measurements resulting from damaged 
specimens (tail broken, ear torn). 


The following are the standard measurements taken by North American collectors 
(Fig. 10). European collectors frequently record head and body length (HB) rather 
than total length and omit the claw for the hind foot measurement. For the sake of 



Fig. 10 Standard body measurements for bats. Total length (TL), forearm length (FA), hind 
foot length (HF), tragus length (TR), ear length (E), tail vertebrae length (TV), and 
wingspan (WS). For other small mammals, only the total length, tail vertebrae, hind 
foot, and ear are recorded. 


consistency, we recommend that collectors use the North American measurements. 

Total Length (TL): straight-line distance from the tip of the nose to the end of the 
last tail vertebra, exclusive of the hairs. Lay the mammal on its back on the ruler and 
measure by extending the specimen, pressing the body flat. Pull the tail (if present) to 
its full length, measuring to the end of the last bone. If no tail is present, measure to 
the end of the backbone. 

Tail Vertebrae (TV): distance from the base of the tail to the tip of the last vertebra, 
exclusive of the hairs. With the mammal on its belly, place the ruler at the point 
where the tail joins the body, pull the tail upward and measure to the end of the last 

Hind Foot (HF): distance from the end of the heel bone (calcaneum) to the end of 
the claw on the longest toe. Stretch the toes and measure from the heel to the longest 
length of the claws. For hoofed mammals, the HF measurement is taken from the tip 
of the hock to the tip of the hoof (Fig. 1 1). 

Ear Length (E): distance from the base of the notch of the lower part of the ear to 
the uppermost margin of the ear. 

Measurements limited to bats are: 

Tragus (TR): distance from where the tragus joins the ear to its tip. 

Forearm (FA): distance from the outside of the wrist to the outside of the elbow. 
Fold the wing when taking this measurement. 

Wingspan (WS): distance between the wing tips when the wing is stretched out. 
Lay the bat on its belly and gently stretch the wings, being careful not to overstretch 

Measure large mammals on level ground using a steel tape measure. The same 

Fig. 1 1 Standard body measurements for a large ungulate. Total length (TL), tail length 
(TV), hind foot (HF), ear from notch (E), and height at shoulder (HS). 


standard measurements (TL, TV, HF, E) are taken for these mammals (Fig. 11). An 
additional measurement is the height of the shoulder (HS) which is the distance from 
the top of the shoulder to the bottom of the foot. For total length, measure in a straight 
line with the body stretched out rather than measuring around the curves of the neck 
and back. 

The measurements for cetaceans recommended by the American Society of 
Mammalogists are shown in Figure 12. 


Specimens should be weighed promptly and before preparing. Weights should be 
taken in grams or kilograms; however, body weights in pounds or ounces are helpful 
when metric scales are not available. For approximate weights, use the abbreviation 
"ca". The body weight of a large mammal is difficult to obtain in the field; 

Fig. 12 Standard cetacean measurements as recommended by the American Society of 
Mammalogists. LENGTH: 1 total, 2 tip of upper jaw to centre of eye, 3 tip of upper 
jaw to apex of melon boss, 4 gape, 5 tip of upper jaw to external auditory meatus, 6 
centre of eye to external auditory meatus, 7 tip of upper jaw to blowhole along 
midline or to midlength of two blowholes, 8 tip of upper jaw to anterior insertion of 
flipper, 9 tip of upper jaw to tip of dorsal fin, 10 tip of upper jaw to midpoint of 
umbilicus, 11 tip of upper jaw to midpoint of genital aperture, 12 tip of upper jaw to 
centre of anus, 13 anterior insertion of flipper to tip, 14 axilla to tip of flipper, 15 
dorsal fin base, 16 distance from nearest point on anterior border of flukes to notch. 
WIDTH: 17 nipper (maximum), 18 flukes (tip to tip). HEIGHT: 19 dorsal fin (fin tip 
to base). GIRTH: 20 on a transverse plane intersecting axilla, 21 maxima, 22 on a 
transverse plane intersecting the anus. 


nevertheless, collectors should weigh large mammals whenever possible because of 
the paucity of weight data. 

High quality spring balances graduated in grams, for example, Pesola scales 
(Bleitz Wildlife Foundation, 5334 Hollywood Boulevard, Hollywood, California, 
USA) are excellent for the field collector. They are made in the following capacities: 
5 g, 10 g, 30 g, 100 g, 500 g, 1000 g, and 2500 g. An Ohaus triple beam balance 
(Fisher Scientific Company, 711 Forbes Avenue, Pittsburgh, Pennsylvania, USA) 
may also be used for weighing mammals to 2000 g. Although more accurate than 
Pesola balances, the triple beam balance is heavier and less compact. For mammals 
greater than 2500 g use a heavy-duty spring balance (Forestry Supplier Incorporated, 
Cox 8397/205 West Rankin Street, Jackson, Mississippi, USA). 

4.4 Determining the Sex of Mammals 

The sex of each specimen should be accurately determined. However, if there is 
doubt, so indicate in your Field notes or catalogue. 

External Genitalia 

The external genitalia of the male usually can be distinguished from those of the 
female by larger size and their position relative to the anus. Most males have a 
prominent penis (Fig. 13), but some small mammals, particularly shrews, have the 
penis retracted into a sheath of a tubular fold of skin during the intervals between the 
breeding seasons. Consequently, the external genitalia may appear to be superficially 
similar in both sexes. With fine pointed forceps, it is usually possible to protrude the 
penis from its sheath. A hand lens is often useful for examining the genitalia in small 
mammals. Many species have a bony or cartilaginous structure in the penis, the 
baculum (os penis). 

In most adult males, the testes occur outside the abdominal cavity, but in a few 
mammals (whales and dolphins) the testes remain permanently within the abdominal 
cavity. When the testes occur outside the abdomen, they are usually situated in a 
scrotum (Fig. 13a). Testes may remain permanently in the scrotum (most primates, 
dogs, ungulates) or they may be intra-abdominal during the nonbreeding season (most 
bats, some rodents). In some mammals (shrews, moles, some rodents, hares), the 
testes are not contained in a distinct scrotum and, although they are located outside 
the abdominal cavity, they remain under the integument in the inguinal region. 

The external genitalia of females consist of a vaginal opening (vulva) that may 
have prominent skin folds in some species. The urethra may also be visible (Fig. 13 
C,D). During the breeding season, teats or nipples of the female mammary glands may 
be enlarged. The number and position of the mammary glands vary greatly in 
different species. In some mammals (primates and bats) there is only a single pair of 
mammary glands confined to the chest region. However, in mammals with numerous 
glands (rodents) the teats are usually situated in parallel rows along the ventral surface 
of the chest and abdomen (Fig. 13E). 

In cetaceans (whales, dolphins, and porpoises), the genitalia are contained in a 
genital groove (Fig. 14). Normally the penis is retracted fully into a pouch in the 
ventral abdominal wall and only the genital slit can be seen. The penis protrudes from 


Fig. 13 External genitalia of a male and female cricetid rodent: a scrotum, b penis, c anus, 
d teats, e vulva, f vaginal opening (perforate), g vaginal opening (imperforate), 
h mammary tissue (stippled area). 

A An adult male with an enlarged scrotum that partially obscures the anus. 
B An immature male without an enlarged scrotum. 
C Vulva, anus, and a perforate vagina on a pregnant female. 
D Schematic drawing of a female showing a membrane covering the vaginal 

opening (imperforate condition). 
E Schematic drawing showing the position of mammary tissue under the teats. 


Fig. 14 Schematic drawing to show the external genitalia of a porpoise (cetacean): 
a umbilicus, b genital slit, c mammary slit, d anus, e penis. 
A Female porpoise illustrating the presence of mammary slits and the position of 

the genital slit. 
B Male porpoise with penis retracted into abdomen. Note position of the genital slit 

relative to the anus. 
C Male porpoise with penis protruding from the genital slit. 


the genital slit only during erection or occasionally on death. The female genitalia are 
also contained in a genital slit but the distance from the centre of the genital slit to the 
anus is much less in females than in males. In some porpoises, the anal and genital 
openings may occupy the same aperture. A number of accessory grooves may flank 
the female genital slit, but one pair — the mammary slits or grooves which contain the 
nipples — is always present and in a constant position in all species of cetaceans. The 
mammary slits are not present in male cetaceans. 

Internal Reproductive Organs 

After the skin has been removed, specimens to be prepared as study skins or skeletons 
should be dissected to verify the sex and to obtain reproductive data (see section 4.5). 
Testes appear as whitish or yellowish oval organs (Fig. 15). During intervals between 
the breeding season, testes may be small, especially in small mammals (shrews). 
Females can be distinguished by the presence of a uterus (Fig. 16). 

Fig. 15 Schematic drawing of the male urogenital system of the Norway rat {Rutins 
norvegicus) as an example of a typical rodent: a kidney, b ureter, c prostate gland, 
d urinary bladder, e vas deferens, f testis, g cauda epididymis, h penis. 


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Fig. 16 Schematic drawing of the female urogenital system of the Norway rat (Raitus 
norvegicus) as an example of a typical rodent: a kidney, b ureter, c ovary, d uterus 
(left horn), e urinary bladder, f vagina, g urethra, h vaginal orifice, i anus. 

4.5 Reproductive Data 

Notes on the condition of the reproductive organs provide important biological data. 
The length and time of the year of the breeding season, litter size, numbers of litters 
per year, and the age of sexual maturity may often be determined for a particular 
species in a given geographic area from these data. 

Mammals prepared as study skins or skeletons (sections 5.2 and 5.4) should be 
dissected immediately after the skin is removed. Open the body cavity by cutting the 
abdominal muscles with scissors or a scalpel and examine the reproductive organs. 
Fluid-preserved specimens (section 5.1) are usually not dissected in the field; 
nevertheless, important information on the breeding condition of the specimen can be 
provided by examining the external genitalia. If the condition of the reproductive 
organs cannot be determined because of decomposition, then indicate so in your field 


notes. Do not guess at the reproductive status! If you have difficulty distinguishing 
embryos or placental scars (see page 32), then preserve the uterus in a vial of 10 per 
cent buffered neutral formalin (with corresponding field number) and include it with 
your collection of specimens. 

A. Males 

Criteria used to distinguish breeding males are the size of the testes and the size of the 
tubules in the cauda epididymis. For dissected specimens, measure the length and 
width of the testes (both if different in size) in millimetres (e.g., 10 mm x 6 mm). 
For fluid-preserved specimens that have a scrotum (section 4.4), determine whether 
or not the testes are grossly enlarged. 

Another useful criteria for breeding males in various small species (shrews, 
rodents, bats) is the occurrence of visible tubules in the cauda epididymis (Fig. 15). If 
the tubules are visible to the eye, they are swollen and usually contain sperm. 
However, if the tubules are not visible, they are probably void of sperm. For 
dissected specimens, note whether or not the tubules are visible and record the 
condition in your field notes. 

B. Females 

Breeding females may be pregnant, lactating, or both. Criteria used to diagnose the 
breeding condition of females are the condition of the vagina, presence or absence of 
embryos, presence or absence of placental scars, and the condition of mammary 


The vagina of females is usually sealed by a membrane until puberty. In some rodents 
and moles (Talpidae) that breed seasonally, the vagina may be closed by a membrane 
during the nonbreeding (anoestrus) of the reproductive cycle (Fig. 13d). The vagina 
in this condition is described as imperforate. In contrast, during heat (oestrus), the 
vagina is not sealed by a membrane and is perforate (Fig. 13c). In some mammals, 
for example the squirrels (Sciuridae), the vaginal region (vulva) may appear to be 
swollen or turgid during oestrus. The condition of the vagina is usually a reliable 
indication of reproductive condition, so note whether the vagina is imperforate or 
perforate and also whether the vulva is obviously swollen. 


Lactation is defined as the secretion of milk. The following criteria are used as 
evidence that a female is lactating: (1) female is observed nursing young; (2) milk can 
be squeezed from teats; (3) heavy deposits of mammary tissue that contain milk are 
present. The criterion of milk in the mammary tissue can be only applied to dissected 
specimens. This mammary tissue is found on the inside of the skin in areas under the 
teats (Fig. 13e) and is usually whitish in colour and may spread beneath the skin some 
distance from the teats. Indicate in your field notes the presence of the mammary 
tissue and whether the tissue contains milk. 



Pregnancy is defined as the condition of having a developing foetus or embryo. 
Females in late pregnancy may have a swollen abdomen and at this stage it may be 
possible to detect embryos by squeezing or pinching the abdomen. 

In dissected mammals, carefully examine the uterus. When examining the uterus, 
it may be helpful to dissect it out and stretch it on a piece of white board or paper. For 
mammals with thick-walled uteri, you may have to open the uterus to examine it but 
this is seldom necessary for shrews, mice, or bats. The presence of embryos is 
positive evidence of pregnancy (Fig. 17c). Count and measure the crown-rump length 
(CR) of all embryos. This measurement (Fig. 17d) is from the top of the head to the 
end of the rump with the embryos in situ (not straightened). If more than one embryo 
is present, measure several and give an approximation of their size (e.g., 5 embryos, 
CR = 16 mm). Some collectors make it standard practice to denote the number of 
embryos in the right (R) and left (L) horns of the uterus (e.g., 5 embryos; 3L, 2R). In 
mammals having more than one embryo, some may die and be resorbed into the 
uterus. Resorbed embryos appear conspicuously smaller and underdeveloped when 
compared with normal ones. Be careful to distinguish any resorbed embryos when 
recording embryo counts (e.g., 5 normal embryos: 3L, 2R, CR = 15 mm; 2 resorbed 
embryos: 1L, 1R, CR = 3 mm). Embryos can be easily preserved by placing them in 
a vial of 10 per cent neutralized formalin. If embryological studies are contemplated, 
preserve in Bouin's solution (see section 6). 

The uterus should also be examined for the presence of placental scars. In some 
mammals (shrews, rodents, carnivores), after a female gives birth placental scars 
form at sites in the uterine wall where embryos were implanted. These scars appear as 
yellow to black pigmented spots on the inside of the uterus (Fig. 17b). Although the 
scars become increasingly paler and smaller with age, they may persist to one year in 
mice and rats. Generally the number of scars corresponds to the number of embryos. 
However, embryos that died during pregnancy will also leave scars on the uterine 
wall and because scars from several litters may be present, the number of scars is not 
always an accurate indication of litter size. Nevertheless, the presence or absence of 
placental scars is important to determine the reproductive history of the animal. If 
scars are readily visible, count them and record the number in each horn of the uterus. 
Where scars are badly faded, scars from several litters are present, or when scars are 
obscured by embryos, it may not be possible to count them accurately. If two (or 
more) sets of scars representing two (or more) different pregnancies are present, one 
set of scars will appear larger than the others. Although it may be impossible to count 
all scars, it is important to indicate that two (or more) sets of scars were observed. 
Examples of placental scar data: 2 sets of placental scars present — not counted; 4 
placental scars (3L, 1R); no placental scars present. 

With the data obtained from examining the uterus, females can be classified as 
follows: nulliparous — no embryos or placental scars; primiparous — embryos or one 
set of placental scars; multiparous — embryos and one (or more) sets of placental scars 
present, or two (or more) sets of placental scars present. 

4.6 Locality Descriptions 

Mammalogists are increasingly concerned with studies of geographic variation and 















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variation within populations. For this research detailed and accurate descriptions of 
collecting sites are required. The following locality data are required: country, state 
or province, county or equivalent, township or equivalent, local area or equivalent 
(town, lake name, name of nearby mountain), and the precise latitude and longitude 
to the nearest minute. Unfortunately, accurate mapping of collecting sites is 
frequently prohibited by insufficient data provided by collectors. Examples of 
problems resulting from incomplete and inaccurate locality descriptions are: locality 
description consisting only of a village name that cannot be located on any map, 
locality description consisting of the name of a town or village (no other data) when 
there may be several towns of the same name in the country, failing to distinguish 
between road distance (speedometer mileage) and map or airline distance. 

To avoid such problems, please follow these guidelines for documenting your 
collecting localities. If possible give the state or province and the county (or 
equivalent if such exists) in the country. This is helpful in limiting the search for a 
locality. The local area designation (a mountain, lake, river, valley) should be given 
when it is a prominent feature. To locate a collecting site precisely, give map or 
airline distance (miles or kilometres) and direction from a permanent reference point 
and latitude and longitude to the nearest minute if possible. A town or a mountain 
peak is a suitable reference point if it appears on small-scale maps (e.g., 1 :500 000). 
If the name of a town or village is the same as a nearby mountain, river or lake, insert 
the word town in parentheses, for example, Rainy River (town). Remember that a 
good reference point must be readily identifiable now as well as any time in the 
future. Transitory features that may be moved, renamed, or eliminated such as 
wayside taverns, small ponds, roads, highway numbers or junctions, and 
campgrounds should be avoided. Also avoid local names that do not appear on 
published maps for they may be impossible to locate. Distance from the reference 
points should be given in map or airline distances and not in road distance calculated 
from a speedometer. Road mileage in mountainous terrain is extremely inaccurate. 
Also, new road construction and reroutings make highway distances transitory. If 
road mileage is used, you must indicate that the distance is road mileage and not 
airline distance. 

Finally, we urge collectors to include with their field catalogues and field notes 
large-scale topographic maps (1 :50 000 or 1 :250 000) with collecting sites indicated. 
Maps of various scales for Europe, Asia, Australia, Latin America, Oceania, and the 
United States may be obtained from Defence Mapping Agency, Department of 
Defence, Topographic Center, Washington, DC 20315, USA. Topographic maps of 
Canada and an index listing available maps may be obtained from Energy Mines and 
Resources Canada, Surveys and Mapping Branch, Canada Map Office, 615 Booth 
Street, Ottawa, K1A 0E9. Topographic sheets for the Province of Ontario may be 
obtained from Ontario Ministry of Natural Resources, Map Office, Queen's Park, 
Toronto, M7A 1W4. 

4.7 Habitat Descriptions 

A description of the habitat in which the specimen was collected gives an indication 
of its ecological distribution. Therefore, provide as much habitat information as 
feasible for all specimens. 


A useful habitat description includes: (1) an indication of the general biome 
(desert, arctic tundra, savannah, coniferous forest, hardwood forest, alpine tundra, 
rain forest). (2) The dominant or most prevalent types of plants (palm grove, white 
spruce and balsam-fir forest, banana plantation, acacia scrub forest). If you know the 
scientific names of plants, use them in the description. (3) Note the elevation, for 
example, 3000 m above sea level (ASL). Elevation can be accurately estimated from 
topographic maps or with a pocket altimeter. For calculations from maps, use the 
units given on the map rather than converting metres to feet or vice versa. (4) Any 
pertinent information on the history of the area should be given (area burned over by 
forest fire in 1968; area cut over for logging in 1970). (5) Any pertinent information 
on soil type or geological formations. 

Rare mammals that are seldom captured by collectors warrant particularly detailed 
descriptions of their habitat. Frequently little, if anything, is known about the ecology 
of these species. For bats, describe roosting sites or netting sites (roosting in humid 
cave, roosting in palm tree, netted over small stream). 

Examples of informative habitat descriptions are as follows: trapped in alpine zone, 
400 m ASL, vegetation scrubby willows and mosses; netted in dense mature forest, 
1000 m ASL; from a large colony roosting in humid limestone cave, near banana 
plantation, 400 m ASL; trapped in treeless, grassland area 200 m ASL; trapped in 
mature jackpine balsam-fir forest. Figure 7 illustrates typical habitat data recorded in 
field notes. 

4.8 Methods of Capture 

Describe how the specimen was collected (trapped, shot, netted, found dead, road 
kill, poisoned). If all your specimens were captured in the same manner, then write 
this at the beginning of your catalogue or field notes to avoid repetition. 

4.9 Miscellaneous Field Notes 

This includes any miscellaneous behavioural or ecological observations that you may 
have made. Although these observations may seem to be trivial, they frequently 
prove to be valuable in natural history studies. Examples are as follows: netted at 
08 00 h; trapped between 08 00 h and 09 00 h; unusual colour phase; observed 
copulating with field number 20; specimen is young of field number 17; species 
observed feeding on papaya fruit; males and females roosting in different parts of the 
cave; tick on right ear; this colony of bats observed flying out of cave at 18 00 h; 
many of this species occupying the same burrow. Also record observations on 
climatic conditions in your field notes. Weather conditions may influence small 
mammal trapping or the netting of bats. Examples of these observations are: rainy 
season, dry season, heavy rains during trapping period. 

4.10 Photographic Records 

Colour slides or black and white photographs of specimens and collecting sites are a 
useful way to supplement documentation data. Photographs of rare mammals that are 


seldom captured by collectors or aberrant specimens (e.g., unusual colour phases) 
may prove invaluable, particularly if these specimens represent species new to 
science. Close-up photographs of the facial regions of live mammals taken in mist 
nets or live traps are an excellent way to illustrate the structure and colour of fleshy 
appendages (e.g., bat nose-leaves) that may fade or shrink in study skins or 
fluid-preserved material. You may also wish to photograph mammals that are 
prepared as complete skeletons without skins in order to make a permanent record of 
pelage. Photographs of carcasses of beached whales and dolphins prepared as skeletal 
material are also valuable. You will enhance your habitat descriptions (section 4.7) 
with photographs of the vegetation on trap lines or at bat netting sites. Photographs 
may be the only way to document rare or endangered species protected by law. 

Most field biologists prefer the 35 mm single-lens reflex camera because of its 
light weight and versatility. Various accessories (bellows, extension tubes, 
macrolenses) are made for close-up work. The authors have found a wide-angle lens 
(35 mm or 28 mm focal length) ideal for habitat photographs. 

5. Methods for Preparing Specimens 

Three kinds of museum specimens are usually prepared from mammals: skins with 
accompanying skulls and/or partial skeletons, complete skeletons, and entire 
mammals preserved in fluid. Each of these has advantages and disadvantages and the 
kind of specimen prepared depends on the objectives of the collector. Ideally, a 
representative series of a species from a given locality should contain all three types 
of specimens. Study skins are essential for analysing pelage colour and moult 
patterns; fluid-preserved specimens are valuable for studying anatomy and histology. 
Skeletons are useful for studies in comparative anatomy, geographic variation, and 
determining age but skeletons of many species are poorly represented in collections. 
Collectors are encouraged to prepare complete skeletons (at least one male and one 
female) for each species collected when feasible. 

The condition of the specimen will often determine the kind of specimen that 
should be prepared. For preserving in fluid, live mammals from mist nets or live traps 
are most suitable. Keep mammals alive in collecting bags or cages until they are to be 
prepared. Mammals from snap traps or specimens that have been frozen can usually 
be prepared as study skins. Mammals from snap traps should be prepared as quickly 
as possible because they decompose rapidly. Once the skin of a mammal begins to 
"slip" (fur falling out), it is difficult to salvage a satisfactory study skin from it. 
Decomposed specimens in which the internal organs have deteriorated and the fur is 
slipping are best prepared as skeletons. A "skull only" should be salvaged if the 
remaining carcass is badly damaged. 

5.1 Preserving in Fluid 

Because changes in tissues occur shortly after death, specimens should be preserved 
immediately after killing. To effectively kill small mammals (bats and mice) without 


damaging the skin and skull, use an airtight jar, can, or plastic bag with a wad of 
cotton containing a few drops of chloroform or ether. Avoid inhaling these chemicals 
as they are toxic to humans. Ether and chloroform are also highly inflammable. 
Larger mammals (hares, foxes) can be humanely and quickly killed by injecting them 
in the heart region with Euthanyl (sodium pentobarbital). The recommended dosage 
is 1 ml per 2.3 kg (5 lb.) body weight. Used by veterinarians, Euthanyl is a restricted 
drug that can only be obtained by prescription. You may find it necessary to calm 
large mammals with ether or chloroform before injecting them. 

The preparation of entire mammals in fluid involves two steps: fixing the tissues of 
the specimen with a solution such as 10 per cent formalin, Bouin's solution, or 
sodium acetate and transferring the specimen for permanent storage to a preserving 
fluid, for example, 65 to 70 per cent ethanol or 45 to 60 per cent isopropyl alcohol. 
"Fixing" halts enzyme processes in tissues and hardens or "sets" the specimen. 
Preservatives prevent the growth of microrganisms and also prevent gradual chemical 
or physical changes in the specimen's structure. 

Unless specimens are to be stored for two months or longer before shipment, the 
collector need be concerned only with fixation. Fluid-preserved specimens received 
from field collectors are washed for 12 to 24 h in water and then transferred to 65 per 
cent ethanol for permanent storage in museum collections. The best fixative for the 
field collector is a solution of 10 per cent buffered neutral formalin. Formalin is a 
solution of formaldehyde. The commercially available formalin is usually a 37 per 
cent (weight/weight) or 40 per cent (weight/volume) solution of formaldehyde. These 
commercial formaldehyde solutions are treated as 100 per cent formalin; therefore, to 
produce a 10 per cent formalin solution, mix one part 40 per cent formaldehyde with 
nine parts water. To reduce volume, most collectors carry full strength (40%) 
formaldehyde solutions in the field and dilute them to 10 per cent formalin before 
preserving their specimens. A wide-mouthed glass jar or a plastic pail makes a 
convenient container in which to dilute formaldehyde and to fix specimens. Because 
formalin solutions are usually acidic (pH 3.0 to 4.6), they tend to decalcify teeth and 
excessively harden tissue. To neutralize acidity, add a teaspoon of powdered borax 
(sodium tetraborate) or a tablespoon of household ammonia to 1 gal. (3.8 L) of 10 
per cent formalin. Best results are obtained by using a precise mixture of salts to 
buffer the formalin to neutrality (pH 7.0). For example, a mixture of 4 g of acid or 
monobasic sodium phosphate monohydrate (NaH2P04H20) and 6.5 g of dibasic 
sodium phosphate anhydrate (Na2HP04) will buffer 1 L of 10 per cent formalin. 
About 40 g of this salt mixture neutralizes 1 gal. (3.8 L). This dry, salt mixture can be 
prepared in advance and carried into the field in plastic bags. Other equipment for 
preserving specimens includes a hypodermic syringe and hypodermic needles for 
injecting the body cavity and larger muscles. If injection equipment is not available, 
mammals can be preserved by making a slit in the abdomen to open the body cavity to 
allow the formalin to enter. 

After specimens have been killed, weighed, measured, and assigned a number (see 
section 4), a field tag should be tied securely to each. If you use paper labels, be 
certain that they will not disintegrate in the preserving fluid. Using waterproof ink or 
a pencil, write the number and sex symbol ( ? for female and a* for male) on both 
sides of the label. 

Once properly tagged, lay the mammal on its back and, with the syringe full of 
formalin, insert the needle into the abdomen and slowly fill the body cavity until it 


becomes turgid. Do not inject too much fluid, but be sure that the body cavity is full 
and firm. For large mammals insert the needle into the larger muscles and inject a 
small amount of the formalin. For mammals with fleshy tails (rats) slit the tail with 
several cuts using a scalpel or sharp knife. 

After injection, the specimen is fixed by placing it in ajar, pan, or pail containing 
10 per cent buffered neutral formalin. Care should be used to keep the specimen in a 
normal, relaxed position for it will retain this shape permanently once it has been 
fixed. If the mouth is not locked tight, prop it open with a small piece of wood or a 
piece of cotton before the specimen is fixed. This permits examination of the teeth for 
identification without damaging the mouth parts. For bats, wings should be partially 
closed in a natural position. Usually the wings are in satisfactory shape if the bat is 
killed in a relaxed position. Do not overcrowd specimens in a container before they 
are thoroughly fixed (usually 12-48 h). Two factors are important: (1) to keep the 
specimen in a natural undistorted position; (2) to make certain there is sufficient 
volume of formalin to properly fix the specimen. A safe rule is to have sufficient 
formalin to completely cover the specimens. 

After 12 to 48 h specimens are fixed and they may be packed more tightly in 
containers for storage. Avoid using metal containers (unless acid-proof lined) as 
formalin causes almost immediate rust and corrosion that will discolour specimens. If 
only metal lids are available, a waxed, cardboard liner will seal the container for only 
a short time because of corrosion. Therefore, place a piece of waxed paper or a sheet 
of plastic over the mouth of the jar before screwing on the lid. If specimens are to be 
stored for a long period before shipment (more than 8 weeks), they should first be 
washed thoroughly in fresh water and then placed in 65 to 70 per cent ethanol or 45 to 
60 per cent isopropyl alcohol. 

5.2 Preparing Skins 

Three types of skins can be prepared for museum specimens: (1) Traditional study 
skins that are filled with cotton or a similar material to approximate the natural shape 
of the mammal. As the leg bones are ordinarily left in the skin, a complete skeleton 
cannot be obtained and usually only the skull is kept. (2) Flat skins, which consist of 
the skin stretched over a cardboard outline. For a flat skin, it is possible to obtain a 
skull and most of the skeleton. (3) Tanned skins, which are first dried and later tanned 
for permanent preservation. A skull and skeleton can be obtained from a mammal 
prepared as a tanned skin with only the terminal digits left on the skin. 

The choice of skin depends on the size of mammal and the collector's objectives. 
Small mammals (bats, rodents, insectivores) are prepared as study or flat skins. The 
study skin has been traditionally used by museum collectors. This type of skin may be 
more time-consuming to prepare, especially for the inexperienced, but it is invaluable 
for studying pelage. Flat skins have the advantage of being quick and easy to prepare 
and they provide both a skull and a skeleton. However, flat skins are difficult to 
compare with study skins when analysing pelage. Skins from mammals larger than a 
fox are too bulky to be made into study or flat skins on cardboard and must be 
prepared as tanned skins. Study skins can be prepared for small fur-bearing mammals 
up to the size of a fox, however, some collectors prefer to make tanned skins for these 


A. Study Skins 

Begin with a midventral incision from the level of the last rib to near the anus (Fig. 
18). Always cut to one side of the penis or vagina so that the external genitalia remain 
attached to the skin. To keep the skin clean and dry, cornmeal, borax, magnesium 
carbonate powder, or sawdust may be sprinkled on the skin or placed in a skinning 
tray or pan to absorb blood and body fluids. Another skinning technique is to make an 
incision that extends across the lower abdomen and down the inside of the leg to the 
heels (Fig. 18). Leave the external genitalia on the skin by cutting between the anus 
and genitalia. For males that have a baculum in the penis, be careful not to cut or 
damage this structure for it may be an important aid in identification. With mammals 
such as bats or mice, the baculum can be left intact in the penis to dry on the skin. If 
bacula studies are anticipated, remove the entire penis and store it in 100 per cent 
glycerine, 10 per cent formalin, or 70 per cent alcohol. For larger mammals, the 
baculum should be extracted from the penis, tagged and dried with the skull and any 
skeletal material. 

With fingertips, a scalpel handle, or blunt forceps, work the skin free of the body 
wall in the vicinity of the incision. Try not to cut into the body cavity. Holding the 
hind foot, push the knee joint upward towards the midline of the body. Peel the skin 
off the leg to the ankle, then sever the hind leg at the hip or knee joint with scissors or 
a scalpel (Fig. 19). When the hind legs are free, work the skin to the base of the tail. 
Use care while skinning around the anus and the anal scent glands found in some 
mustelids (Mustelidae). 

If the mammal has a nonfleshy tail (e.g., some bats), cut it close to the trunk and 
leave the tail vertebrae in the skin. If a fleshy tail is present, slip it out of the skin and 
later replace it with a wrapped wire. Rolling the tail on a table top or skinning board 
will help loosen the connective tissue that attaches the tail vertebrae to the tail sheath. 
For shrews, mice, and other small specimens grasp the tail at the base of the sheath 
with the thumb and index finger (Fig. 20) of one hand. Press the thumb- and 
finger-nails firmly against the tail vertebrae. Then with the other hand, slowly pull the 
tail vertebrae until they are free of the skin. If the tail vertebrae break off in the tail 
sheath, you must split the skin of the tail and remove the vertebrae. After inserting a 
tail wire, the incision should be sewn with a fine needle and thread. For larger 
mammals, it may be necessary to hold the base of the tail sheath with heavy forceps 
or two blocks of wood. For mammals larger than a squirrel, it is usually necessary to 
split the tail by a longitudinal incision in order to remove the vertebrae. 

With the tail free, the skin can now be peeled back to the region of the front legs 
(Fig. 21). Do not pull the skin off the body, as this will result in an overstretched 
study skin. A recommended method is to use one hand to gently push the skin off the 
body and with the scalpel held in the other hand, sever any connective tissue holding 
the skin to the carcass. Remove the skin from the front legs down to the ankle. For 
bats, detach the skin as far as the elbow joint. With scissors or a scalpel, cut the front 
legs (or wing bones of bats) at the shoulder joint (Fig. 22). Peel the skin over the chest 
area to the base of the skull. 

Probably the most difficult stage of the skinning operation is to remove the skin 
from the head region without damaging the ears, eyelids, lips, and skull. Use a sharp 
scalpel for skinning the head region. Carefully work the skin over the head until you 
reach the cartilaginous bases of the ears (Fig. 23). Pick away any fatty tissue that may 


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Fig. 18 Initial incision is made midventrally (dashed line) or across the heels (solid line). 



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Fig. 19 Severing the hind leg at the knee joint. 


Fig. 20 Removing tail vertebrae from the tail sheath. 

obscure the ear cartilage and sever the cartilage at the base of the ear. Continue to peel 
the skin over the head until the eyes are exposed. With the skin held away from the 
head, cut the membrane that covers the eyes (Fig. 24). The skin should be still 
attached in the eye region at the front corner of the eyelid. Carefully cut this 
attachment with your scalpel but avoid cutting into the eyelid, as the skin of the eye 
region will tear when the skin is stuffed. Work the skin to the lips and sever the 
connective tissue that attaches the lips to the skull. Finally, peel the skin forward until 
it is attached to the body only at the tip of the nose. Cut the nasal cartilage being 
careful not to cut into the nasal bones of the skull (Fig. 25). 

Once the skin is removed, dissect the carcass for reproductive data (section 4.5) 
then direct your attention to the skin. Remove all fat and excess flesh from the skin. 
To accelerate drying and inhibit insect damage, rub a drying agent into the flesh side 
of the skin. At the ROM, we use magnesium carbonate powder for study skins and flat 
skins. This powder is available from most biological or chemical supply companies. 
Borax can also be used as a drying agent-preservative. However, there is some 
evidence that borax may affect red-coloured pelage; therefore, try to keep borax 
powder off the fur. Arsenic, arsenic and borax, arsenic and alum, alum and potassium 
nitrate, and arsenic soap have been used in the past as drying-preserving agents. 
Because alum may affect fur colour and arsenic is toxic, these chemicals are not 


Fig. 21 Skin being peeled off the carcass. 


Fig. 22 Location of cut for severing the front legs. 

Fig. 23 Cutting the cartilage at the base of ears. 


Fig. 24 Skinning around the eye. 

Fig. 25 Removing the skin from the nose. 


recommended and they should only be used if magnesium carbonate or borax is not 

For smaller mammals (mice, shrews, and bats), fat and flesh can be picked off the 
skin with your fingers; however, it may be necessary to use a dull knife to remove this 
material from skins of larger mammals. A simple method for degreasing skins that 
have heavy fat deposits is to dip them in naphtha or white gas. Shake off any excess 
gas and roll the skin in sawdust to hasten drying. Do not wring the skin as this will 
stretch it. A liberal dusting of fur with magnesium carbonate or sawdust before and 
during skinning usually prevents blood from adhering to the skin. Heavily matted 
blood around wounds may be removed with water or alcohol on a cotton wad, 
followed by dusting with magnesium carbonate or sawdust. 

Remove the muscle tissue from the leg or wing bones with scissors, scalpel, or 
forceps (Fig. 26) and rub the bones in magnesium carbonate. Restore the legs to their 
approximate original shape by wrapping the bones with cotton to replace the muscles. 
With the skin still reversed, sew the lips together (Fig. 27). 

Fig. 26 Removing muscle tissue from the leg bone. 


Fig. 27 Stitch used to close the lips. 

Now the skin is ready to fill with a body and head made from a single piece of 
cotton. Some collectors prefer to use fine tow rather than cotton. Roll the cotton into a 
smooth cylindrical bundle that is slightly longer and thicker than the body of the 
mammal (Fig. 28). Form the head region of the cotton filler with a pair of forceps by 
pressing in the centre at the end of the roll (Fig. 29). Grasp the two corners on either 
side of the forceps, fold together, and take a new hold of the pointed end and shape as 
a smooth cone (Fig. 30). 

Place the cone into the head of the skin and reverse the skin over the points of the 
forceps (Figs. 31, 32). Adjust the eyes, ears, and mouth, then continue to reverse the 
skin slowly over the cotton until the specimen is completely filled. The length of the 
cotton body can be trimmed with scissors to fill the skin properly (Fig. 33). For study 
skins of small species of bats, mice, and shrews, some collectors prefer to construct a 
separate head rather than use a single piece of filling. Best results are obtained by 
constructing the head from fine tow and wrapping it with a thin wisp of cotton. Use 
the skull as a guide for size. Instead of sewing, hold the lips in position by inserting 
small pins into the tow. After the head is shaped, fill the body with a single piece of 

If the tail vertebrae were removed, then an appropriately sized wire wrapped with 
cotton must be inserted into the tail for support (Fig. 34). The collector will require 
the following gauges of wire for preparing skins: 12 gauge (hares), 16 gauge (large 
squirrels), 18 and 20 gauge (small squirrels), 22 and 24 gauge (mice, rats, shrews), 
26 gauge (small shrews and small bats). If available, use Monel wire as it does not 
corrode. Cut the wire to a length that extends from the tip of the tail to midway into 
the body. A loop at the body end of the wire provides added strength and stability to 
the finished specimen. Wrap thin wisps of cotton to form a shape similar to that of the 
original tail vertebrae (Fig. 35). It may be necessary to moisten the tail wire with 
saliva to make the cotton adhere. Species with long tails that taper to a fine tip 
(jumping mice), present a problem because the tail tip is too narrow to accommodate 
the usual tail wire. A very thin wire (26 gauge) can be tapered to a fine point with a 
file. However, a better method is to prepare tapered wires in advance of fieldwork by 
dipping precut wires into an acid solution to form a long, tapering point that will fit 
the tail sheath. Dipping Monel wire for 5 min in a solution of one part hydrochloric 
acid (HCL) and two parts nitric acid (HNO3) will effectively produce a tapering 
point. Oxide deposits on the wire can be removed by placing wires for several 
minutes in an enamel tray containing hydrochloric acid. Pad the portion of the looped 


Fig. 28 Rolling cotton into a cylindrical bundle. 

Fig. 29 Grasping the cotton with forceps. 

Fig. 30 Forming the head with fingers and forceps. 

tail wire that extends beyond the tail into the body cavity with a thin piece of cotton 
then stitch the midventral incision with a fine needle and thread (Fig. 36) and tie a 
field tag (see section 4.2) to the hind foot of the skin. 

The next step is to anchor the study skin to a pinning board (cardboard, corkboard, 
Styrofoam, or wallboard) for drying. Careful pinning is the key to a well-prepared 
skin. For most mammals the front and hind feet are positioned parallel to the body 
and held in place with pins through each foot and a pair of pins at the outer side of 
each hind foot near the heel (Fig. 37). Anchor the tail by a pair angled across its base 
and by one pair angled across the tip. To shape the ears and head, use pins placed 
against the side of the skin. 

Check to be sure the head is symmetrical and, if necessary, a thin (insect) pin may 
be used to anchor it in place. The eyelids may be held open by pulling through a small 
bit of cotton from the head. A final check of the specimen should be followed by 
cleaning the fur with a small brush (a toothbrush works well) to remove dirt or dust 
(Fig. 38). 

A recommended method of pinning bat wings (Fig. 39a) is to place a sharp pin 
(insect pins are preferable, but any sharp pin will do) in the wing joint near the thumb. 
Position the wing so that the forearm is nearly parallel to the body and the upper arm 
joins the body in a natural position at a slight angle. Some of the membrane above the 
forearm and upper arm should be exposed, but the wings should not be overstretched. 
The second pair of pins is inserted at the elbow to hold the forearm in the desired 


Fig. 31 Cotton body is inserted into the head region. 

Fig. 32 Skin is reversed over the cotton. 


Fig. 33 Cutting the cotton to the appropriate size. 

Fig. 34 Wrapping the tail wire with cotton 

Fig. 35 Inserting the wrapped wire into the tail sheath. 

Fig. 36 Stitch used to close the body incision. 

Fig. 37 Skin is pinned out for drying and a field tag is attached to hind foot. 


Fig. 38 Brushing the fur with a small brush. 

position. A third pair of pins may be required to hold the end of the upper arm in the 
appropriate position close to the body. Pin each foot, pulling the body into the desired 
position. If a tail is present, extend and anchor it with a pin at the tip and with crossed 
pins over the base of the tail near the body. If a membrane is present between the tail 
and legs, extend and anchor it by placing pins near the end of the calcar (the 
cartilaginous structure extending along the edge of the membrane from the heel of the 
foot). The wing bones are then pinned so that each digit or finger is spread slightly 
away from adjoining ones. 

The pinned specimen must be dried thoroughly before shipping or transporting. 
Drying time will vary considerably with local conditions. In hot dry climates, stuffed 
skins may dry in one day; however, in humid climates it may be extremely difficult to 
dry skins completely. A shaded area with good air circulation provides the best 



Fig. 39 A Dorsal view of a bat study skin pinned for drying. 

B Corresponding skull is shown in a gauze bag with field number attached. 

conditions. Do not place skins in direct sunlight as this fades pelage and intense heat 
may cause excessive shrinking of skins. Under poor drying conditions, it may be 
necessary to unpin specimens and to expose their undersurface to the air by turning 
them upside down. Take care not to bend or break the ears when doing so. When 
skins are dried, remove the pins and store them in chests or shipping boxes (see 
section 7). 

Ants and egg-laying flies can cause extensive damage to skins and most collectors 
protect their drying skins by storing them in special wooden cases that have screened 
openings for ventilation. A properly screened drying chest should be part of your 
standard field equipment; however, a temporary drying container can be constructed 
in the field using a few pieces of wood and cheesecloth. At the ROM we use wooden 
drying chests (90 cm x 50 cm x 35 cm; 35 inches x 20 inches x 14 inches), that 
contain 80 cm x 42 cm (32 inches x 17 inches) sheets of 2.5 cm (1 inch) Styrofoam. 
The sheets of Styrofoam are separated by 2.5 cm (1 inch) high, wooden frame 
spacers. For small mammals (shrews, mice), a single spacer is sufficient for 
separating Styrofoam sheets, for larger mammals (hares), two or three spacers are 
required. Study skins are pinned to the Styrofoam for drying. When they are 
sufficiently dried, pins are removed and each layer of skins is covered with cotton. 

B. Flat Skins 

Flat skins mounted on cardboard can be prepared for such species as shrews, mice, 
squirrels, and small carnivores. Many collectors use the method described by 


Anderson (1965) where both front and hind feet are left on the skin. We recommend 
the following procedure because it enables the collector the obtain both a flat skin and 
a skeleton from the specimen. The skeleton is complete except for the one front and 
one hind foot that are left on the skin. 

Rather than making a cut along the midline of the abdomen, begin the cut at one 
heel, cutting through the skin at the back and inner side of the leg, across the base of 
the tail between the anus and external genitalia, and then extend the incision to the 
opposite heel (Fig. 18). Leave the external genitalia on the skin. Detach the skin from 
the legs and cut through the leg bone at the ankle of only one leg; leave that foot on 
the skin. On the opposite leg, detach the skin to the ankle and then cut the skin there. 
This foot is left attached to the body of the mammal as part of the skeleton. Now 
remove the skin from the tail and pull it towards the front legs. Do the same for the 
front legs as the hind legs — leave one foot on the body of the mammal and detach the 
other foot with the skin. Remove the skin from the head region being careful not to 
damage the ears or lips. Dissect the carcass for reproductive data (section 4.5). Clean 
all fat and excess flesh from the skin and, to hasten drying, rub a drying agent (borax, 
magnesium carbonate) into the flesh side of the skin (see section 5.2A). 

Unlike the conventional study skin, the flat skin is stretched on a piece of 
cardboard or corrugated pasteboard. To prepare the stretcher, lay the skin flat on the 
board and trace its outline. Shape the card with scissors, leaving a sufficient amount 
of the card behind the shaped outline to support the tail and to permit writing of field 
data (Fig. 40). Use a board sufficiently thick to support the skin fully. Cardboard 
sheets of different thicknesses should be carried into the field by collectors who 
intend to prepare flat skins. Pull the skin over the stretcher board, being careful not to 
overstretch. For small mammals, the skin is put on the board, fur side out, but for 
mammals larger than a squirrel, the skin should be stretched on the board flesh side 
out for a few days. Then, after the skin has partly dried, reverse it and pull it on to the 
board, fur side out. 

Insert a wrapped tail wire for support as with conventional study skins (see section 
5.2A) and tie the hind foot and base of the tail to the board with thread (Fig. 41). You 
will need heavy sail-makers' needles for piercing heavy cardboard. Small pins are 
used for holding the front foot in position for drying and for shaping the lips if 
necessary. Use a toothbrush for a final cleaning of the fur. Tie the field number (see 
section 4.2) to the stretcher card and write the field number on the card in case the tag 
is lost (Fig. 41). 

A modified flat skin is used by museum collectors for hares and rabbits (Fig. 42). 
Because the hind legs are supported on a wooden stick, this type of skin is sturdier 
than the conventional study skin. Also it requires less space for storage. A heavy 
stretcher (corrugated cardboard) is cut to fit the skin. A wooden stick which extends 
from the head to beyond the hind feet is attached to the board by means of cord or 
wire. Wrap the cardboard stretcher with a thin layer (0.5 cm; 0.25 inches) of cotton 
and insert it into the skin. It may be necessary to trim the posterior end of the board 
with shears to fit the skin properly. Wires should be inserted in each leg to provide 
support. Secure the hind legs to the stick with wire or heavy thread; the front legs are 
positioned parallel to the head. The ears can be held together flat against the skin in a 
natural position with a single stitch of thread. Sew the skinning incision and tie a field 
number to one hind leg. At the ROM, we are using a modified version of this method 
for our hare skins. The skin is prepared in the same manner as the traditional hare flat 




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skin except that only one hind foot and one front foot are left on the skin. The other 
feet are left on the carcass as part of the skeleton. 

Flat skins dry quickly and under optimum conditions they may be sufficiently dried 
in 24 h. Follow the precautions for drying study skins, that is, keep flat skins out of 
direct sunlight and protect them from insect pests. After they have been dried, flat 
skins can be packed compactly into boxes for shipping. 

C. Skins to be Tanned 

Mammal skins larger than a fox must be prepared for tanning, although smaller 
fur-bearing mammals (foxes, ermine, beaver) may also be prepared for tanning. 
Skinning pelts for tanning may be either "cased" or "open". Pelts from fur-bearers 
other than beaver are usually prepared "cased" by professional trappers; however, 
the choice of skinning method is really a question of personal preference. Large 
mammals such as bears, deer, moose, and seals should be skinned "open". For a 
discussion of techniques for mammals that require special treatment, see Anderson 

"Cased" skins are removed from the mammal in much the same manner as the flat 
skin (see section 5.2B). Make an incision from foot pad to foot pad along the hind 
legs. Detach the skin from the hind legs to the foot. The skin can be separated from 
the carcass by pushing down with thumb and fingers between the skin and carcass. 
Skin out the feet leaving only the claws on the skin and remove the tail vertebrae from 
the tail sheath. For mammals larger than a squirrel, split open the tail on the ventral 
side. With the tail vertebrae free, pull the skin down the carcass to the front legs. 
Detach the skin from the front legs, leaving the claws on the skin, then pull the skin 
over the head region. Carefully skin around the ears, eyes, mouth, and nose. 

Cased skins should be stretched, fur side in, over a frame for drying. For the 
collector who prepares the occasional pelt, a stretcher can be improvised from soft 
wood, wire, or corrugated cardboard. However, if you plan to collect a number of 

Fig. 41 Example of a flat skin prepared from a chipmunk (Sciuridae). 


Fig. 42 Modified flat skin used for hares. Hind feet are tied to a stick which is attached to a 
cardboard outline covered with a thin layer of cotton: a leg wires, b cardboard 
outline, c wooden stick. 

fur-bearers, then you should make a drying board for the various species that you 
intend to collect. Drying boards approved by the fur trade for different species are 
available commercially (Southeastern Outdoor Supplies, Route 3, Box 503, Bassett, 
Virginia, USA). Adhesive-backed templates in various standard sizes and shapes are 
also sold commercially. The template is placed on a board which is then cut to the 
appropriate size and shape. 

For "open" skins begin with a midventral incision from the throat extending 
posteriorly to the tip of the tail, being careful to cut to one side of the genitalia (Fig. 
43a). Cuts are then made from this midventral incision along the inside of each leg to 
the foot. Usually the claws or hoofs are left on the skin; however, if the carcass is to 


Fig. 43 Cuts for skinning a large mammal. 
A Ventral view of cuts. 

B Enlarged view of cuts for skinning leg and hoof, 
c Cuts for skinning around antlers. 

be prepared as a skeleton, then leave only one hind foot and one front foot on the 
skin — the other feet remain on the carcass. The skin of some mammals (deer and 
caribou) can be pulled off easily by hand, but for other mammals (bears), the skin 
must be removed by careful cutting. Figure 43c illustrates the cuts that should be 
made for skinning around horns or antlers. 

Once the skin is off, remove as much flesh and fat as possible and sponge off any 
blood from the fur. Be extremely careful when defleshing. If too much of the skin is 
removed, the fur will fall out when the pelt is tanned. An ordinary dull table knife 
makes a good fleshing tool. Spread a layer of salt (sodium chloride) evenly on the 
entire flesh side of the skin and rub it in thoroughly with the hands. Stretch the skin 
out, flesh side exposed, and allow it to dry for about 24 h (longer in humid 
conditions). Cheesecloth or mosquito netting can be used to keep egg-laying flies off 
the skin. After 24 h, shake off any water and excess salt. Resalt the skin, then fold the 
head and legs in and roll the skin into a bundle. The skin should be periodically 
checked for signs of decomposition and the presence of fly eggs or larvae and it may 
be necessary to apply another treatment of salt. The skin must be dried as thoroughly 
as possible prior to shipment or transport. After the skin is received from the 
collector, museums usually send it to a commercial tanner for tanning. It is important 


not to treat the skin with chemical preservatives other than salt in the field as these 
may interfere with the tanning process. 

5.3 Preparing Skulls 

As soon as the study skin is prepared, direct your attention to the skull. Care should 
be taken to prevent damage to any part of the skull. Separate the skull from the 
carcass by severing at the joint of the skull and the first vertebra (atlas). Fragile skulls 
from small mammals (shrews, mice, small bats) are dried without any cleaning. 
However, the brain, eyes, tongue, and heavy muscle layers should be removed from 
skulls about the size of squirrels or larger. A piece of wire with a small hook on the 
end can be used to pick the brain tissue out of the cranium, or the brain tissue can be 
flushed out of the skull with a syringe. Cut the muscles attaching the eyes and tongue 
with scissors and then pull these organs from the skull using forceps. Heavy muscle 
tissue can be removed by using a scalpel or scissors, but be extremely careful not to 
damage the thin processes on the skull. For skulls with antlers in velvet (deer, moose, 
etc.), it may be necessary to split the velvet with a knife to facilitate drying. 

Thoroughly dry skulls in the field. An efficient technique for drying skulls is to 
place them in cloth bags. Gauze skull bags are usually supplied to ROM collectors; 
however, if you do not have these bags, similar ones can be constructed from 
cheesecloth or other porous material. Write the field number in pencil on a heavy 
paper or cardboard tag and tie it to the skull. The same number can be attached to the 
outside of the bag (Fig. 39b). Skull bags can be strung on a wire and put out in a 
ventilated place to dry. Ensure that skulls cannot be reached by animals that might be 
attracted to them. When using a vehicle in the field, an excellent technique for drying 
skulls is to tie wire strings of skulls securely under the hood where the heat of the 
engine and air flow during travelling quickly dries them. It may be necessary to store 
skulls inside the vehicle at night to protect them from such predators as racoons or 
cats. After skulls have completely dried, they can be compactly packed for shipping. 
As with study skins, the time required to dry skulls varies with climatic conditions. 
Before packing, ascertain that there is a skull for every skin and note in your 
catalogue any damaged skulls. 

5.4 Preparing Skeletons 

If the specimen is to be prepared as a flat skin or tanned skin, then remove the skin by 
the methods described in section 5.2. However, if the mammal is decomposed with 
the fur slipping, remove and discard the skin in the quickest manner possible, being 
careful not to damage the skeleton. Because wing membranes must be removed from 
the bones of the hand, bats prepared as skeletons should be skinned as follows. 
Detach the skin from the legs as far as the ankle, then cut the skin with scissors at the 
ankle joint. Peel the skin from the carcass until you reach the wings. Detach the skin 
from the arm of the bat; then by slowly pulling towards the wing tips, peel the wing 
membrane off the wing bones, being careful not to break the delicate bones. Remove 
the skin from the head region in the usual manner (see section 5.2). 


When the skin is off the carcass, dissect for reproductive data (see section 4.5), 
then extract all internal organs from the body cavity. Any organs left in the carcass 
will decompose rapidly. Next, remove the larger muscle tissues from the bones. The 
amount of defleshing required depends on the size of the mammal. To avoid 
damaging the delicate skeletons of such small mammals as shrews, mice, or small 
bats, do not attempt to cut off the small amount of flesh. Simply leave the skull 
attached to the skeleton and allow the entire carcass to dry. For medium-sized 
mammals (squirrels, hares, fruit bats, small carnivores), remove the muscles with 
scissors and forceps. Although it is not necessary to disarticulate the skeleton for 
these mammals, that is, to cut the ligaments that hold the bones together, you must 
separate the skull from the skeleton and extract the brain, tongue, and eyes (see 
section 5.3). Considerable work is involved in defleshing the skeletons of large 
mammals (deer, bears, seals) and a sharp skinning knife is essential for cutting the 
heavy muscles and ligaments. You must at least partially disarticulate large mammals 
in order to reduce the skeleton to manageable sections. Separate the skull and remove 
the brain, tongue, and eyes (see section 5.3). 

After defleshing, skeletons must be dried thoroughly before shipping or 
transporting. The best method for drying skeletons of small- to medium-sized 
mammals is to place them in gauze bags that protect them from egg-laying flies but 
provide ventilation for rapid drying. If you do not have these bags, make them from 
cheesecloth or similar material. Tie a field number to the skeleton and place a single 
skeleton in each bag. For bats tie the wing bones against the body to prevent them 
from breaking off. After tying the top of the bag tightly with string, hang it to dry 
following the precautions described for drying skulls (see section 5.3). Burlap bags 
make excellent drying bags for skeletons of large mammals. Each bag should have 
the field number securely attached. Two important rules to follow when preparing 
skeletons are: (1) do not place skeletons in plastic bags, as they will decompose 
instead of drying; (2) do not treat skeletons with any chemicals as they will inhibit the 
activity of dermestid beetles that are used in many museums to clean skeletons. 

6. Special Techniques 

6.1 Karyotyping 

Slides of somatic chromosomes can be prepared in the field using the in vivo bone 
marrow technique. The method is simple, quick, and produces slides of good quality 
for conventional staining. Most workers use chromosomes from tissue culture for 
banding studies. 

A large metal toolbox is invaluable for carrying karyotyping equipment in the field 
(see Appendix 2). Sodium citrate solutions will support growth by bacteria and 
consequently must be prepared fresh each day. Weighing and separating the required 
daily quantities of sodium citrate crystals before a field trip saves considerable time. 
Small vials or plastic bags are suitable containers for sodium citrate. Although slides 
can be stained in the field, staining solutions add considerable bulk and weight to the 


karyotyping kit. Slides can be stained with good results 1 to 2 months after 
preparation in the field. Most hand centrifuges hold four centrifuge tubes; therefore 
one can process four specimens at a time. Although the volume of suspension in 
centrifuge tubes could produce a dozen or more slides, we generally prepare four to 
six slides for each specimen. 

Practise the procedure in the laboratory before attempting to do it in the field. This 
will enable you to become familiar with the various steps and to verify the quality of 

Mammals that have been karyotyped must be kept as voucher specimens (study 
skins or preserved in fluid), for chromosome slides without voucher specimens are 
virtually worthless. 

The following technique modified from Baker (1970) has been used by ROM staff 
to karyotype bats, small rodents, and small carnivores in the field. A list of equipment 
required for karyotyping is given in Appendix 2. 

1. Dilute one or more vials (10 mg) of Velbe (Eli Lilly & Company, Indianapolis, 
Indiana, USA) with distilled water to produce a 0.025 per cent solution. One method 
is to take a serum bottle containing 100 ml of bacteriostatic sodium chloride (Abbott 
Laboratories, Montreal, Canada) and remove and discard 20 ml of sodium chloride 
solution with a sterile syringe. Dissolve the contents of two 10 mg vials in the serum 
bottle to produce an 80 ml stock solution of Velbe. Keep this refrigerated. For a 
working solution, transfer 10 ml of stock solution to any empty Velbe container. If 
working with colchicine, use a 0.04 per cent solution. 

2. With a 1 ml tuberculin syringe, inject the mammal intraperitoneally. The dosage 
of Velbe is 1 unit (0.01 ml) per gram of body weight. Use the same dosage with 
colchicine. After injection, the mammals must be kept alive for 1.5 to 2 h. Bats can 
be kept in individual collecting bags; rodents or carnivores in cages. 

3. After 1.5 to 2 h, anaesthetize the mammal, dissect out the humerus (bats) or the 
femur (rodents, carnivores) being careful not to damage the proximal end. Remove 
muscle tissue from the bone. 

4. Using a 5 ml Luer Lock syringe, flush out bone marrow with 3 ml of 1 per cent 
sodium citrate solution (made fresh daily). The size of the syringe needle will depend 
on the size of the bone. Generally, no. 21, 24, or 26 needles are required for small 

5. Vigorously break up and suspend cells with a pipette. Then let the solution stand 
for 5 to 20 min (more than 20 min may rupture cells). 

6. Centrifuge at 500 to 1500 rev/min for 2 to 4 min. 

7. With a pipette remove and discard the supernatant fluid being careful not to 
disturb the clump of cells. 

8. Slowly add one pipette of Carnoy's fixative (three parts absolute methanohone 
part glacial acetic acid) to the centrifuge tube. By carefully adding the fixative down 
the inside of the tube with a pipette, you do not disturb the clump of cells. With the tip 
of the pipette close to but not touching the cells, carefully extract the fixative. Add 
fresh fixative, suspend the cells and let stand for 10 to 12 min. 

9. Centrifuge for 2 to 4 min, discard supernatant, then resuspend the cells in fixative. 

10. Repeat this procedure two or three times. 


11. After the final centrif ligation, resuspend the cells in fixative. 

12. With a pipette, place two or three drops of suspension on a clean microscope 
slide. Ignite by quickly passing a match over the slide. Do not touch the slide with the 
match. Some workers prefer to air dry slides. Use microscope slides with frosted ends 
and record the field number of the specimen on the slide with a lead or diamond 

13. Slides can now be stained in the field or sealed in a slide box and later stained in 
the laboratory. To prepare the stain, mix eight parts of warm distilled water with one 
part Giemsa stain (filter the Giemsa before mixing with water). Slides are stained for 
13 min in a Coplan jar. About 50 ml of stain are required to fill a Coplan jar. The 
destaining process requires five steps in Coplan jars: (1) rinse in acetone; (2) 1 min in 
acetone; (3) 1 min in acetone xylol (1:1); (4) 1 min in xylol; (5) 2 min in xylol. 
Mount cover slips using permount while slides are still wet with xylol. 

If slides are not stained immediately, store them in slide boxes. It is important in 
the field to keep slides dry and free from dust. Place a small packet of silica gel 
crystals in each box and tape the sides of the box to make it airtight. Store slide boxes 
in plastic bags. 

6.2 Collecting Parasites 

Mammals are usually hosts of many parasites. Those found externally on the body are 
ectoparasites; parasites found in internal organs are referred to as endoparasites. 


Ectoparasites are insects (parasitic flies, fly larvae, lice, and fleas) or arachnids (mites 
and ticks) that feed on the body fluids, dead skin, tissues, or hair of the host mammal. 
In bats usual parasites are bat flies and mites, whereas lice, mites, ticks, and fleas are 
commoner on rodents and other mammals. Although many techniques exist to obtain 
ectoparasites, the following is a simple method suitable for field use. If the mammal 
is alive, put it in a clean plastic or cloth bag with a piece of cotton soaked in 
chloroform or ether to kill the ectoparasites and the host. Shake the dead mammal and 
the inverted bag over a white porcelain tray or a piece of white paper and recover the 
parasites which will be conspicuously dark on the white background. A vigorous 
brushing of the fur with a toothbrush will remove any ectoparasites that are caught in 
the fur. Because ticks imbed in the skin, they are difficult to remove from the host 
without damaging the mouth parts. For fluid-preserved specimens, leave ticks in 
position on the mammal and make a note on your data sheets that these ectoparasites 
are present. For study skins detach the ticks by snipping away some of the skin of the 
host in which it is imbedded. 

There are several precautions to follow when collecting ectoparasites. It is essential 
to separate different species of mammals in collecting bags. If different species of 
mammals are put in the same collecting bag, then host data will be meaningless, for 
most ectoparasites may move from one mammal to another in the bag. Handle 
ectoparasites carefully, as legs, wings, and other body parts are delicate. Use 


jeweller's forceps or fine-tipped brushes moistened in alcohol to pick up 
ectoparasites. Ensure that instruments, killing bags, and pans are washed and clean 
before processing a new specimen. 

Preserve all ectoparasites in small, screw-cap or rubber-stoppered vials with 70 per 
cent ethanol (do not use formalin, as it hardens specimens). Write the field number of 
the host and any other pertinent data (e.g., site on body of host) in waterproof ink or 
pencil on sturdy paper and place it in the vial. Note on your catalogue sheets or field 
notebook that ectoparasites were preserved for that particular specimen. If you wish 
further information on parasites, refer to the appropriate publications listed in the 


Endoparasites most frequently found in mammals are helminth worms (trematodes, 
cestodes, and nematodes). Trematodes or flukes are small, flattened worms that occur 
in the digestive tract, liver, lungs, and other internal organs. Cestodes or tape worms 
are long, many-segmented worms that live in the intestine of the host; nematodes or 
round worms are unsegmented worms found in most organs, including muscles. It is 
beyond the scope of this manual to describe the many special techniques for 
preserving endoparasites. But for the field collector who may occasionally find these 
endoparasites and wishes to preserve them for identification, we recommend that they 
be stored in 70 percent alcohol. A small amount of glycerine (5 ml in 100 ml of 70% 
ethanol) will keep parasites pliable and reduce hardening of tissues. Label vials with 
the field number of the host and note in the catalogue or field notebook that parasites 
were preserved. For descriptions of special methods for preserving endoparasites, 
consult references given in the bibliography. 

6.3 Tissues for Biochemical Study 

Fresh tissues from various organs (heart, kidney, liver, ovary, testes, and muscle) can 
be preserved in the field by quick freezing. 

Remove the desired organs from freshly killed specimens. Cut a 5 to 10 mm cube 
of tissue from the organ with scissors or a scalpel and wash in physiological saline to 
remove contaminating blood. The saline solution should be prepared fresh each day 
and each tissue sample should be rinsed separately. Ideally the tissue should be 
quick-frozen in liquid nitrogen, wrapped in aluminium foil and stored in dry ice. 
Label each sample with a field number and code (e.g., H for heart, K for kidney). 
Liquid nitrogen is suitable for working at a field station facility but may be 
impractical and dangerous to use in other field situations. If liquid nitrogen cannot be 
used, wrap tissues in foil, label, and immediately store in dry ice. A small Styrofoam 
cooler with a tightly fitting lid makes an ideal dry ice chamber (Sudia et al., 1970). 
Tissue samples will keep for three days in such a container with dry ice without 
deterioration. Storage time may be increased by replenishing the dry ice every two 
days. Once samples are brought to the laboratory, they can be stored in a freezer at 
-70°C for 6 months. 


6.4 Blood Samples 

Blood samples may be required for biochemical studies of haemoglobin and serum or 
plasma proteins, blood parasite studies, or immunological studies. Although blood 
samples taken in the field may be stored temporarily on wet ice and brought to the 
laboratory for analysis, generally this is not practical. For studies of blood parasites, 
slides can be prepared directly in the field. Haemoglobin or serum samples obtained 
for biochemical analyses should be quick-frozen with liquid nitrogen and stored in a 
Styrofoam cooler with dry ice as described in section 6.3. Once samples are brought 
to the laboratory, they should be transferred to a freezer where they can be kept for 
several months. 

Mammals should be anaesthetized before blood is taken. Ether is an effective 
anaesthetic but an overdose may kill the animal. Sudia et al. (1970) recommended 
carbon dioxide as an anaesthetic. Mammals can be bled from the heart using a 2 or 
5 ml disposable syringe. Clean the skin of the area to be punctured with cotton soaked 
in water and allow to dry. If you require plasma, rinse syringes with heparin to 
prevent coagulation. For such mammals as mice, use a 25 gauge % inch or 5 /s inch 
needle; for mammals to the size of hares, a 23 gauge 1 inch needle; and for mammals 
the size of foxes or racoons, an 18 gauge 1 V2 inch needle. For small rodents and 
bats, some workers prefer to take blood from the orbital sinus. Hold the mammal in 
the left hand with the thumb exerting sufficient pressure behind the eye to cause the 
eye to bulge out slightly. Insert a 50 or 100 microlitre micro-sampling pipette 
(Corning Glass Company, Corning, New York, USA) into the posterior corner of the 
eye and gently rotate it to rupture the capillaries against the bone. Other methods that 
may be used to obtain blood include venipuncture and skin puncture (Miale, 1972). 

Blood from syringes or micro-sampling pipettes should be discharged into vials 
(Sudia et al. , 1970) for freezing and storage. Label each vial with the field number of 
the specimen. After the blood sample has been taken, the mammal can be humanely 
killed (see section 5.1) and prepared as a voucher specimen. 

Slides of blood smears for parasite studies are easily prepared in the field. 
However, there are several precautions to follow when preparing blood slides. Use 
new slides that are either precleaned by the manufacturer or cleaned with soap and 
water and rinsed in 95 per cent alcohol. Wipe with a lint free cloth before using. If 
blood is obtained by cutting a vein, pricking tissue, or cutting the tail, discard the first 
drop from the syringe to get rid of cellular debris. Use the second drop to prepare the 
blood smear. With blood obtained directly from a syringe inserted into the heart or a 
vein, it is not necessary to discard this first drop. Heparinized blood is unsuitable for 
blood smears. 

Place a clean microscope slide on a flat surface and put a small drop of fresh blood 
about 3 cm (1 inch) from the end. The end of a second slide (spreader slide) is placed 
on the slide in front of the drop of blood. The spreader slide should be maintained at 
an angle of about 30 degrees. Pull the spreader slide back onto the drop of blood. 
When the blood has spread the width of the slide, push the spreader slide forward 
with a fast, steady motion. Keep the slide flat and allow the smear to air dry. Once 
fixed or dried, slides can be stored in microscope slide boxes. Label each slide with 
the field number of the specimen. Follow the precautions listed in section 6.1 for 
keeping slides free of dust. Freshly prepared films always stain best but if the slides 
cannot be stained promptly, fix your slides in absolute ethanol or methanol. For 


detailed information on preparing blood smears and blood stains, consult Miale 
(1972) and Faust et al. (1970). 

For most biochemical studies it is essential to separate plasma and red blood cells 
and best results will be obtained by centrifuging. If electricity is not available, several 
suitable hand centrifuges are available (Fisher Scientific Company, 711 Forbes 
Avenue, Pittsburgh, Pennsylvania, USA). Transfer the fresh whole blood samples to 
centrifuge tubes and centrifuge at 3000 rev/min for 20 min. The buff-coloured layer 
(plasma) can be separated from the packed cells by aspiration with a micropipette. 
Transfer the plasma to glass vials or tubes, label, and freeze. If haemoglobin is 
required, red blood cells should be haemolyzed. Wash the cells once with cold 0.85 
per cent saline and then centrifuge at 3000 rev/min for 20 min. Remove the 
supernatant by aspiration with a micropipette. Add two volumes of distilled water to 
the packed cells. After mixing thoroughly, freeze and thaw the solution three times. 
Haemolysates are then centrifuged at 4000 rev/min for 5 min. Transfer the 
supernatant to vials and quick-freeze with liquid nitrogen. 

6.5 Preserving Stomach Contents 

Because the food of such small mammals as rodents and insectivores is usually 
ground completely by the teeth, identification of food items requires microscopic 
techniques. These usually involve preparation of microscope slides of stomach 
contents and comparison with reference slides (Drodz, 1975; Williams, 1962). 
Preparation of these slides can be tedious and time-consuming and the collector may 
wish to preserve the contents of stomachs in the field and analyse the stomach 
material at his/her convenience in the laboratory. 

Dissect stomachs from freshly killed specimens. If a freezer is available, the 
simplest method is to place whole stomachs in plastic bags or vials and freeze them. 
A label with the specimen's field number should be placed in bags or vials. Another 
method is to remove the contents of stomachs and allow the material to dry. Dried 
stomach material can be stored in paper envelopes. In the laboratory the desiccated 
material from stomachs is soaked for 24 h in water and then examined. This 
technique, however, may not be suitable for humid climates. A third method is to 
preserve stomachs and their contents in vials with 80 per cent ethanol. Make an 
incision through the stomach wall to allow the solution to reach stomach contents. 
Record the field number on a sturdy label with waterproof ink and include it in the 

6.6 Preparation of Sperm Slides 

For Study with the Light Microscope 

Generally, testes from freshly killed specimens are used for sperm studies; however, 
Hirth (1960) obtained satisfactory results with smears of the Cauda epididymis of 
specimens preserved in 10 per cent formalin. Spermatozoa can be obtained from the 
cauda epididymis (Fig. 15) or the seminiferous tubules. 


The following technique was used by Forman (1968) to study spermatozoa in 
North American bats. Place the whole testis in a fixing solution consisting of two 
parts 100 per cent methanol, four parts 95 per cent ethanol, one part acetone, two 
parts chloroform and one part 100 per cent propionic acid. To permit rapid fixation, 
cut the testis into 5 to 10 mm squares. Put a short section of seminiferous tubule on a 
slide with one drop of lactophenol-cotton blue stain. Lactophenol-cotton blue consists 
of 20 g phenol crystals, 0.05 g cotton blue (Poirrier's Blue, National Aniline 
Division), 20 ml lactic acid, 40 ml glycerol, and 20 ml distilled water. Dissolve 
liquids by heating under hot water tap, then add the phenol crystals and cotton blue. 
Tease the tubule apart to permit spermatozoa to enter the staining solution. Place a 
cover slip over the slide and seal the edges with balsam. 

Genoways (1973) used the following method for comparing spermatozoa in 
various species of spiny pocket mice of the genus Liomxs. Remove the epididymis 
from freshly killed specimens. Take a small amount of fluid containing sperm and 
suspend in an isotonic solution of sodium citrate (prepared fresh each day). Place a 
few drops of the suspension on a microscope slide and let dry. Then fix the 
spermatozoa with a solution of one part glacial acetic acid and four parts absolute 
methanol for 10 to 15 s. Then stain the sperm slides in a Coplan jar for 30 min with a 
0.02 solution of toluidine blue in water. 

For Study with the Scanning Electron Microscope (SEM) 

Several recent studies have shown that there is great potential for studying the 
morphology of mammalian spermatozoa with the SEM. If such studies are 
contemplated, fix the testes in gluteraldehyde in the field. Remove the testes from 
freshly killed specimens and wash in physiological saline. Place the testes in a vial 
with a 1 or 2 percent solution of gluteraldehyde. Gluteraldehyde is sold commercially 
as a 25 per cent solution in water (J. T. Baker Chemicals, Canadian Laboratory 
Supplies, Toronto, Canada). The testes should be cut into 5 to 10 mm squares to 
permit rapid fixation. In the laboratory tissue is washed and centrifuged to remove the 
fixative and sperm samples are freeze-dried for later examination with the SEM 
(Gould et al., 1971). 

6.7 Fixing Tissues for Histological Study 

If histological studies are contemplated, tissues should be fixed from various organs 
immediately after the specimen is killed. Proper fixation is essential for preventing 
any post-mortem deterioration in tissues. In section 5 we discussed the use of 
buffered neutral formalin for fixing entire mammal specimens. Buffered neutral 
formalin is also an excellent general purpose fixative for histological work and it can 
be used to fix many different tissues. For certain histological procedures, however, 
other fixatives may be desired because they penetrate tissues more rapidly than 
neutralized formalin and may render tissues more easily stained by certain 
histological dyes. Tissues can be left in some fixatives (e.g., buffered neutralized 
formalin) for several months; with other fixatives (e.g., Bouin's), tissues must be 
transferred to alcohol immediately after fixing. Three of the more widely used 


fixatives are described here. For more information on fixatives and histological 
methods, consult Luna (1968). 

Bouin's Solution 

*picric acid, saturated aqueous solution 750 ml 

37-40% formalin 250 ml 

glacial acetic acid 50 ml 

*if not stored in an aqueous solution, picric acid is highly volatile. 

Bouin's solution is frequently used for fixing gastrointestinal tracts, reproductive 
organs, endocrine glands, and brain tissue. It is a rapid fixative and will fix blocks of 
tissues in 4 to 12 h depending on their size. Once tissues are fixed, however, they 
must be washed in two or three changes of 40 per cent alcohol for 4 to 6 h to remove 
all picric acid. If picric acid is not removed, tissue will undergo deleterious changes. 
After washing tissues, store in 70 per cent alcohol. 

Alcohol-Formalin-Acetic Acid Solution (AFA) 

37-40% formalin 10 ml 

alcohol, 80% 90 ml 

glacial acetic acid 5 ml 

A good fixative for rapid fixation, AFA solution has been used for reproductive 
and gastrointestinal tracts. Small pieces of tissue (2 mm thick) will completely fix in 
4 to 6 h. AFA is not suitable for tissue storage and the fixed tissue should be 
transferred to 70 per cent alcohol. 

Formalin-Sodium Acetate Solution 

37^0% formalin 100 ml 

sodium acetate 20 g 

tap water 900 ml 

This is an excellent fixative in which to store gross blocks of tissue (e.g., whole 
brains of such small mammals as bats, shrews, or mice). 

7. Shipping Specimens 

7.1 Methods for Shipping 

Fluid-preserved Specimens 

Once properly fixed, specimens can be shipped "damp packed", that is, only a small 
amount of 10 per cent formalin is added to the packing material in the container to 
keep specimens moist in transit. About 0.25 L (0.5 pt) of fluid is sufficient for a 


container of 3.8 L (1 gal.). Cotton wool or newspaper should be added to each 
package, both to limit the movement of specimens and to retain the dampness. 
Wide-mouthed plastic jars are the most suitable shipping containers but if they are not 
available, plastic bags may be used if they are well sealed. Glass jars are the least 
desirable type of container but can be used if properly packed in a strong container 
and insulated with at least 5 cm (2 inches) of wadded paper or similar material on all 
sides of each jar. If plastic bags are used, it is recommended that the specimens, 
cotton wool, and fluid be placed in one bag and sealed by tying the top of the bag; be 
certain to remove excess air from the bag. Then place the sealed bag inside a second 
bag, which is in turn sealed by careful tying. The double plastic bag should then be 
placed in a light, strong container. A tin can with a lid is ideal. If the tin is too large, 
fill the remaining space with cotton wool or crushed paper. The container should be 
large enough to absorb the fluid should the bags leak. Seal tin cans with adhesive tape 
to ensure they will not open or leak in transit. Generally, packages should be kept 
small enough so that they may be shipped by ordinary parcel post. 

Skins and Skulls 

It is advisable to ship study skins in small, strong containers, preferably plywood 
boxes, although strong cardboard boxes may be used. Begin packing by placing a 
layer of about 5 cm (2 inches) of cotton wool or similar material on the bottom, 
followed by a layer of dried skins. Add another layer of cotton wool and repeat the 
process with another layer of skins, until the box is almost full. Allow a top layer of 
about 5 cm (2 inches) of cotton wool. 

If thoroughly dried and free of insects and insect eggs, skulls may be packed in the 
same box with skins. Otherwise, ship them in a separate strong container that cannot 
be smashed or crushed under normal shipping conditions. If the skins are likely to be 
in transit for longer than a few days, add moth balls (paradichlorobenzine) to the 
container of skins, to protect them from moths or dermestid beetle eggs. Skulls and 
skeletons should not be placed together in a container with moth balls because the 
chemical released may inhibit the activity of dermestid beetles that are used by most 
museums to clean skulls and skeletons. 


Skeletons must be dried before shipping. You can pack small skeletons in the same 
container used for the skins if moth crystals have not been added to the container. 
Place the skeletons on the bottom of the container, then cover with alternate layers of 
cotton and skins. Heavy bones from large mammals should be packed separately in 
sturdy wooden crates. If a large skeleton is properly dried, place it in a plastic bag just 
before shipping to reduce the odour. Collectors shipping large skeletons should notify 
the receiving museum prior to shipping. 


Pages of the original field catalogue, field notes, and any pertinent topographic maps 
should be mailed separately from specimens as first class or air mail and registered if 
from a country where mail service is inadequate. 


7.2 Import/Export Regulations 

Convention on International Trade in Endangered Species 

Canada, the United States, and approximately 40 other countries have signed a 
Convention on International Trade in Endangered Species. The Convention prohibits 
all imports and exports of protected species except under permit. These regulations 
apply to all international shipments and, in addition to living mammals, to parts and 
derivatives of endangered species. Permit requirements must be observed for any 
shipment to or from a member country, even if the other country involved is not a 
member of the Convention. Species protected are listed in three Convention 
appendices. Appendix I of the Convention lists species threatened with extinction; 
Appendix II lists species that must be monitored to avoid the threat of extinction; and 
Appendix III lists species placed there by individual countries to reinforce domestic 
conservation measures. All shipments of species listed in Appendix I of the 
Convention require two permits — one from the importing country and another from 
the exporting country. Export permits must be issued from the country of origin for 
species listed in Appendix II. International shipments of Appendix III species require 
either an export certificate from the country that listed the species or a certificate of 
origin from any country. 

Shipments to Canada 

All shipments to Canadian museums should be labelled with a declaration form that 
indicates the total number of specimens, the general kind of specimens (bats or 
rodents), and that specimens are preserved (e.g., 100 preserved bats) rather than live 
or frozen. Shipments of living mammals must have Canadian Department of 
Agriculture, Health of Animal Branch permits. 

The shipment should also be marked "No Commercial Value (NCV), For 
Scientific Research, No Endangered Species". If the collector insures the shipment 
for more than $150.00, he must complete special Canada Customs invoice forms for 
shipments from outside Canada; these invoice forms are available from museums 
upon request. Shipments should also be labelled "In Bond to Destination" which 
ensures that the shipment can only be inspected by customs officials at the destination 
and not at border points. Notify the museum of a forthcoming shipment and, when 
sending specimens by air freight, forward a copy of the Airway Bill. 

Permits to import or export species listed under the Convention on International 
Trade in Endangered Species may be obtained from the Canadian Wildlife Service in 
Ottawa or any provincial or territorial wildlife headquarters. 

Shipments to the United States 

Packages should be clearly marked on the outside with the name and address of the 
shipper and of the consignee and give an accurate statement of the contents (species 
and numbers of each species). Label the package "Scientific Specimens; No 
Endangered Species; No Commercial Value". 



Nonendangered mammals and parts thereof from countries other than Canada may 
enter or leave the US without a permit only through eight designated ports of entry: 
Los Angeles, San Francisco, Miami, Honolulu, Chicago, New Orleans, New York, 
and Seattle. Species protected under the Convention on International Trade in 
Endangered Species and the US Endangered Species Act must enter the US through 
these same eight cities. Mammals other than endangered species may be imported, 
for final destination only, into Alaska through Anchorage, Fairbanks, Juneau, or Tok 
Junction; or into either Puerto Rico or the Virgin Islands through San Juan, Puerto 

Specimens obtained legally in Canada may enter the USA without permit through 
any of 25 border points if no endangered species are included. Contact the Division of 
Law Enforcement, US Fish & Wildlife Service, Washington, D.C., USA for a list of 
these border points. If no endangered species are included, mammals collected 
legally in Mexico may enter the US without permit through any of seven border 
points: Calexico or San Diego-San Ysidro, California; Nogales or San Luis, Arizona; 
Brownsville, El Paso, or Laredo, Texas. 

Scientific specimens of nonendangered species may enter or leave the US at 
nondesignated ports of entry under permit. 


To import specimens through the designated ports of entry previously given, the 
collector needs a valid collecting permit and an export permit (if required by the 
country of origin) and a Declaration of Importation of Fish & Wildlife (Form 3-177). 
You must file copies of these documents with the District Director of Customs at the 
port of entry. 

For importation of mammal specimens through nondesignated ports of entry, you 
must satisfy the previously given requirements and have a permit for the 
nondesignated port from the U.S. Fish & Wildlife Service. 

Permits for importing specimens of mammals listed in the Convention on 
International Trade in Endangered Species of the federal Endangered Species Act can 
be obtained from the Federal Wildlife Permit Office, U.S. Fish & Wildlife Service, 
Washington, DC 20240, USA. Note that although some species of mammals are 
listed in both the convention and the Endangered Species Act, the two lists do not 
contain identical species. 

A permit is required to import living material, including tissue cultures, cell lines, 
and blood and serum that could serve as a vector for pathogenic organisms. Apply to 
the U.S. Department of Agriculture, Animal and Plant Health Inspection Service, 
Veterinary Services, Washington, D.C., USA for this permit. 


Packages or containers in which specimens are transported must be clearly marked on 
the outside with the same information as described for shipments from outside the 
US. Because it is illegal under the Lacey Act to import or ship in interstate commerce 
any wildlife taken in violation of state or local laws, scientific collectors must 
familiarize themselves with appropriate laws for the state or region concerned. 


8. Public Health Hazards 

It is beyond the scope of this manual to discuss the numerous potentially dangerous 
diseases that are carried by wild mammals. Several useful references are given in the 
bibliography. Because diseases are often local in their distribution, it is impossible to 
list specific diseases that the collector may encounter. As a result, you are advised to 
familiarize yourself with the disease hazards that may be present in your local area. 
Consult your physician about vaccinations that may be taken against diseases in your 
area, for example, rabies, plague. All collectors should be vaccinated against tetanus. 
Take care to avoid being bitten when handling live mammals and use leather gloves 
to protect the hands. If you are collecting species that are susceptible to rabies (bats, 
foxes, mongooses, skunks), regard all captures as potentially rabid. Some common 
sense precautions to reduce health hazards can be followed when preparing 
specimens. Wear rubber gloves when dissecting mammals and, for additional 
protection, use disposable paper face masks. Avoid contact with urine and faeces, 
which often contain infectious agents. Dissecting instruments should be cleaned and 
disinfected with Dettol or a 3 per cent phenol solution after use. Immediately wash 
any cuts or abrasions with soap and treat with an antiseptic. Use similar precautions 
when handling road kills. If symptoms such as chronic respiratory distress, 
influenza-like sickness, swelling of lymph nodes, high temperature, vomiting, or 
diarrhoea occur in conjunction with the handling of specimens, they should be 
regarded with suspicion and medical advice should be sought. 

9. Acknowledgements 

We gratefully acknowledge the assistance and encouragement of staff of the 
Department of Mammalogy, ROM, during the preparation of the manuscript. The 
following staff deserve special thanks: J. R. Tamsitt for his advice on special 
techniques and for critically reviewing the manuscript, Judith Eger for critically 
reviewing the manuscript, Sophie Poray for drawing some of the illustrations, and 
Nancy Grepe for typing the numerous revisions of the manuscript. Other illustrations 
were done by Anker Odum and Julian Mulock, Exhibit Design Services, ROM. The 
cover drawing is by Peter Buerschaper, Exhibit Design Services, ROM. Many of the 
techniques described in the manual for preparing study skins were developed by John 
G. Williams. 


10. Selected Bibliography 

General Reference 


1965 Methods of collecting and preserving vertebrate animals. 4th ed. rev. National Museum of 
Canada, Bulletin 69:1-199. 


1968 Instructions for collectors no. 1. Mammals (non-marine). 2nd ed. British Museum (Natural 
History) Publication 665:1-55. 

brown, j c and d m stoddart 

1977 Killing mammals and general post-mortem methods. Mammal Review 7:63-94. 

deblase. a f and r e martin 

1974 A manual of mammalogy with keys to the families of the world. Dubuque, W. C. Brown. 
329 pp. 

Giles. R.H.. Jr., ed. 

1971 Wildlife management techniques. 3rd ed. rev. Washington, DC, The Wildlife Society. 633 

HALL, e r 

1962 Collecting and preparing study specimens of vertebrates. University of Kansas, Museum of 
Natural History, Miscellaneous Publication 30:1-46. 


1966 Biological techniques; collecting, preserving, and illustrating plants and animals. New York, 
Harper & Row. 525 pp. 


1965 Collecting bat specimens for scientific purposes. Toronto, Dept. of Mammalogy, Royal 
Ontario Museum. 8 pp. 


1977 A guide to the management of recent mammal collections. Camegie Museum of Natural 
History, Special Publication 4:1-106. 



1976 Supplies and suppliers for vertebrate collections. Museology, Texas Tech University, 

Locality Data 


1965 More on locality data and its presentation. Systematic Zoology 14:64—66. 

REIMER. w j 

1954 Formulation of locality data. Systematic Zoology 3: 138-140. 

Mammal Collecting 


1972 A live trap for pocket gophers. Journal of Wildlife Management 36:1320-1322. 



1968 Bats and bat banding. United States Dept. of the Interior, Bureau of Sport Fisheries and 
Wildlife, Resource Publication 72:1^47. 


1965 The Manitoba trappers' guide. Rev. ed. Winnipeg, Dept. of Renewable Resources and 
Transportation Service. 122 pp. [good review of trapping and skinning methods] 


1974 A conical pitfall trap for small mammals. Northwest Science 48:102-103. 


1962 Game biology and game management: a laboratory manual. Minneapolis, Burgess. 143 pp. 
[trapping methods] 


1978 A collapsible bat-trap and a comparison of results obtained with trap and with mist 
nets. Austrialian Wildlife Research 5:355-362. 


1974 An improved trap for bats. Journal of Mammalogy 55:475-477. 

1976 Collecting techniques. In Baker, R.J. et al., eds.. Biology of bats of the New World family 
Phyllostomatidae Part I. Texas Tech University, Musuem, Special Publications 10:71-88. 

twigg, G I 

1975 Catching mammals. Mammal Review 5:83-100. 


1972 Relative efficiencies of four small mammal traps. Journal of Mammalogy 53:868-873. 


1977 A field test of six types of live-trap for african rodents. Zoologica Africana 12:215-223. 


1975 Karyotype of the eastern mole (Scalopus aquaticus), with comments on the karyology of the 
family Talpidae. Journal of Mammalogy 56:902-905. [description of a live trap for moles] 



1945 Color changes in mammal skins during preparation. Journal of Mammalogy 26:128-132. 

QUAY, w M 

1974 Bird and mammal specimens in fluid — objectives and methods. Curator 17:91-104. 

setzer. H w 

1968 Directions for preserving mammals for museum study. United States National Museum, 
Smithsonian Institution, Information Leaflet 380:1-19. 

smith. D A 

1968 On the use of tapered wires for study skins of small mammals. Journal of Mammalogy 

Public Health 


1972 Possible health hazards associated with the collection and handling of post-mortem zoological 
material. Mammal Review 2:43-54. 


McDIARMID, A , ed. 

1969 Diseases in free-living wild animals. Zoological Society of London, Symposium 24. 
London, Academic Press. 332 pp. 



1977 Index (o U.S. federal wildlife regulations. Lawrence, University of Kansas, Museum of 
Natural History. 278 pp. 


1973 Convention on international trade in endangered species of wild flora and fauna. Canadian 
Wildlife Service reprint. Ottawa, Environment Canada. 20 pp. 

EDWARDS. S R and L D GROTTA. eds. 

1976 Systematics collections and the law. Lawrence, Association of Systematics Collections. 29 


1976 Federal regulations pertaining to collection, import, export and transport of scientific 
specimens of mammals. Journal of Mammalogy 57(2) suppl.: 1-9. 


1976 State laws as they pertain to scientific collecting permits. Museology, Texas Tech 
University, 2:1-81. 


1977 Endangered species legislation in Canada. In Mosqin, T. and C. Suchal, eds., Canada's 
threatened species and habitats. Canadian Nature Federation, Special Publication 6: 19-21. 

Reproductive Data 


1955 Embryo resorption and placental scar formation in the rat. Journal of Mammalogy 


1956 Morphology of the reproductive tract. In Parkes, A. S., ed., Marshall's physiology of 
reproduction, vol. 1, pt. 1. London, Longman Green, pp. 43-155. 


1950 Determining fecundity in male small mammals. Journal of Mammalogy 31:433^*36. 


1967 Correlation of embryo and placental scar counts of Peromxscus manic ulaius and Microtus 
ochrogasier . Journal of Mammalogy 48:317-319. 

Special Techniques 


1970 The role of karyotypes in phylogenetic studies of bats. In Slaughter, B. H. and D. W. Walton, 
eds.. About bats: a chiropteran biology symposium. Dallas, Southern Methodist University 
Press, pp. 303-312. 


1975 Analysis of stomach contents of small mammals. //; Grodzinski, W., R. Z. Klekowski, and A. 
Duncan, eds., Methods for ecological bioenergetics. IBP handbook 24. Oxford, Blackwell 
Scientific, pp. 337-341. 



1970 Craig and Faust's clinical parasitology. 8th ed. Philadelphia, Lea and Febiger. 890 pp. 


1968 Comparative gross morphology of spermatozoa of two families of North American 
bats. University of Kansas Science Bulletin 16:901-928. 


1973 Systematics and evolutionary relationships of spiny pocket mice, genus Liomvs. Texas Tech 
University, Museum, Special Publications 5:1-292. [technique for preparing rodent sperm 


1971 Mammalian gametes: a study with the scanning electron microscope. Chicago, Proceedings 
of the Fourth Annual Scanning Electron Microscope Symposium, part 1, pp. 289-295. 

hirth. M F 

1960 The spermatozoa of some North American bats and rodents. Journal of Morphology 


1968 Simple field techniques and laboratory method for recovering living ticks (Ixoidea) from 
hosts . Journal of Parasitology 54 : 1 88- 1 89 . 


1968 Manual of histological staining methods of the Armed Forces Institute of Pathology. 3rd 
ed. New York, McGraw Hill. 258 pp. 


1971 Essentials of parasitology. Dubuque, W. C. Brown. 305 pp. 


1972 Laboratory medicine: hematology. 4th ed. Saint Louis, C. V. Mosby. 1318 pp. 


1971 Direct and repeatable bone marrow chromosome preparations from living mice. Stain 
Technology 46:285-288. 


1970 Collection and processing of vertebrate specimens for arbovirus studies. Washington, 
United States Dept. of Health, Education, and Welfare, Public Health Service. 65 pp. 

watson. g e and a b amerson. Jr. 

1967 Instructions for collecting bird parasites. United States National Museum, Smithsonian 
Institution, Information Leaflet 477:1-12. 


1962 A technique for studying microtine food habits. Journal of Mammalogy 43:365-368. 


1 1 . Appendices 

11.1 Appendix 1 

Checklist of Field Equipment 

For a listing of suppliers of field equipment, consult Dowler and Genoways (1976). 

Collecting Bats 

mist nets— 12.5 m x 2 m (42 ft x 7 ft), 9 m x 2 m (30 ft x 7 ft) 

hand net 

machete — for cutting poles 

headlamp and batteries — for working in caves or checking nets at night 


long forceps (25 cm; 10 inches) — for collecting bats in attics or rock crevices 

cloth collecting bags 

leather gloves — for handling live specimens 

Collecting Other Mammals 

museum special snap traps 

Victor rat-traps 

Sherman live traps 

juice or coffee cans — for pitfall traps 

cloth collecting bags 

bait — peanut butter, rolled oats, dried fruits, nuts 

coloured, plastic flagging tape — for marking trap sites 

compass and topographic maps of study areas 

tobacco tin or plastic jar — for carrying bait 

knapsack — for carrying traps 

cord, wire — for tying traps to stakes 

guns, ammunition 

leather gloves 

cotton wool — nesting material for live traps 

Measuring and Preparing Specimens 

weight scales (metric) — Pesola spring balances 

metal tape-measure (metric), plastic ruler (metric) 

catalogue sheets 

field tags, waterproof paper 

hand lens — for examining external genitalia and internal sex organs 

notebook — for field notes 

pencils, pen, and waterproof ink 


small waterproof tags — for labelling vials 
small vials (\-4 dram size) — for embryos and parasites 
borax or magnesium carbonate powder 
gauze or cheesecloth bags — for skulls and skeletons 
disinfectant (Dettol or 3% phenol) — for sterilizing dissecting instruments 
sawdust or cornmeal 
toothbrush— for brushing fur 
disposable gloves 

chloroform or ether — for killing small mammals 
Euthanyl (sodium pentobarbital) — for killing larger mammals 
70 per cent ethanol — for ectoparasites, cleaning blood off fur 
40 per cent formaldehyde, salts for neutralizing formalin 
hypodermic syringe and needles — for injecting specimens 
plastic bags, various sizes 
moth balls (paradichlorobenzene crystals) 
cotton — for stuffing study skins 

wire for tails of skins — no. 12, 16, 18, 20, 22, 24, 25 gauge (preferably Monel wire) 
glass-headed pins, various sizes 
needles, various shapes and sizes 
pinning board, corkboard, or wallboard 
assortment of cardboard or art board — for flat skins 
white linen thread, sizes 40, 30, 25 
white cotton thread, sizes 50, 60 
pliers, wire cutters 

drying chest — for storing drying skins 
scalpel handle, no. 3 and no. 4 sizes 

scalpel blades, no. 20 size (for no. 4 handle) and no. 10 size (for no. 3 handle) 
scissors, one small pair with sharp points, one large pair 
forceps, one pair with fine tips, one pair with blunt tips 
bone cutters 

skinning knife — for large mammals 
sharpening stone 

table salt (NaCl) — for drying large skins 
burlap sacks — for skulls and bones of large mammals 
masking tape 

electrical or adhesive tape — for securing lids on glass vials and taping lids of mailing 

11.2 Appendix 2 

Karyotype Kit for 120 Small Mammals 

Sodium citrate 15 packages of 0.5 g quantity 

5 packages of 0.75 g quantity 
1 hand centrifuge 
24-15 ml glass centrifuge tubes 
24-1 ml tuberculin syringes 


2-2 ml Luer Lock syringes 

4-5 ml Luer Lock syringes 

1 doz. no. 26 syringe needles (Vi inch) 

1 doz. no. 24 syringe needles (1 inch) 

1 doz. no. 21 syringe needles (1 l /z inches) 

250 (1 box) disposable glass pipettes (5.75 inches) 

48 rubber pipette bulbs 

1 plastic centrifuge rack 

2 centrifuge tube cleaning brushes 

5 gross (720) precleaned microscope slides with frosted ends (75 mm x 25 mm size) 

3 pints absolute methanol 
2 pints glacial acetic acid 
4-10 mg vials Velbe 

2-100 ml serum bottles of bacteriostatic sodium chloride 

1-125 ml nalgene Erlenmeyer flask — for sodium citrate solution 

9 slide boxes (100 slide capacity) 

1-100 ml nagrene Erlenmeyer flask — -for fixative 

1-50 ml nagrene graduated cylinder for measuring fixative 


silica gel crystals 

grease pencil — for labelling centrifuge tubes 

100 ml distilled water 

For Staining 

500 ml xylol 

500 ml acetone 

100 ml Giemsa stain 

100 ml permount 

5-1 oz. boxes cover slips (no. 1, 24 mm x 50 mm) 

6 Coplan jars 

1-50 ml polypropylene graduated cylinder — for measuring stain 
1 polypropylene funnel, filter paper — for filtering stain 


ISBN 0-88854-255-0 
ISSN 0082-5093