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The scientific value of avian research specimens is immense, but the accumulation rate of this resource is too low to meet either present or future needs. This may be due, in part, to the fact that few students are currently being taught to prepare specimens. Modern specimen preparation is a routine but detailed and meticulous process in which comparatively few are expert. I summarize methods for obtaining bird specimens and preserving them both for the short term and for the long term as high quality scientific research specimens. The preparation method outlined preserves skin, partial skeleton, stomach contents and two duplicate tissue samples for every specimen, maximizing the scientific usefulness of each bird. The resulting skins and skeletons augment current samples, simultaneously increasing the sample sizes available for studies involving either type of specimen. These methods allow a diverse array of data to be taken from every individual, and are thus suitable for general preparation or focused, single-species research projects. These archival quality methods assure that, if prepared as outlined, the skin and skeleton specimens possess a useful life of half a millennium or more. I suggest that this is an unparalleled opportunity to make a personal, signed, long-term contribution to science with relatively little time investment.
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OBTAINING, PRESERVING, AND PREPARING BIRD SPECIMENS
K
EVIN
W
INKER
University of Alaska Museum
907 Yukon Drive
Fairbanks, Alaska 99775 USA
Abstract.—The scientific value of avian research specimens is immense, but the accumulation
rate of this resource is too low to meet either present or future needs. This may be due, in
part, to the fact that few students are currently being taught to prepare specimens. Modern
specimen preparation is a routine but detailed and meticulous process in which compara-
tively few are expert. I summarize methods for obtaining bird specimens and preserving
them both for the short term and for the long term as high quality scientific research spec-
imens. The preparation method outlined preserves skin, partial skeleton, stomach contents
and two duplicate tissue samples for every specimen, maximizing the scientific usefulness of
each bird. The resulting skins and skeletons augment current samples, simultaneously in-
creasing the sample sizes available for studies involving either type of specimen. These meth-
ods allow a diverse array of data to be taken from every individual, and are thus suitable for
general preparation or focused, single-species research projects. These archival quality meth-
ods assure that, if prepared as outlined, the skin and skeleton specimens possessa useful life
of half a millennium or more. I suggest that this is an unparalleled opportunity to make a
personal, signed, long-term contribution to science with relatively little time investment.
OBTENIENDO, PRESERVANDO Y PREPARANDO ESPECI
´MENES DE AVES
Sinopsis.—El valor cientı´fico de especı´menes de aves para investigacio´n es inmenso, pero la
tasa de acumulacio´n es muy baja para satisfacer las necesidades presentes o futuras. Esto se
puede deber en parte al hecho de que pocos estudiantes esta´n siendo entrenados apreparar
especı´menes. La preparacio´n moderna de especı´menes es un proceso rutinario pero deta-
llado y meticuloso en que hay pocas personas expertos. Resumo los me´todos para obtener
especı´menes de aves y preservarlos tanto en corto tiempo como para especı´menes de inves-
tigacio´n de gran calidad cientı´fica a largo tiempo. El me´todo de preparacio´n descrito pre-
serva la piel, esqueleto parcial, contenido estomacal y dos muestras de tejido para cada
especı´men, maximizando la utilidad cientı´fica de cada ave. Laspieles y esqueletos resultantes
aumentaron las muestras presentes, aumentando simulta´neamente los taman˜os de muestras
disponibles para estudios requiriendo cualquier tipo de especı´men. Estos me´todos permiten
una obtener una diversidad de datos de cada individuo, y por lo tanto son apropiadospara
la preparacio´n general o enfocada, y de proyectos de una sola especie. Estos me´todos de
calidad de archivo aseguran que, de prepararse tal como se describe, los especı´menes de
piel y esqueletales poseen una vida u´til de al menos 500 an˜os. Sugiero que esta es una
oportunidad sin paralelos para producir una contribucio´n personal de larga duracio´n fir-
mada a la ciencia con una inversio´n de tiempo relativamente corta.
The importance of specimens to the science of ornithology and the
conservation of biodiversity would be difficult to overstate (see Remsen
1995; Winker 1996, 1997). Yet, at a time when the need for avian speci-
mens has increased across a broad range of studies, it seems that few are
being added to collections, even though the world’s bird collections in
aggregate represent a grossly inadequate documentation of extant birds
(e.g., Goodman and Lanyon 1994, Winker 1996). This may be due in part
to the fact that few students are being trained to prepare specimens (cf.
Rogers and Wood 1989). Given the state of literature on the subject
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(mostly outdated) and the changing nature of specimen preservation,
even a motivated novice would have a difficult time learning to produce
high quality research specimens. While still containing some usefulpoint-
ers on various aspects of specimen acquisition and preparation, earlier
guides to specimen preparation (e.g., Swainson 1836, Coues 1874, Ridg-
way 1891, Chapin 1946, Blake 1949, Hall 1962, Proctor and Lynch 1993)
are outdated and not sufficiently comprehensive. Johnson et al. (1984)
provided an excellent contribution emphasizing tissue preservation and
updating older preparation methods with important new variations, but
was neither comprehensive nor of sufficient detail for the novice.
With a little foresight, understanding, and practice, field ornithologists
can preserve the specimen research material that is needed today and
that will be highly useful in the coming centuries. Because a specimen is
extremely useful for many types of research, and because proper prepa-
rations can last for centuries, it behooves all ornithologists to promote
the proper collection, preparation, and preservation of bird specimens.
It should be remembered that the populations these specimensdocument
represent renewable resources.
Here I outline practical methods for obtaining, preparing, and pre-
serving bird specimens that maximize the usefulness of each bird. These
methods are intended to be useful to the novice for learning to prepare
and preserve scientific bird specimens, but also to the professional, who
might find new ideas and methods that increase the research value of
each specimen. The methods outlined here are for general purpose spec-
imens, and are therefore not all-encompassing. Although more intricate
than many previous methods, these methods are not overly cumbersome,
given the long-term scientific value of the products, and they have proven
suitable for general preparation procedures. These methods are also suit-
able for focused research projects examining broad suites of morpholog-
ical and genetic characters.
PHILOSOPHICAL APPROACH
Today, specimen-based ornithologists practice a wide range of prepa-
ration procedures (e.g., Johnson et al. 1984). Many of these methods are
specialized for the immediate needs of the researcher, and workersoften
needlessly discard material desperately needed in the broader scientific
community simply because it is too much time or trouble to properly
preserve portions of the specimen they don’t use. This situation applies
even to some dedicated collectors, who will throw away skin, skeleton, or
tissues because they personally don’t use that particular type of bird sam-
ple. As a general methodology this is difficult to justify.The skin, skeleton,
and tissues of every bird can and should be preserved whenever time
allows.
At the root of the method presented here is a ‘‘total evidence’’ ap-
proach: the more evidence that we are able to bring to bear on a question,
the more confidence we can have in the answer or results derived from
the specimen material. This suits not only individual research projects,
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but also brings to the present and future research community mostof the
components of a dead bird that we know can be useful, whether for
traditional studies of phenotype and genotype, or for stable isotope and
contaminant analyses, or for future methods (and questions) as yet un-
developed. From the specimen use requests I have seen in the past few
years, it is certain that museum collections will continue to provide an
increasingly broad research community with immensely valuable historic
research material for an amazing variety of questions. Preserving speci-
mens with this future in mind significantly enhances both our science
and our personal contributions to it—particularly at a time widely viewed
as a watershed period of biodiversity loss and climate change.
The preparation method presented here emphasizes a combination
skin-skeleton specimen that enables the skins to enhance existing samples
of skin collections. Thus, skins without bills (often called ‘‘shmoos’’ or
‘‘muppets’’) are de-emphasized, because these remove the important
ramphotheca from the skin, making these skins incomparable (orincom-
pletely comparable) to previously existing specimens. Consequently, the
skeletons produced using this method are partial, in that the full skull is
not preserved. Although there are many differences, this method is most
similar to Method 2 outlined by Johnson et al. (1984).
Preserving both skin and skeleton from each individual whenever pos-
sible is fully justified. The need for skeletal material has been emphasized
for at least 140 yr (Newton 1860), and the now outdated world inventories
of skeletal specimens (Wood et al. 1982, Wood and Schnell 1986) had a
profound influence on complete skeleton preparations. But plumage
probably reveals more of the developmental similarities and differences
among populations than skeletons, and is also useful for examining age-
related differences among individuals. Although we are too ignorant at
present to use and decipher all of the information available in plumage
characteristics, tools such as reflectance spectrophotometry are providing
increased resolution and rigor when studying plumage characteristics
(e.g., Graves 1997). Skeletal characters are useful at multiple levels and
are not subject to the fading and wear that plague plumage studies. Fur-
thermore, series of skeletons are as important as series of skins for incor-
porating individual variation into data sets, and sample sizes of bothskins
and skeletons are maximized by preserving both from each specimen.
Finally, and importantly for focused projects, studies of evolution are im-
proved when broad suites of genetic and morphological characteristics
can be assessed from the same individuals (see also Johnson et al. 1984).
What should be preserved and prepared
?—The world’s systematics collec-
tions are so weak in so many areas, and so few specimens exist to docu-
ment today’s populations, that almost every dead bird has scientific value.
In ongoing morphometric studies I have found it difficult or impossible
to assemble 30 individuals of each age and sex class of many common
eastern North American passerines. This stems in part from the fact that
older specimens have fewer data than modern preparations (most lack
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data on degree of cranial ossification and body mass, for example), but
it is also due to a general paucity of specimens.
As members of a research community concerned with increasing our
knowledge of birds in the face of accelerating global changes, we will
achieve much greater success by pulling together than by blindly pursuing
only our own narrow research agendas. A certain degree of altruism ben-
efits us all, and, importantly, provides immense benefits to future re-
searchers. If preceding generations of ornithologists had not made gen-
eral collections and preserved them, our science would be greatly impov-
erished today. Whenever we go into the field we might be the onlytrained
ornithologists to visit an area in decades. Consequently, anyone in the
field should be supported and encouraged to sample broadly: few areas
are adequately sampled, and temporally adequate samples essentially do
not exist. Depositing specimens in research collections that have wide
accessibility is an important corollary of this philosophy.
With the growth of the animal rights movement has come an increas-
ingly strident opposition to the active collection of birds. Personalbeliefs
in this regard are to be respected, but I have argued elsewhere that in
many cases this opposition is misguided, often arising from an ignorance
of biological principles and the value of the specimens themselves (Wink-
er 1996). Here I wish to emphasize that one’s beliefs about how animals
should be treated (i.e., allowed to die) should have little or no effect
upon one’s dedication to preserving specimens for science. The avian
carnage caused by domestic cats, glass windows, automobiles, and com-
munications towers is far greater than the comparatively tiny numbers of
birds collected by scientists each year (Banks 1979, Churcher andLawton
1989, Klem 1990b). It seems a terrible breach of ethics to allow this car-
nage to go to waste when the specimens are often of great scientificvalue.
And, in fact, probably most specimens added to museum collections today
come from the salvage of birds found dead. A good salvaged specimen is
one less that might have to be collected. Thus, individuals ethically op-
posed to the collection of birds might be looked upon to become some
of the best and most active preparators, wishing to see something bene-
ficial resulting from the many avian casualties arising from nonscientific
anthropogenic influences.
The bird bander represents an important category of avian researcher
whose activities provide unique opportunities to obtain and preserve re-
search specimens. Unfortunately, these opportunities are rarely exploited.
Although the work of banders has benefitted enormously from informa-
tion derived from bird collections, and though banding regularly gener-
ates dead birds through accident, there is little return to collections from
banding efforts. I am a bird bander, and have learned throughexperience
that the quality of my work is greatly improved through a combinationof
banding and selective collecting. If a bander is really motivated to learn
about birds, rather than simply deriving enjoyment from capturing and
banding them, then it is a very short step to discovering how much more
can be learned through judicious collection.
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I regularly use skins 100–150 yr old in my skin-based research. Although
the purposes for which I am using them are probably different from the
reasons for their initial collection, their scientific and physical integrity
remain as great or greater than the day they were prepared. In fact, their
scientific value often increases with time through label annotations and
biotic changes occurring among populations and environments. Judging
from old books, which are also products of plant fiber and animal skin,
we can realistically expect the archival-quality skin preparations outlined
here to last for at least 500 yr or more if housed under properconditions.
I can think of no other comparatively short term time investment with
such staggering long term scientific impact.
PERMITS
The collection and possession of bird specimens is generally regulated,
and many types of permits are often required, depending on the country
or countries where these activities take place. The long-term scientific
value of a specimen is dependent upon its being accessible to the scien-
tific community. The scientific community abides by all permitting re-
quirements, and therefore requires that specimens be legally acquired
and possessed. As scientists, it is our duty to obtain permits that enable
us to perform our jobs to the best of our abilities, but which at the same
time provide adequate safeguards to ensure that wildlife resources remain
available for continued existence and for use and enjoyment by society
at large. There have been conflicts between scientists and permitting
agencies in obtaining permits agreeable to both parties (for some discus-
sion, see Remsen 1995, Winker 1996). However, it is usually possible to
work with permitting agencies to achieve mutually agreeable permits.
Make every attempt to provide permit-granting agencies with the infor-
mation they need to make informed decisions. In general, scientific col-
lecting does no harm to avian populations (see Banks 1979, Remsen1995,
Winker et al. 1991, Winker 1996). If one wishes only to salvage dead birds,
special salvage permits are usually issued, and are generally easier to ob-
tain than collecting permits.
An important category of permit is that allowing the international im-
port and export of specimens. In some countries the collecting permit
also serves as a valid export permit, while in others two separate permits
are needed. In the U.S., a national U.S. Fish & Wildlife Service (USFWS)
import-export permit is required for any international transaction, and,
in the case of import, an export permit is usually needed from the coun-
try of origin. In the U.S., specimen import is also regulated by the U.S.
Department of Agriculture’s Animal and Plant Health Inspection Service
(USDA-APHIS; www.aphis.usda.gov). Thus, in the U.S., permits to trans-
port and possess wild bird specimens should be obtained from both
USFWS and USDA. Learning of the specific requirements prior to any
field collecting or international transaction can save much time and trou-
ble. Realistically, plan for permitting processes to take 6–12 mo before
field work can be conducted. And be aware that many different permits
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might be required, depending on where your activities are being per-
formed. For a recent collecting trip to Canada, for example, I needed a
total of seven permits.
Specimens of species listed under national laws concerning threatened
or endangered species, and species listed in the Convention on Interna-
tional Trade in Endangered Species (CITES) appendices usually require
special permit procedures. Many international specimen shipments re-
quire CITES permits (sometimes even if the taxa involved are not CITES-
listed!).
In some countries, researchers at institutions must obtain permission
from Institutional Animal Care and Use Committees (IACUC, or an equiv-
alent) to study birds using the methods outlined by the researcher in a
proposal to the committee. The duty of these committees is to assure the
ethical and humane treatment of animals in science, and workers adher-
ing to professional guidelines (Gaunt and Oring 1997) should have little
difficulty in obtaining IACUC approval. Some funding agencies (e.g., in
the U.S., the National Institutes of Health and the National Science Foun-
dation) will not make grant awards available until IACUC approval is
obtained. Although it is not within their purview, some IACUC commit-
tees do not approve of the scientific collecting of birds, and it may be
difficult to convince them of the need to actively collect specimens. In
these cases and in permit applications it is often useful to point out the
scientific benefits and justifications for collecting birds, and that the num-
bers taken are small in relation to natural and other human-related mor-
tality factors. Publications where these and other useful data and argu-
ments have been assembled include Banks (1979), Winker et al. (1991),
Goodman and Lanyon (1994), Remsen (1995), Winker et al. (1996), and
Winker (1996, 1997). Educating permit-granting agencies, committees,
and personnel, while at times rewarding, has become a time consuming
and often frustrating part of the business.
Permitting procedures change frequently, and I know of no better
method to proceed than writing or telephoning the appropriate agencies
to request a set of current guidelines. Association of Systematics Collec-
tions (1993), searches on the internet, and inquiries to biologists familiar
with local permitting conditions can speed the process.
IN THE FIELD
Methods of obtaining birds are too varied to be treated in great detail,
but a brief summary is useful for the novice. Most birds added to research
collections today probably come from the salvage of birds found dead,
but collecting birds using shotguns or traps remains an important means
of acquiring specimens not obtainable through salvage. Bub (1991) gave
methods of trapping; the mist net is probably the most commonly used
trap in general use today (see Appendix for addresses of suppliers).
Collecting birds using firearms has become an arcane aspect of field
ornithology, despite its historic prominence. For larger birds, the standard
firearms and ammunition used in game bird sport hunting are effective.
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For smaller birds, however, specialized equipment is necessary. Museum
collecting has become centered on double-barrelled shotguns, usually us-
ing either 12, 16, or 20 gauge weapons with auxiliary barrels. Auxiliary
barrels are machined metal inserts that enable one to fire a smaller cal-
iber shell from a weapon chambered for something larger. For example,
auxiliary barrels can enable one to fire .410 or .22 caliber shotshells from
a 12, 16, or 20 gauge shotgun. ‘‘Aux’’ barrels are easiest to use when less
than about seven inches long. They drop into the shotgun chamber like
a long shell, and they are removed for reloading or when one wishes to
use a full-sized shotgun shell. Carrying a 12-gauge, double-barrelled shot-
gun with .410 and .22 auxiliary barrels enables one to collect anything
from geese or cranes to hummingbirds with the same firearm. The weap-
on is normally carried with a different caliber in each barrel to enable
the proper load to be used when required, depending on species and
distance (e.g., .410 and .22, or 12 gauge and .410). Firearms safety and
laws regarding firearms are as applicable to bird collection as to any other
use of firearms. Always assume the gun is loaded and be sure it is pointed
in a safe direction.
When collecting, one wishes to dispatch a bird quickly and cleanly while
at the same time minimizing damage to the specimen. Here, shot size
and distance are the two most important considerations. Proper distance
can be gauged through experience and practice, and varies with load and
firearm. Most museum collecting of smaller birds using the smaller shot-
shells (.410 and .22) is done with No. 12 lead shot. No. 12 shot is only
available in lead, and usually must be specially ordered. Loads containing
shot of this size are commercially available in the U.S. in .22 caliber shot-
shells, but not in .410. For the latter, most active North American collec-
tors are probably now reloading their own custom ammunition. Larger
shot sizes (No. 9 is available commercially in .410, for example) are ap-
propriate for larger birds (e.g., jays), but are not as effective as 12 shot
for smaller birds because the pattern (spatial distribution of the shot) is
not as dense.
Dispatching birds is probably the most difficult task faced by the col-
lector—not physically, but psychologically. Generally, those of us studying
birds do so because we have developed a strong liking for them. Conse-
quently, actually killing them is decidedly unpleasant. However, having
the specimen, which you know is valuable for the science of ornithology,
is often a great pleasure, regardless of the specimen’s source. The col-
lecting of birds for science is unquestionably justified, and this justifica-
tion keeps the collector active. ‘‘But let all your justifiable destruction of
birds be tempered with mercy; your humanity will be continually shocked
with the havoc you work, and should never permit you to take life wan-
tonly. Never shoot a bird you do not fully intend to preserve, or to utilize
in some proper way.’’ Coues (1874:30).
Two methods are generally used to dispatch birds in the hand, whether
wounded or in traps: thoracic compression and cervical dislocation. The
former is preferred in small birds because it is simple, quick, and causes
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no damage to the specimen. As I practice and teach it, thoracic com-
pression involves slipping thumb and forefinger in under the feathers on
each side of the bird (positioned between the spine and sternum), softly
feeling the ribs to be sure the position is correct, placing the forefinger
of the other hand against the front of the sternum, and instantlybringing
all three fingers together with a great deal of pressure focused on the
heart and lungs. This instantly stops the heart and lungs, and must cause
blood pressure to skyrocket. For small birds, it seems thatunconsciousness
occurs instantly; death follows very quickly. Pressure is held until it is
certain that the heart won’t restart. Twitches of the muscles can often be
felt quite plainly; pressure should be maintained until they stop. This
method is fast and, in my opinion, essentially painless. I get no enjoyment
from this difficult task, and use the method because it is so quick and
seemingly humane. I stress to students that anyone who might enjoy this
should find another field of study.
Cervical dislocation involves quickly stretching the neck (not twisting)
until the spinal cord is broken. Because thoracic compression is not suf-
ficiently rapid in larger birds, cervical dislocation is preferred. The latter
method is not usually acceptable in small birds because it can easily dam-
age the specimen and makes skinning more difficult. Methods used by
veterinarians to dispatch birds usually involve drugs or gasses that are
often illegal, inconvenient to carry in the field, damaging to the speci-
men’s value, or of some risk to personnel.
Preparation is made easier by preventing body fluids from getting onto
the plumage. This process begins as soon as one picks up a bird. Living
birds should be held with the cloaca pointed downward. Freshly shot
birds, or any bird that is wet or leaking body fluids, can be placed in a
bag of cob dust, sawdust, or corn meal to absorb moisture and keep the
plumage clean. Also, a wad of absorbent cotton or tissue paper should be
put down the throat—this is the most common source of fluids leaking
onto the plumage. Perforations of the skin (e.g., from shot, cats, raptor
talons, impact wounds, etc.) can also be plugged using cotton or absor-
bent paper to prevent leakage. Washing and drying feathers adds a lot of
time to the preparation process, so preventing them from becoming dirty
in the first place is worth the time invested here.
Specimens should be kept as cool as possible until they are prepared
or temporarily preserved through freezing or fluid preservation. When
external body fluids have become stabilized or dried in a bag of dust, I
generally place the specimen into an open plastic bag kept or carried in
shade away from my body until it can be labelled and frozen. Plastic bags
may be used rather than the paper cones of yesteryear: while continuing
to prevent plumage disarray they do not need to be made up in the field,
and they do not contribute to subsequent desiccation of the frozen spec-
imen. Every opportunity should be taken to allow birds to cool and to
keep them cool until short- or long-term preservation is possible. A thin
canvas bag, small pack in cool weather, or even a fishing creel or outer
coat pocket can be used to carry dead birds in the field. Just remember
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to allow the body heat to escape and to prevent feather disarray. Dead
birds should always be handled in a manner that preserves the integrity
of the feathers. Bending feathers and causing them to be ruffled back-
wards should always be avoided. One need not be particularly delicate;
just be sure that contact with the bird goes with, rather than against, the
grain of the feathers. Never grab or hold a bird by its tail or wings.
Freezing birds.
—The best way to preserve birds when there is not time
to prepare them immediately is to freeze them. Tissue decomposition
begins immediately upon death, and substantial degradation of important
elements can occur within the first hour. For example, the gonads of
immature passerines can become unrecognizable within1hofdeath
(even 30 min in warm conditions), and proteins also deteriorate, making
it necessary to freeze birds (or a tissue sample) relatively quickly after
death if they are to be useful for studies of allozyme genetic markers.
Freezing birds within 0.5–2 h of death is best (cf. Johnson et al. 1984).
Degradation of DNA also begins following death. Freezing or otherwise
preserving tissue samples for DNA studies is highly desirable as soon as
possible after death (see below). Specimens should be sexed when tissues
are taken.
Often, freezing the whole bird is desirable. Preparation is usuallyeasier
in a lab than it is in the field, and, when it is possible to freezespecimens
for later preparation, the return on the investment of field time can be
maximized. If a freezer is not available, dry ice (solid CO
2
) is preferred
for temporarily freezing birds in the field (and also for shipping them).
If kept well insulated, a 23-kg block of dry ice can last as long as two
weeks in rather warm conditions, but this is exceptional and requires little
use of the ice: few exposures to ambient temperatures and only a few
small specimens frozen. Dry ice in blocks is far superior to pellets, pucks,
or small blocks of condensate compressed straight from a CO
2
tank.
Whenever freezing a specimen collected or found dead, each bird
should have a label containing data on date and locality (at least), and
habitat and other notes as well, put into a plastic bag with it or tied to its
leg prior to freezing. When using dry ice, place specimens directly on the
ice in a cooler.
A standard chest-type cooler can be modified for improved use with
dry ice by making a nested series of ‘‘dense styrofoam’’ boxes, with lids,
to fit inside (J. Klicka, pers. comm.). Standard styrofoam is subject to
severe deterioration under the temperatures of dry ice and liquid nitro-
gen; the blue kind of dense styrofoam is the best to use when exposures
to dry ice or liquid nitrogen are expected. As the ice sublimates and
shrinks, the cooler’s contents are transferred to the next-smallest box,
which is inserted into the cooler. This apparatus works quite well, but
caution must be used to keep some ice on top of the specimens; I have
had temperature differentials of over 45 C develop inside such a cooler
under hot conditions.
A less common and more expensive method of freezing specimens in
the field involves the use of liquid nitrogen (LN
2
) in dewars, or flasks.
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LN
2
is commonly used in the field to freeze tissue samples, and works
very well for this purpose (Johnson et al. 1984). It works less well for the
preservation of whole specimens, however. With care, it can be used suc-
cessfully with small birds, but some specimen damage is to be expected.
Upon immersion, liquid nitrogen seeps into the external air pockets of
a specimen. When the specimen is removed from the liquid, the nitrogen
expands very rapidly into a gas, causing explosive destruction of the walls
of these air pockets. The rachides (shafts) of flight feathers (rectricesand
remiges) are often riven, or shivered, as a consequence. Structural dam-
age and even partial loss can occur in these feathers when specimens are
frozen whole in liquid nitrogen. Less frequently, bills or other peripheral
parts are damaged. This damage can be minimized by preventing rapid
temperature change in the frozen specimens. You can actually hear the
damage occurring as small pops and clicks upon rapid removal from LN
2
.
Holding specimens in the vapor phase of LN
2
for several hours or over-
night diminishes damage considerably.
Using this method, specimens are not placed directly into LN
2
; they
are first wrapped. Plastic bags do not survive immersion in LN
2
. Wrapping
birds first in multiple layers of paper tissue (bathroom or facial tissues
work well), then enveloping them in two layers of aluminum foil (label
inside) works reasonably well (G. Graves, pers. comm.). These envelopes
are then dropped into a wide-mouthed dewar for transport or placed
inside the canes of small-mouthed dewars. This method is only usable
with relatively small birds (ca.
60 g). When removed from LN
2
, birds
should be put directly into a
80 C freezer or onto dry ice pending
preparation to maintain tissue quality. A final source of damage to LN
2
-
frozen specimens is that they are so cold that they are very brittle. An
aluminum foil sarcophagus or envelope prevents most damage, but care
must be taken to treat these specimens very gently. Do not drop them
onto a hard surface.
For brief periods of freezing, ‘‘blue ice’’ (reusable plastic vessels filled
with a refreezable fluid or gel) of at least
20 to
80 C can be used to
bring birds from the field to the freezer. Although messier, bags of water
frozen to
40 to
80 C also work if blue ice is not available.
Storing frozen specimens.
—Freezing provides only temporary preserva-
tion; deterioration also occurs in frozen birds. Freeze-drying, the evapo-
ration or sublimation of water out of the tissues, is the most common
problem with frozen birds: water escapes from the carcass into the air
surrounding it and recondenses outside the tissues. To minimize this
problem, it is important to (1) minimize the time a bird is in the freezer;
(2) minimize the amount of air that the specimen is exposed to (use
plastic bags with as much air as possible removed from them); (3) main-
tain a temperature of
20 C or colder, if possible; and (4) minimize
temperature fluctuations. Frost-free freezers are to be avoided at all costs.
They undergo dramatic fluctuations in temperature designed to thaw,
vaporize, and remove water condensation in the freezer compartment.
This action mobilizes water in bird carcasses also, resulting in its recon-
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densation in the bag, but outside of the tissues. Buffering specimens from
temperature changes and exposure to air is important. Small specimens
that have been triple-wrapped in plastic bags with all of the air possible
removed from them, then placed inside styrofoam coolers in a constant
temperature freezer, can be prepared quite easily up to three years fol-
lowing freezing. This is an exceptional length of time, however, and
freeze-drying is usually detectable in the outer joints of small birds within
the first year of freezing. Preparation of frozen specimens that have suf-
fered freeze-drying can be assisted by soaking the affected parts (using
soaked cotton) or even the whole bird in water in a refrigerator for aday
or two.
Taking, preserving, and archiving tissue samples.
—Specimens frozen
whole on dry ice or LN
2
very quickly following death provide tissue sam-
ples of high quality for genetic studies. It is important to take tissue sam-
ples from these specimens as soon as possible to maintain this usefulness.
Whole frozen specimens held at
80 C prior to preparation can be
brought slowly up to room temperature by being held in an ordinary
(
20 C) freezer for hours or days first, or brought directly to room tem-
perature under close observation. Tissues should be taken and refrozen
as soon as the specimen has thawed sufficiently to be worked. Usually,
tissues can be taken while the organs are still frozen. With experience in
preparation, in small birds the skin can be removed from the carcass and
tissues taken before the internal organs have thawed. Two tubes oftissues
should be preserved whenever possible. These should be stored separate-
ly, if possible, for both the short and long term to prevent loss of critical
material through freezer failure or other catastrophe. Also, tissue tubes
should be filled to within 2–3 mm of their tops whenever possible. This
maximizes the amount of tissue available for future studies, minimizes
wasted freezer space, and prevents tissue desiccation.
Taking tissue samples from each specimen should be a priority for any-
one collecting regularly, and can be done by anyone who has the oppor-
tunity to work with dead birds. Winker et al. (1996) emphasized the cru-
cial nature of voucher specimens when taking tissue samples. I will not
discuss sampling protocols that do not preserve voucher material (e.g.,
bleeding and releasing live birds). Such cases should be exceptional. Tra-
ditional allozyme studies use several different tissues because of the dif-
ferent proteins available in each. In general, heart, liver, muscle, and
kidney are saved for these studies (Johnson et al. 1984). Although allo-
zyme studies have declined in popularity with the advent of improved
DNA technologies, when one is freezing fresh tissue samples it is a good
idea to preserve all of these tissues in each tissue tube. Doing so in the
order above insures that the types are separable later in the molecular
laboratory. For modern DNA studies, frozen tissues are not absolutely
required, and freezing has become less popular because of the extra field
logistics required (LN
2
or dry ice). This trend is short-sighted, however;
frozen tissue samples are still preferable. When freezing tissues is not
possible, one can place minced tissues in a vial of buffer (Seutin et al.
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1991) or place small pieces into 95% ethanol for dehydration. New ar-
chival methods of taking and storing genetic samples will be or are being
developed (e.g., using blood-soaked paper that has been chemicallytreat-
ed), but thus far ultracold freezing represents the closest thing to a mu-
seum-quality tissue preservation method. Given DNA chemistry, aqueous
solutions do not represent long term, archival-quality storage conditions
(see Cann et al. 1993, Poinar et al. 1996). Thus, aqueous field storage
methods should not be trusted over the long term (years), and should
be stored in cryosystems upon return to the museum or laboratory.
Labels and field catalogues.
—Proper data recording and management is
perhaps the most important aspect of specimen preservation, and doing
it right begins in the field. Labelling is also where most errors occur. An
improperly labelled specimen is of limited scientific value, and a specimen
with no data may be useless. ‘‘. . . never put away a bird unlabelled, not
even for an hour; you may forget it, or die.’’ (Coues 1874:70).
Historically, proper data management could be achieved using thelabel
alone. This becomes much more difficult when several pieces of the same
animal are preserved, however, because these pieces need to be easily
linked when being archived and, eventually, used. The method outlined
here is centered upon a field catalogue, and the link among these sepa-
rate pieces is the unique field catalogue number. Consider that each pre-
served part is destined for a different physical collection (e.g., skin, skel-
eton, tissue), and that pieces often come to reside in different institutions.
The field catalogue becomes the central data source for all of the pre-
served parts, because full labels will not be written for each. The field
catalogue is never meant to be housed with a specimen, however, and
time, together with specimen loans and transfers, assures that the field
catalogue is often not available. Attached labels are therefore the most
important and most used documents for data retrieval. My method uses
the skin label as ‘‘the final label;’’ it will be as complete as the field
catalogue entry and will be the only label with all of the data.
Personal field catalogues are a tradition in ornithology, but, as a con-
sequence, many have been lost. Thus, beginning a personal catalogue
should only be done when there is not another immediate cataloguing
option. I regularly use project-specific and laboratory ‘‘field’’ catalogues
to bring together under one system the efforts of many individuals who
would otherwise generate an uncontrollable proliferation of idiosyncratic
note taking. I have found a form style of field catalogue (Fig. 1) superior
to the blank page because it prompts the wandering mind and results in
more consistent quality and completeness of data. It can easily be gen-
erated on 100% cotton paper using a computer and laser printer. Using
a loose-leaf catalogue format on standard-sized paper enables several in-
dividuals to use the catalogue simultaneously, makes photocopying easy
(to distribute to project participants, for example), and allows completed
originals to be immediately archived in the museum to prevent loss. The
loose, unused sheets are kept together in a folding aluminum clipboard/
container, and usually just one sheet at a time is removed for use. Com-
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F
IGURE
1. A form-style field catalogue page improves the consistency and quality of speci-
men data. A loose-leaf catalogue on 100% cotton paper is easily made up and printed
using a word processor and laser printer. It is also easy to make copies when the sheets
are completed, and one can take only blank sheets into the field, offering less risk of
losing completed pages and their data. Key to abbreviations used in catalogue and on
labels (Fig. 2): HAB: habitat; WCH: wing chord (Baldwin et al. 1930:76); TL: tail length
(Baldwin et al. 1930:92); TS: tarsometatarsus (‘‘tarsus’’) length (Baldwin et al. 1930:
107); BL: bill length from anterior edge of nostrils to tip (Baldwin et al. 1930:16; see
Parkes 1988 for discussion on bill measurements); BLH: bill height from anterior edge
of nostrils (Baldwin et al. 1930:20); BLW: bill width from anterior edge of nostrils (Allen
1889:188); SKL: skull length from rearmost part of skull to bill tip; TE/OV:testes/ovary;
SK: stage of ossification (most useful among passerines), stated in approximate per-
centages.
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pleted sheets and those in the process of being completed during speci-
men preparation are stored in filing cabinets in the museum and can be
bound if desired.
Assignment of unique numbers (i.e., using a field catalogue) becomes
necessary as soon as subsamples are taken or preparation begins. The
unique number assigned to a specimen is a combination of the catalogue
abbreviation (full initials if personal, a project or laboratory abbreviation
if not), and an incremental number (Fig. 1). The field catalogue should
contain all of the data associated with the specimen, and recording the
appropriate data for each specimen should begin with collection and be
steadily completed during the preparation process. Until preparation is
complete, each entry in the field catalogue is a working document. The
collector and preparator (often the same person) work in tandem (per-
haps months or even years apart) to complete the entry.
The final label should be written immediately after preparation and
the field catalogue entry are completed. The data associated with a spec-
imen are what make it valuable to science. Completing the field catalogue
entry and the skin label during and immediately following preparation
(and before doing anything else) not only provides an immediate review,
assuring that all of the appropriate data are recorded, but generates a
second original record of these data, a prudent safety precaution. When
finished, then, two complete original documents exist containing the full
data for the specimen and its preserved parts.
To streamline data recording and information retrieval, I use two mu-
seum labels during the collection and preparation process. I consider it
essential that full field data are attached to the specimen right after col-
lection or salvage, and use the first label in the field, writing in (at least)
date, locality, and collector. This label is later used for the partial skeleton
following preparation; the only additions to it are sex, a note of the dis-
position of the specimen (how it was prepared), and a second writing of
the field catalogue number in case the label becomes damaged in the
skeleton preparation process (it may be partially eaten by dermestids if it
is exposed to body fluids). The second label (a new one) is made up
following preparation as the final label for the skin, and contains all of
the data associated with the specimen. This final label for the skin is
written as the last step in preparation and is tied to or pinned unambig-
uously beside the drying specimen. Always record a specimen’s complete
data before going on to another. Never try to remember these details to
write them later.
At a minimum, a label should contain the date and locality of collection
(or salvage) and the name of the collector. In montane regions elevation
should be included. A temporary or field label with this information is
all that is needed if a specimen is being frozen for later preparation, but
additional information is often included at this time (e.g., soft part colors,
behavior, habitat description). Such a label is far too incomplete for a
final specimen, however. A complete specimen label should contain data
on date, locality, collector, preparator, field number, sex (including con-
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F
IGURE
2. A completed specimen label. Actual dimensions are 19 86 mm. Note that it is
flipped from front to back on the short axis. Abbreviations given in Fig. 1.
dition of gonads and any external reproductive characters such as cloacal
protuberance or incubation patch), skull ossification (in species like pas-
serines where it is a useful aging character; see below under Aging by
Skull Ossification), mass, fat, molt, and disposition (what parts of the
animal were preserved). Additional data are very desirable, but may be
dispensed with if necessary: habitat where the bird was obtained, soft part
colors (e.g., bill horn, iris red, feet lead blue, etc.), measurements, age,
stomach contents, behavior, time of day, parasites, any noteworthy re-
marks, and species identification. The identification of a specimen is per-
haps the least important, except for very young birds and complete skel-
eton preparations when no skin is preserved, in which case it is essential.
Errors are regularly made in field and laboratory identifications; those
preparing complete skeletons should preserve skin material as well for
vouchers. Skins are identifiable to species and usually subspecies in the
museum collection, and serve an important role as vouchers, especially
for molecular studies (Winker et al. 1996).
There are standard ways to record most of these data so they are in-
stantly recognizable and fully and unambiguously recoverable from every
label. Figure 2 shows a widespread method. The positions on the label
where the data are recorded are important also; following these standards
assures that your labels will match those in most museum collections.
Date is given as day (number), month (written out in letters), and year
(four-digit number).
Never
represent the month as a number! This error
has caused many specimens to be removed from studies because of un-
certain dates. Locality is written in full, with the major geographic division
first, followed by greater detail and ending with the most specific infor-
mation (preferably latitude and longitude). Collector and preparator
should be clearly indicated (front of the label for the former, back for
the latter) so the quality and veracity of the data can be judged and credit
given to the people responsible for the specimen’s collection and pres-
ervation. Field number connects all of the parts of the animal to thesame
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individual and points to the field catalogue entry where original data are
recorded. The sex of the bird can only be truly learned by dissection,
making the preparator fully responsible for accurately determining and
describing the sex organ(s). This information is crucial, and is often ac-
companied by other sexually-related data (e.g., incubation patch or clo-
acal protuberance present). Body mass must be taken as shortly after
collection as possible and prior to removing anything from the bird. A
bird’s mass is related to size, fat load, and reproductive condition, and is
a useful datum for many types of studies. Fat load gives important clues
to a bird’s physiological or migratory condition, and molt provides ad-
ditional important data about the condition of the individual and the life
cycle of the population or species. If no evidence of fat or molt is found,
write ‘‘none.’’ An absence of information means that you didn’t look.
Degree of skull ossification is useful for aging many passerines. Remarks
provide additional information about the specimen; it may be the parent
or offspring of another individual (give catalogue number), heavy parasite
load, disease, or aberrancy; or it may have been washed in preparation.
Finally, disposition informs the future user and museum cataloguer what
else exists from this individual, greatly aiding recovery and use of all avail-
able preserved parts.
These constitute the essential data that should accompany each speci-
men, and it can be seen from Figure 2 that both the front and back of
the label record this information in traditional positions. The rest of the
data that most specimens should contain (e.g., habitat, measurements,
remarks, etc.) are written on the back in the remaining space. I find that
a free-form text method enables me to fit everything on a single label,
even if six or seven lines are required. Using a stereotypical order dupli-
cated in the catalogue (Fig. 1) speeds writing and information recovery.
Our data entry software is also set up in this order to speed the comput-
erization and cataloguing of specimens. A generic, preprinted label for-
mat (Fig. 2, without the writing) is preferable because it allows maximum
flexibility and can be ordered in volume. Custom stamps are often used
to fill in parts of the label when working for a long period in a particular
locality; preprinted labels can also serve this purpose. Foster and Cannell
(1990) discussed most categories of label data at greater length. Using a
second label may be necessary occasionally, especially when descriptions
of soft part colors are long, or when sketches of gonads, bills, or skulls
are used. In these cases the back of the second label serves as another
page.
The final label attached to the skin is an original document written by
the collector/preparator to the future users of the specimen. It should
be complete and it should retain its originality. Never replace an original
label. If more space is needed for writing, add a second label. It isamazing
how much can be learned of past preparators and collectors—their
strengths and weaknesses—through reading their labels. Further,the sam-
ples of handwriting borne by each label are often essential for interpret-
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ing comments subsequently written on labels (in pencil) in museum col-
lections.
The labelling practice of affixing only a number to specimens, to have
labels completed later (e.g., upon return from the field, or after prepa-
ration), is to be assiduously avoided. Having spent inordinate amounts of
time trying to track down the field catalogues associated with such spec-
imens, I can state assuredly that the small amount of time saved by the
collector or preparator regularly dooms the product. An appropriately
labelled specimen is immediately useful, regardless of its final destination
and uncertain preparation, deposition, and cataloguing future.
A third label is eventually made up for the finally prepared skeletal
specimen in the museum, but these labels are often computer generated
and added to the specimen months or even years after the skin has been
catalogued.
Labelling tissue vials presents a special problem in that, for frozen sam-
ples, labelling must be completed prior to freezing: one cannot write on
a frozen vial. Some curators have gone so far as to give tissue samples
final tissue catalogue numbers during preparation (in field or laboratory)
by issuing blocks of blank tissue catalogue pages or numbers to collectors
and preparators. This requires the preparator to fill out two catalogue
entries during preparation, just to issue the specimen a (usually) second
unique number. I find this to be needlessly cumbersome and rarely com-
prehensive—not all specimens entering a collection will be generated
using the in-house numbering system. Cryovial manufacturers make in-
serts for the vial lids that can be written upon and inserted when samples
are finally arranged for cryostorage and catalogued. A specimen does not
need multiple unique numbers prior to being catalogued into a collec-
tion, and the traditional field catalogue number is sufficiently unique to
serve the purpose. This number is also borne by all of the other parts of
the animal. Although I haven’t seen a duplication in the traditional ini-
tials-and-number field catalogue system (Fig. 1), the date, taxon, and lo-
cality information accompanying the number make later confusion seem
impossible should duplication ever occur.
However, given the labelling constraints on tissue vials (even tissues in
buffer or alcohol should be frozen for long term archiving), the method
outlined in Figure 3 is preferred, since it provides all of the information
necessary to assure rapid and accurate sample recovery from the freezer
whether the sample has been catalogued into a collection or not. Writing
the field number twice is insurance against the loss of information
through abrasion, which is not uncommon, especially in LN
2
dewars.
Finally, no matter how rushed you seem to be, make a conscious effort
when writing labels to do a slow and careful job. You are notwriting notes
to yourself. It is imperative that your penmanship be exemplary. For ex-
ample, never write a label in script handwriting; always print carefully.
Localities, remarks, strange abbreviations, and people’s names are often
uninterpretable on specimen labels because someone was in a hurry.
Temporary fluid preservation.
—Freezing specimens is not always possi-
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F
IGURE
3. Label as written on outside of cryovial of tissue samples, stomach contents vial
label, and traditional skin label knot.
ble. Yet, when one has only a short time in the field, preparing every bird
as it is obtained would seriously limit the number of specimens that could
be collected. This is particularly true for brief visits to remote locations.
Weber et al. (1984) presented a method of fluid preservation that enables
one to maximize field time in such situations and bring specimens back
to the laboratory for preparation. Specimens preserved this way are more
difficult and time consuming to prepare, and on average the skins are
not as good (aesthetically) as those freshly prepared. However, the meth-
od enables one to collect more individuals when in the field, increasing
the return on the investment. I have had some success with this method
of temporary preservation. It involves a dilute solution of formaldehyde,
phenoxyethanol, and salts. Concentrated chemicals are carried into the
field and mixed with water in a pickling container. Gloves are used when-
ever contact with the preservative is possible, and care should be taken
to minimize inhaling the low-level formaldehyde fumes.
Tissues are removed from the specimen prior to pickling (formalde-
hyde causes serious damage to DNA; Vachot and Monnerot 1996) and
are placed in buffer or LN
2
. The bird is sexed, eviscerated, and labelled,
its stomach preserved in alcohol, the plumage and body cavity cleaned of
blood, then it is wrapped and pinned in a single layer of loose gauze
prior to immersion in the pickling solution. Do not use pins with plastic
heads; the heads dissolve and may stain the plumage. When the specimen
is pickled its position should be close to that of the final preparation,
because the feathers become essentially pickled into place and are far less
workable than in a fresh or frozen specimen. Blood should be assiduously
removed prior to wrapping in gauze; it becomes fixed in the solution and
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stains the plumage with a dark, blackish-brown color. Washing bloody
plumage in clean water and swabbing out the body cavity with cotton
prior to wrapping are recommended.
Specimens should be stirred once or twice daily in the solution to as-
sure complete pickling. Upon leaving the field, the fluid can be drained
and the moist specimens wrapped in cloth and placed within many plastic
bags and/or a leak-proof container for transportation back to the labo-
ratory. There, a fresh solution can be made up to complete any pickling
not finished in the field, as well as for storage of the specimens until
preparation. I have detected no change in such specimens stored this way
for up to 4 yr following collection. Prior to preparation, the pickled spec-
imens are rinsed under clean water for 1–3 d to leach out most of the
chemicals. Gloves are used in skinning, and each skin must be completely
washed and dried.
Soft part colors.
—The colors of the bill, legs and feet, irides, and things
like wattles, external air sacs, etc. generally change rapidly upon death.
The colors of these parts often vary among age and sex classes as well as
seasonally and geographically. Describing these colors while the bird is
still fresh using standard color references such as Ridgway (1912), Smithe
(1974–1981), and Munsell (1990) enhances the value of the specimen
both to science and to artists who might use the specimen for subsequent
illustrations.
Measurements.
—Many specimen measurements change upon prepara-
tion and drying (e.g., Winker 1993). A bird should be weighed and mea-
sured before anything is modified or removed (e.g., tissues or stomach).
Occasionally, one encounters birds whose mass is clearly affected by sub-
stantial food items; such individuals can be re-weighed with the food items
removed (note this in Remarks). A researcher may take numerous mea-
surements from a prepared specimen, and it is not up to the preparator
to anticipate these. However, there are several commonly taken measure-
ments that change or are unobtainable in the completed specimen. I
generally take seven measurements prior to preparation (Fig. 1). Taking
these properly requires practice, but they can be performed rapidly and
accurately, with precision to 0.1 mm when using a vernier caliper is pos-
sible. Further discussion can be found in Winker (1999). Units of measure
are always grams for mass and millimetres for morphological characters,
and I omit writing ‘‘mm’’ for morphological measurements.
TOOLS OF THE TRADE
Although a perfectly good specimen can be prepared using nothing
more than a good pocket knife, some cotton balls, and makeshift mate-
rials, proper tools speed the job considerably. A sharp scalpel,two or three
sizes of forceps, small and large scissors, vernier calipers, a millimetretape
measure (for larger birds), pens with black, waterproof ink, cotton paper
and labels, heavy and light thread, needles, and pins constitute a good
preparation kit. In addition, cryogenic vials, vials for stomach contents,
pinning boards (e.g., styrofoam), gauze, cotton, sticks, cob dust (or saw-
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dust or cornmeal), excelsior (for large birds), heavy thread and string,
and alcohol constitute supplies consumed in the preparation process.
Some sources for supplies are given in the Appendix. Finally, a personal,
project, or laboratory field catalogue should be available for recording
full data and assigning a unique catalogue number to the specimen.
No single pen functions well for all of the writing that preparation
requires.
Never use pencil.
It doesn’t xerox well (important for field cata-
logues), it often fades or smudges, and it can completely disappear with
grease and long term label color changes. Only black, waterproof inks
should be used, and your pens should be tested for known short-term
problems. For example, alcohol may dissolve a waterproof ink, which is
important to know before putting labels in stomach content vials. Tech-
nical pens, used by draftsmen and artists, are ideal for labels and field
catalogues, but are expensive and inferior for cryovial labeling because
the ink flakes off at ultracold temperatures. Fine point permanent mark-
ers (e.g., ‘‘Sharpie’’) are good for writing on cryovials, but are usually
inferior for labels and catalogues, and they usually dissolve with alcohol.
Inexpensive ballpoint pens with permanent ink work well for field cata-
logues, but can rarely be found with point sizes small enough for the
delicate writing of legible labels. And some ballpoint inks dissolve on fat-
soaked labels or when exposed to naphthalene (K. C. Parkes, pers.
comm.). I use three pens: technical for labels, permanent ballpoint for
catalogues, and permanent marker for cryovials.
PREPARATION
1. Put a wad of absorbent cotton or facial tissue down the throat to
prevent fluid leakage. If one is already there, replace it.
2. Record soft part colors if bird just died (Ridgway 1912, Smithe 1974–
1981, Munsell 1990).
3. Weigh bird and take any desired, irreproducible measurements if
not already done. Record in catalogue. Note if bird has been in freezer
for a long time prior to weighing; desiccation causes loss of mass.
4. Label tissue cryovials with Sharpie marker. Neveruse a technical pen.
Include catalogue number, species, locality, date, and catalogue number
again (Fig. 3). Abbreviate species and locality if necessary; make sure the
catalogue number is written legibly and indelibly
twice
; they can become
lost through abrasion or illegibility. Usually, take two tissue samples.Label
vials as ‘‘1 of 2’’ and ‘‘2 of 2.’’
5. Make incision in skin from furculum to cloaca. Begin separatingskin
from body (Fig. 4). After making the initial incision, separate skin from
body using forceps to push body surface inward while gently pulling skin
outward or, better, pushing skin outward using forceps and/or a finger
(Fig. 4).
6. Make incision into body cavity; carefully remove stomach/gizzard
(big, hard, in the way) by tearing or cutting the esophagus and intestine
free (Fig. 4). Set it aside for later.
7. Be careful not to disturb or mess up the body cavity too much: you
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IGURE
4. Initial incisions through skin and abdomen and relative positions of majororgans
occupying the abdominal cavity, illustrated, anterior to posterior, as heart, liver, stom-
ach/gizzard, and intestines.
F
IGURE
5. Determining sex requires direct observation of gonads, here illustrated in a non-
breeding male (left) and female (right). Gonadal tissue almost always lies on top of the
posterior third of the bird’s left adrenal. Adrenals occur in both sexes in pairs near the
anterior edge of the kidneys. Males always have two testes, but most females have only
one ovary, the left, as illustrated.
must be able to examine the sex organs, which lie along the dorsalsurface
of the cavity near/upon the anterior side of the kidneys (Fig. 5). Intes-
tines can usually be removed—carefully, so that fluids don’t get on feath-
ers and gonads aren’t disrupted.
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IGURE
6. Poke the knee into visibility and disarticulate. Skin leg to next joint (articulation
of tibiotarsus and tarsometatarsus). One leg should be removed for the skeleton by
cutting the skin (center, lower dotted line; cleaned leg at right). In some cases disartic-
ulation of a leg is required to obtain one unbroken tibiotarsus for the skeleton (center,
upper dotted line).
8. Take tissue samples and sex bird. First remove heart (Fig. 4). If from
a small bird, place whole heart in cryovial 1. If bird is larger (e.g.,
Cath-
arus, Passer
or larger), cut heart in half and put half (or some) in each
vial. Next, put some liver into each vial, then sex the bird (see Determin-
ing Sex below; Fig. 5). Record gonad type and size in the catalogue.After
sexing, kidneys can be removed (this disturbs the gonads); put somekid-
ney tissue into each tissue vial. At this point your tissue vials should each
be approximately
½
full, except for very small birds (e.g., many Parulidae,
Sylviidae, Trochilidae). Fill each to within ca. 2 mm of top with breast
muscle. On small birds this involves fileting out the entire breast with a
scalpel. Replace caps onto vials very tightly and freeze tissues immediately.
Caps that are not replaced very tightly often come off in LN
2
and the
samples are lost.
When you have attained the ability to skin out a bird in about 10 min
(separating skin from body), you can take tissues after the body is out.
Taking tissue samples and sexing a bird is much easier with the body
removed, but tissue freshness is extremely important. I try to get tissues
frozen within 30 min or less of thawing to maximize their usefulness in
molecular studies. With the carcass skinned, more tissue can be taken in
small birds. When skinning small birds, I generally take all large muscles
(breast, wings, legs) and all internal organs except intestines and stomach.
Lungs and kidneys must be lifted up out of their bony recesses, and with
practice can usually be taken out in one or two pieces.
9. Skin. Begin with knees, and proceed with legs (Fig. 6), tail, and
shoulders, to head (making incision in rear of head/neck for large-head-
ed birds such as Picidae and Anatidae), then pinch and roll out ears, and
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IGURE
7. Body is skinned to shoulders, which are disarticulated. Skinning proceeds to
head, then past the ears, which are pulled free, and then past the eyes, from which the
skin is carefully cut behind the eyerings, to the base of the bill, where scissors are used
to cut the skull free in a plane across the anterior portion of the eye sockets (dotted
lines). Care must be used not to cut the tongue, which should come free with the body
for the skeleton.
cut skin away from eyes back far enough so that eyering remains intact.
Skin can be separated from body to base of bill, just in front of eyes (Fig.
7). To minimize holes, use forceps and fingers to separate skin from body.
Scalpels are used only where it says ‘‘cut’’ in details below. I use scissors
only to cut the vertebrae at the tail, to make the skull cuts, and, later, for
cutting thread.
Details: After the initial skin incision (step 5), work on one side at a
time. In general, throughout the skinning process you wish to avoid pull-
ing on the skin. Instead, work at the point where the skin still adheres to
the body, pushing the skin free using a thumbnail or forceps where pos-
sible. Poke the knee (by holding the lower leg) in toward the body while
holding the skin away from the body (sort of lifting the abdomen skin in
a bunch over the knee while the latter is held and pushed inward). Pop
the knee into visibility (Fig. 6). You don’t need to see much of it. Using
a wiggling motion (rather than slicing), ‘‘worry apart’’ the knee jointand
the surrounding muscle.
After the leg is detached below the knee from the body (attached only
to leg skin), push the skin free with your thumbnail down to the ‘‘ankle’’
joint (Fig. 6). This is the joint between the tibiotarsus and the tarsometa-
tarsus, and in most birds it is where feathering ends and skin or scales
begin. One leg will be removed and saved for the skeleton—the right if
unbroken. To remove one leg, when reaching the ‘‘ankle’’ joint cut aring
around the skin to separate it from the limb and remove the whole leg
for the skeleton (Fig. 6). If the foot is too large, cut the muscles away
from the tibiotarsus to get it out through the hole you’ve made at the
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base of the feathered leg skin, or make a lengthwise incision in the leg
skin to enlarge the hole at its end. Remove the muscles from the tibio-
tarsus and, if the bird is very small, add them to a tissue vial. Set this
removed leg aside for the skeleton (Fig. 6), and put the leg skin back
right side out. On the remaining leg, cut through the knee joint and skin
down to the ‘‘ankle’’ joint again. Now remove the muscles from this ti-
biotarsus, wrap a little cotton around the bone to replace the muscle mass,
and gently pull the leg back inside right and quickly brush the feathers
back into place. Leaving this bone in the skin with its cotton wrapping
adds a lot of strength to the leg and makes it comparable to historicskins.
If you have already saved a whole leg for the skeleton, cleaning the muscle
from the tibiotarsus remaining with the skin can be performed very rap-
idly by breaking this tibiotarsus near the proximal end, peeling thewhole
muscle mass away from the bone, cutting the tendons in one slice near
the distal end, and using a little dust and two fingers to rub away any
small amounts of meat remaining.
If a leg (tibiotarsus or tarsometatarsus) is broken, it stays with the skin.
If neither leg is broken, the left will remain with the skin. Decisions about
which bones stay with the skin and which go with the skeleton are made
before cutting the knees and after carefully feeling and visuallyexamining
each leg. In most cases the left leg can remain with the skin. Be sure that
one whole bone of both tarsometatarsus and tibiotarsus is available for
the skeleton. In some cases (commonly in shot birds) it is necessary to
disarticulate the tibiotarsus from the tarsometatarsus that will remain with
the skin to obtain a complete tibiotarsus for the skeleton (Fig. 6). Insuch
cases the remaining leg is only attached by a little bit of skin. It can be
given extra support through replacing the missing tibiotarsus with the
addition of a small stick and some cotton, but I generally skip this in
small birds to save time.
Now, working on each side separately to prevent the skin from being
stretched and ripped, separate skin from around the pelvic area. Work
down into this area; don’t try to open it up and lay the skin out flat or
you’ll tear it. When the skin is relatively loose down the sides, hold the
tail base between thumb and forefinger, putting the forefinger forward
enough to cover the muscular tail base and serve as a stopping point for
your scissors as you cut the vertebrae from the inside. From above, cut
the lower abdomen posterior to the upward-projecting public bones and
aim your scissors to cut the vertebrae behind these bones and in front of
the muscular base of the tail. Care must be taken not to cut the bases of
the tail feathers or they will fall out; however, the synsacrum or pelvic
bones should not be damaged either. When the vertebrae are cut, use a
scalpel to free the remaining tissue so the tail is separated fromthe body.
Free the lower body from the skin and evert skin with fingers to the
shoulders, disarticulating these with the scalpel (add dust). When skin-
ning down the back, push skin away from body using thumbnail andtake
care not to tear the skin by stretching it too much around the body. It
often helps to get it off of the high point of the breast as soon as possible.
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This can easily be done by grasping the whole skin-body interface all
around the bird with all five fingers simultaneously and pushing-pulling
this skin mass past the broadest body area. Disarticulate the shoulders to
separate the wings from the body (Fig. 7). This loosens things up consid-
erably. Be sure to add a lot of dust during this phase or you’ll get fluids
on the feathers.
Continue peeling to head, making an incision in rear of head/neckfor
large-headed birds such as Picidae and Anatidae (see Variations below).
With a little dust for friction, pinch and roll out ears, which prevents any
holes being made, then cut skin away from eyes back far enough so the
eyering remains intact. Skin can be separated to base of bill (just in front
of eyes). Using scissors, cut skull away from skin at the base of the bill.
Use three cuts, and don’t cut the tongue: a) Upper, frontal portion of
head, done straight down from the top of the head with the scissors in
the frontmost area of the eye sockets (don’t cut through the eyes them-
selves); b) Left mandibles: insert scissors (sharp tip inwards helps) straight
upwards on each lateral side of the upper and lower mandibles to cut
these bones; c) Right mandibles, same as left (Fig. 7). Gently twist and
pull to finish separation, keeping in mind that the tongue comes withthe
body. Also, don’t pull very hard, or you’ll rip the bill away from the skin.
You can make auxiliary cuts if necessary through the palate, but keep the
tongue (and its bones) intact. The body should now be separated from
the skin. If you haven’t taken tissues and sexed the bird, do so now (see
Determining Sex below; Figs. 4 and 5).
General points: Always be sprinkling corncob dust, sawdust, or corn-
meal onto any fresh surface to keep fluids and stickiness from getting
onto feathers. Feathers must be kept from contacting body fluids or you
will have to wash and dry the bird—a time consuming task to be avoided
whenever possible. Whole bones are removed for the skeleton until one
of each is preserved; broken bones can remain with the skin.
10. Skin, clean, and tie wings (Fig. 8). Setting the body aside, sexed
and with tissues taken, take up the skin and skin out the wings (Fig. 8).
Begin by getting the shoulder muscles up away from the skin so you can
peel down the humerus, then work the skin away down to the joint where
the radius and ulna begin. The secondaries are attached to the ulnae,
and they must be separated carefully to prevent tearing. When skinning
down the radius and ulna, place a thumbnail directly onto the ulna and
forcibly push downward (or outward, toward the distal end of the wing)
to separate each secondary from its bony attachment. In very large birds
a dull knife may be necessary to help make this separation. Skin down to
the ‘‘wrist’’ joint(where the radius and ulna form a joint with the ‘‘hand’
bones), and cut through this joint, having skinned down far enough that
you don’t cut the skin. Skin the other wing similarly. Leaving one wrist
intact is desirable so that the skin is comparable in wing measurement to
the great number of historic specimens. Make the choice based on skel-
eton quality: if the radius and ulna are broken on one side, leave the
distal portions of these bones with the skin. If the bones on both sides
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IGURE
8. Skin wing to distal ends of radius and ulna, separating bases of secondariesfrom
ulna with thumbnail. With wings cleaned and bones removed for skeleton, consider
interscapular distance on the carcass as one unit (center) and firmly tie wings together
so that the wrist stubs are 1 units apart.
are unbroken, I leave the left wrist intact, breaking the left radius and
ulna in half (a traditional method) and leaving the distal half with the
wing. In large birds, skin out the hand bones of one wing; I do the right.
This must be done carefully, because the primaries are attached to these
bones; the most distal bone is the most difficult. There is very little skin
down here, so work carefully. With practice, all of this becomes easy.
Thread a rather heavy needle with heavy thread (I prefer white button
and carpet thread, used also for labels) and make a good, big knot at the
end of the thread. With the distal ends of the wings still everted, push
the needle through the ‘‘hand’’ bones (but not the skin) of one wing
and pull the thread through to the knot so it’s tight. The other wing can
be tied with the thread to the broken ulna, or push the needle through
the other wrist, making sure first that the two wings are oriented correctly
(with their dorsal surfaces facing inward; Fig. 8). If the hand bones are
too sturdy to be run through with a needle, the wings can be tied together
by running the needle through skin on the underside of the wing beside
the distal joint of the radius and ulna. Be careful to use the underside,
where disarrayed feathers are hidden in the final skin, and not to put the
thread too close to the bases of primaries or secondaries, which often
become twisted as a consequence.
Now, arrange the everted wings so the dorsal surfaces face each other
and the primaries extend straight backward, almost touching each other.
Hold the two everted wing stubs in one hand with your first threefingers
at a distance from each other equal to the distance between the scapulars
on the carcass (we will use the carcass for a model several times). This is
‘‘one unit’’—the distance between the two wings on the living bird (Fig.
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8). Pull the thread so it is taut between the wing stubs when they areheld
at this distance (it is held on one side by the knot, on the other side by
your other hand). Now, with a pair of forceps, pull back into the inter-
stub space enough thread to equal
of the ‘‘one unit’’ of distance you
continue to maintain between the two everted wing stubs. Carefully tie
off the loose end of thread so that the two everted wing stubs are firmly
tied together at 1
times the ‘‘one unit’’ distance (Fig. 8). If tying to the
broken ulna on one wing, use about 1
interscapular ‘‘units.’’ This dis-
tance is important: if you’re off, the skin will be difficult to arrange prop-
erly later.
11. Clean tail, being careful of the thin skin. Be sure to cut or scrape
out the uropygial (oil) gland on the dorsal surface. Also, don’t cut off
too much muscle or other material here, especially in small birds, or tail
feathers will begin to fall out. But you do want it relatively clean and
grease-free. When most of the meat and glands have been scraped away,
use dust to rub remaining meat and oils off.
Between skinning and cleaning up the tail, a lot of work is done in this
area. In small birds this is a difficult part of the job—you are working in
a tight space where there is little skin and little room for error. You don’t
want to tear off the tail (easily done in small birds), or to have tail feathers
fall out (also easily caused). Conversely, you have to get out the uropygial
gland and most of the meat so the preserved skin isn’t ruined by rot or
oil leakage.
Tail feathers are arranged at their bases in tight rows whose basal tips
outline a rearward-projecting arrow. They are attached to the pygostyle,
a long, narrow bone that projects posteriorly down the middle of the two
rows of tail feathers. This bone and most of the muscle (all of the muscle
in large birds) can be removed (if done carefully) without compromising
the integrity of the tail feathers and the skin surrounding them. In small
birds, I generally evert the base of the tail, then hold it by folding the tail
back against the outside, dorsal surface of the skin, then grasping the tail
and the skin firmly together in one hand. This prevents stress on the little
skin keeping the tail attached, and the tail stub is exposed for careful
work with a scalpel. Meat edges are carefully teased off of the skin with
the scalpel edge and the skin worked back off of the tail stub on all sides
using a thumbnail. On the dorsal surface you need to expose most of the
uropygial gland, a white or yellowish bilobed organ. Usually you can’t
expose the whole organ without tearing the skin, so be careful. Cut and
scrape all of this organ away (some scooping action is often necessary to
get back under the skin). A few short feathers are present where this
organ connects to the surface (this is where the bird wipes its bill to oil
its feathers); cut this small ‘‘nipple’’ away from the inside or you’ll make
a hole. With the tail stub skinned, remove the remaining muscle by whit-
tling it carefully away. Some of the remaining bone can be easily cut or
broken away, but it is best to leave the pygostyle itself present in small
birds—it keeps the tail feathers firmly in place and too often the tail is
damaged in trying to remove it. Don’t cut the bases of the tail feathers
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(rectrices)! Give the cleaned tail and surrounding skin a dusting and rub
to clean it further and dry the surface, then put things back inside right.
12. Fleshing. Scrape off any remaining meat or fat from the inside of
the skin. This can be a very rapid step (e.g., in passerines with no fat),
or a very long, drawn out process (e.g., in migrating waterfowl and sea-
birds). For fat birds, special tools such as toothed spoons, scraping wires,
and wire wheels can be of great assistance in fleshing. With very greasy
birds it is often best to wash them thoroughly after fleshing, first in soapy
water, then in a solvent that dissolves fat (e.g., white gas, mineral spirits,
paint thinner, hexane). Such cleaned skins dry up very nicely. Oils left in
the skin leach out onto the plumage and also acidify over time, hastening
deterioration of the skin. Well-fleshed skins and formerly fatty birds that
have been washed in a fat-dissolving solvent are some of the best-pre-
served in museum collections.
At this point it is possible to put everything aside for awhile. Making
sure all relevant data have been recorded, the skin can be moistened and
frozen double-wrapped in a plastic bag
with a label
attached to the bird.
I leave the head inverted (except for species with sharp bills orlong head
feathers), but put the rest feather-side-out (especially the wings), then
place the bird with its label into a plastic bag, squeeze out all the air, and
bag it again before freezing. Some workers place moistened paper towels
or tissue against the skin in the cavity before freezing. The skeleton can
be made up now or set aside as the skin was.
13. Put the skin back inside right. Before putting the skin right side
out, moisten it thoroughly so that it is maximally flexible and each feather
can be adjusted (not possible with a dry skin). Saliva and fingers, or a
dish of water with a paintbrush or cotton swab can be used effectively to
put moisture exactly where it is needed. Moisten the head well, but be
careful not to get water on any feathers (especially through the eyes or
other holes). Don’t put on too much water—you just want the skin to be
limber. Dust the skin thoroughly to take off the stickiness, then put it
back inside right. The tail should already be reverted. Next revert each
wing to feather side out, spreading out the skin and feathers on each side
and rearranging the flight feathers and coverts into their proper posi-
tions. No bunched-up skin should remain anywhere between the wrists of
each wing (to minimize wing feather disarray, revertwings before freezing
if stopping before step 13). I should mention that it is fast and convenient
to ‘‘strip’’ the secondaries from the ulna, but that for this convenience
we must be assiduous in putting the secondaries back into their proper
position. Beginners often fail to do so, leading some authorities (e.g., Van
Tyne 1952) to condemn the practice. Now is a good time to check wheth-
er you’re satisfied with the distance between the tied wings and to change
it if necessary. To pull the head back through the neck, push the bill
along until you can see and grab it from outside. Pull it carefully out and
begin rearranging feathers generally.
This is the time to carefully inspect the plumage. Notes on molt and
fat, much of it best seen while the skin was everted, should be made now.
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Molting body feathers are easily seen on a fleshed skin as dark spots or
short lines, which represent the blood associated with growing feathers.
Flight feather molt must be assessed from outside. Details on fat are dis-
cussed below.
14. Washing. If dirt, blood, or other body fluids are present on the
plumage, you will have to wash these places clean. Use a mild soap and
water, and wash the dirty spots well. Rinse well, then dry. Whenonly small
areas or the head require washing, restrict the washing to these areas.
Drying a bird is a time-consuming process, so spot-washing should be
employed whenever possible. When the area (or bird) is well washed and
rinsed, it should first be patted dry with some toweling, then thoroughly
dried with forced air or with a combination of dust and forced air. In the
lab, the forced air is usually compressed air or,for larger birds, an electric
hair dryer (set on warm only—never hot). These tools speed the drying
of washed birds considerably and make laboratory preparation preferable
to field preparation whenever specimen washing is required. In the field,
the forced air is from one’s lungs.
When you have an adequate amount of dust, and when you are not
working with downy birds (e.g., owls), you can dry a washed bird relatively
quickly by shaking it about in a large, closed bag of cob dust. With a few
double handfuls of cob dust in the bag, close it and firmly shake it about
for 10–15 minutes, occasionally opening the bag and manipulating the
skin to knock off sticking wet dust and allow dry dust to enter wet areas.
In the later stages, remaining wet areas should be manually treated with
dust, brushing dry handfuls repeatedly over the wet feathers. When it is
nearly dry, remove the bird from the dust bag and shake most of the dust
free. Then, in a well ventilated area where airborne dust won’t offend
people or equipment, blow the bird clean and dry using forced air or by
blowing on it. If you don’t do this carefully, you’ll developa ‘‘dust cough’
from inhaling too much cob dust. A bird is dry when its downy under-
plumage is fully fluffy again. Note that it takes physical stimulationduring
the drying process to get the feathers to fluff up again. Much of the
drying process consists of providing this stimulation by rubbing, thump-
ing, or beating the drying feathers.
A ‘‘dry wash’’ is sometimes possible when mud or blood is present in
small amounts on generally dark plumage. In these cases, the dirty plum-
age can be held firmly in one hand while with the other hand clean, dry
cob dust (or cornmeal or sawdust) is rubbed firmly and repeatedly onto
the plumage with the grain of the feathers. White or very pale plumage
generally can’t be cleaned this way. Dried blood or mud can often be
chipped out by spot-rubbing with a thumbnail, with finer particles
brushed away using a stiff brush (e.g., toothbrush).
15. Build and insert a three-piece cotton head: two rolled eyes the size
of the bird’s eyes (the whole eyeball, compared in the orbit with the
cotton one; Fig. 9), and a piece of cotton formed to resemble the skull
without eyes in the orbits (Fig. 9). Preen the feathers and shape the head
while the forceps holding the last piece (the cotton skull) are still insert-
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IGURE
9. Shape smooth, firm, cotton eyeballs roughly the size of the original (compare
with carcass), and firmly insert first one, then the other. Next, roll a cotton skull in a
size and shape similar to the original skull without the eyes; firmly insert and arrange
head skin and plumage with other hand before releasing cotton.
ed. It is also easiest to arrange all of the pieces separately, and as a whole
each time one is placed inside, by working from inside with the forceps
still holding the cotton and from the outside with your other hand. Spe-
cific and subspecific identifications frequently rely on characters around
the eyes—make skins slightly bug-eyed. Symmetry is a strongly encour-
aged objective here.
16. Roll a cotton body onto a stick with a sharpened point, using the
carcass as a model and making sure the neck isn’t too long (Fig. 10). You
want a sturdy stick that is at least the total length of the final specimen.
Having a stick that projects to the tail tip protects against tail damageand
provides a sturdy anchor for the leg and label. Wet the stick so the first
cotton sticks to it, then wind long, narrow, thin pieces of cotton rather
firmly onto the stick with one hand while you rotate it with the other. In
this winding you are trying to mimic the length and girth of the main
body of the bird, which you should have in front of you for comparison.
Ignore the neck of the carcass, and make the body begin down from the
pointed end of the stick approximately the distance from the nostrils to
the base of the skull. Except in very long-necked birds such as waterfowl
and shorebirds, in which a neck of cotton must be built, in the final skin
you want the head cotton to rest against or very near the cotton body.
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IGURE
10. A cotton body is rolled in narrow, thin pieces onto a sharpened stickto resemble
the size and form of the original body without its neck; the symmetrical rolled body is
pressed into an asymmetrical shape resembling the original. The sharpened stick is
inserted up the neck and into the remaining spongy bone of the upper mandible, and
the skin is pulled onto the body.
Leave only enough stick emerging from the cotton body to allow this to
occur. When the length, girth, and rough shape of the carcass are mim-
icked by the rolled cotton body and the surface is generally smooth,make
one side the dorsal surface and the other the ventral by compressing one
side and pushing cotton up onto the other, obtaining a final shape very
similar to that of the carcass (Fig. 10). This enables the final skin to lie
flat on its back, with the bulk of the cotton lying in the ventral portion
of the skin, and prevents the skin from rolling in the museum tray.
17. Insert the pointed end of the stick into the skin (Fig. 10). Use
forceps to help open the passage to the mouth (made easier by dusting
away surface stickiness prior to reverting the skin). When you have the
pointed end of the stick coming out of the mouth, withdraw it to the
point where it goes over the edge of the severed base of the bill, then
ram it into the bone of the upper mandible, aiming roughly for the stick
to pass through the spongy upper mandible bone to come to rest in one
of the nostrils. Get the bill on straight, and don’t plug up the nostril or
it will be difficult to get a bill measurement from the skin. Preen the head
again.
18. Laying the skin and partially inserted body ventral side up on the
table, use forceps to compress the cotton body locally and draw the skin
up onto the body with your fingers, working carefully and evenly until
the body lies in the skin, essentially in its final position (Fig. 10). Any
adjustments to size must be made now. If thebody is too large, take cotton
off. If it is too small, add cotton in smooth, symmetrical contours where
necessary. A small piece of cotton inserted into the skin pocket at the
base of the tail firms it up. You should now be able to put the ventral
incision edges back in their proper relative places with only slight com-
pression of the cotton body.
19. Sew the skin back up. Tie leg to stick. Close bill. Bill must be both
pinned and tied. The lower mandible must be put into its proper place
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against the upper mandible, and is best held there with a pin through
the base of the gonys into the upper mandible. This maintains horizontal
alignment. A thread passed through the nostrils with a needle and tied
around the bill and behind the pin (rather tightly, but not crushing light-
billed birds) will keep it closed. In species with closed or inaccessible
nares, or species in which the operculum is of taxonomic value (e.g.,some
swallows), an alternative method of bill closure is to push the pin com-
pletely through the bill and pass the tying thread around the bill behind
the pin on both the dorsal and ventral surfaces. For long-billed birds such
as hummingbirds and shorebirds, a second tying out near the tip (without
a pin) assures permanent bill closure along its entire length.
Few stitches are required to close the ventral incision. Sewing should
always be done by bringing the needle up from the inside of the skin
through to the outside. I generally make about 4 loose, looping stitches
in passerines, ending on or beside the cloaca. Closing the ventral incision
is usually a rapid process that leaves gaps through which the cotton body
is visible. Plumage arrangement covers this in most cases. The ventral
incision should be carefully closed with closer stitches only in species with
dense ventral plumage (e.g., waterfowl), in individuals with a bare venter,
and in specimens that have been temporarily fluid preserved. When the
ventral stitches have been placed, insert forceps below the stitches and
on top of the cotton body, compress the body and, holding the needle
in one hand and forceps in the other, draw the stitches firmly but not
tightly closed in one pull. Holding the forceps in place, I next arrange
the ventral feathers with the other hand to be sure everything looks fine,
then hold and compress the whole body while carefully slipping the for-
ceps out. The thread is then tied firmly to the stick. Before cutting itfree,
I make sure that the tail is centered and doesn’t flop excessively when
the specimen is lifted. At this point one or two additional stitches in the
skin at the base of the tail can center it and hold it firmly against the
stick if required.
The remaining leg should be tied to the stick midway along the tarso-
metatarsus with the leg in a relaxed, ‘‘natural’’ position that leaves it with
the sole of the foot pointing toward the tip of the tail and with the tar-
sometatarsus fully exposed and available for measurement (e.g.,passerine
in Fig. 11).
20. Preen and rearrange feathers. Most feathers are best rearranged by
working from their bases. Long, narrow forceps and in some cases a long
rigid wire are very effective; brushes are not. In many species with narrow
tracts of feathers on the neck (e.g., most passerines), the neckskin usually
must be unstretched and slightly bunched (as in the natural bird) by
pulling the skin near the bend of the wing upward towards the cheek;
also try pulling a little skin downward from just below the cheek toward
the bend in the wing. Tuck any visible plumage bases into their proper
places below surrounding contour feathers and arrange these contour
feathers over the bases still exposed. If the neck of the cotton body and
stick was too long, you can try to push the whole body forward to un-
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F
IGURE
11. The sewn skin’s plumage is carefully arranged, then the whole is pinned out in
final form to dry. Long-necked birds should have their necks bent to shorten the spec-
imen, minimizing space required for packing and in the museum tray.
stretch the neck skin; this is best noticed and done before the stitches in
the ventral incision are tied off. Twisted feathers disrupt or break the
smooth contours of the feather surface, and can usually be untwisted with
forceps. Wings and the thread tying them together must sometimes be
hiked forward into their proper places. Primaries, secondaries, tertials,
and scapulars should all be properly aligned and placed before the bird
is finally laid on its back onto the pinning board.
21. Pin skin out in its final form to dry. Pinning on drying boards (I
use styrofoam) is preferable to the traditional wrapping in a thin layer of
cotton or gauze, because it is easier to learn and the preparator has full
control and access to most of the specimen (Fig. 11). The wings should
be on the back, with the wingtips under the tail (i.e., lying on its dorsal
surface); I hold it all thus with one hand until the first four to six pins
are in place. I begin pinning by crossing a pair of pins over the end of
the stick near the tip of the tail and another pair over base of the bill to
anchor the whole bird, then pin the wings in place. Pins are then placed
wherever required to obtain a smooth, symmetrical form. The whole spec-
imen should be symmetrical and well preened; I usually have a narrow
pair of forceps in hand for this during final pinning. At times, pinning a
thin piece of cotton over part of the venter helps to smooth it (especially
in fluid preserved birds), and wrapping the whole in gauze after a day of
drying can smooth the contour of the whole bird. The drying skin can
be unpinned or unwrapped each day to be checked and adjusted until
dry. Large birds usually require a combination of pins and gauze, and
should be rotated almost daily to allow complete drying and to prevent
mold and rot. See below for more details on drying specimens.
22. Remove the muscular outer sheath of the stomach by cutting it
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open longitudinally and peeling out the inner sac, in which the contents
lie. Open this sac halfway without spilling the contents and put the whole
thing, contents and all, into a small vial. Fill vial with 70% ethanol, tightly
screw on the cap, and shake vigorously. Examine.Write a briefdescription
of the contents in the catalogue. Some large birds lack a removableinner
sac (e.g., some raptors, waterfowl). In some species a crop may be present
that contains more intact food; preserve some or all of this material sim-
ilarly.
23. Review the catalogue to be sure all data are recorded there. See
Labelling and Field Catalogues above and Figure 1.
24. With all of the data now complete in the catalogue, and with clean
hands, write labels: one for the skin, one for the skeleton, and a small
one for the stomach contents vial (Figs. 2 and 3). If skin and skeleton
are going to be catalogued into the same collection, the skeleton label
can be less detailed than the skin label (e.g., leaving the back largelyblank
except for mass, sex, number, and disposition). But the field number
should be written twice on the skeleton label and care should be taken
not to get body fluids or blood on it, or the affected parts may be eaten
by dermestid larvae during processing. At the University of Alaska Mu-
seum we catalogue the skin first (when it has dried), directly from the
complete label onto the computer. This computerizes all of the data as-
sociated with a specimen and enables the printing of a final label for the
skeleton, whose original label is retained but is usually marred, stained,
and perhaps even physically damaged from exposure to dermestids dur-
ing the process of skeleton preparation.
25. Tie the labels properly (Fig. 3), and attach the skin label to the
skin, tying it around both the leg (midway on tarsometatarsus) and the
stick to add strength and prevent future damage through accidental pull-
ing.
26. Prepare skeleton for drying. Remove eyes from sockets. Pop them
and squeeze out the fluid. Put them into the body cavity. No skeleton is
complete without the sclerotic ring, a ring of bone plates in the eye. Skin
or at least open up the skin on the unfeathered portion of the leg and
toes to facilitate dermestid access. After cleaning the bulk of the meat
from the leg and wing bones, place these lengthwise into the body cavity
(Fig. 12). Using a light cotton thread, and holding the head against the
side of the carcass, firmly (but not too tightly) wrap up the skeleton,
making sure the pieces in the body cavity are nestled well inside and their
ends wrapped to keep them in place. Tie off the end, then loosely tie the
skeleton label around the neck with clean hands, using forceps to insert
the label thread around the now wrapped neck (Fig. 12). Hang whole to
dry or place in a screened box in a well ventilated place safe from scav-
engers and generally free of insects. Watch drying skeletal material to
prevent mold growth and insect infestations.
27. Put small label for stomach contents into vial with contents and
alcohol after the ink has dried (Fig. 3). Be sure it is legible in thealcohol.
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F
IGURE
12. The carcass should be cleaned of large pieces of meat and internal organs. Eyes
should be removed and the fluid squeezed out. Loose pieces, including eyes, shouldbe
put inside the body cavity and the whole well wrapped together in light thread, then
labelled and dried. Second tibiotarsus and distal half of one radius and ulna can be left
with skin to increase strength and to leave one wrist intact.
I also write the field number (twice) and species in indelible marker on
the vial cap.
SEXING
,
AGING
,
FAT
,
AND SPECIMEN CARE
Determining sex.
—Properly sexing each specimen is important. Some-
times it is impossible because a bird is too decomposed, or has been too
badly shot, but it is generally possible, and a specimen of known sex is
much more valuable to science than one whose sex is unknown or has
been inferred from external features (in species where this is possible).
Direct examination of the gonads is the preferred way to sex every bird.
In many species sex can be inferred from plumage (sexually dichromatic
species) or from the presence of sex-specific incubation patches (some
species), but these inferences give us little or no detail about the present
reproductive state of the individual.
Care must be taken not to disturb the dorsal region of the body cavity
near the adrenal glands and the proximal ends of the kidneys until sex
has been determined. Most errors in sexing, and the greatest source of
unsexed specimens, arise from careless and overly exuberant removal of
organs from the body cavity when taking tissues and gutting the specimen.
Both sexes have a pair of adrenal glands that lie against the dorsalsurface
of the body cavity at the anterior, proximal ends of the kidneys (Fig. 5).
Finding the adrenals helps orient the search for the gonads and prevents
mistaking adrenals for testes in females by maintaining familiarity with
the variation that occurs in adrenal size and color. In most immature
birds, the basal third of the left adrenal is usually overlaid by gonadal
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tissue, whether the bird is male or female. A stereo microscope or a hand
lens in good light is an asset when preparing autumn and early winter
specimens.
When found, gonads should be measured and described. Measure-
ments include the maximum length and width of the organ(s), and the
diameter of the largest ovum in females. Descriptions can include such
things as the state and size of developing ova or eggs, condition of the
oviduct, degree of differentiation of the ova, and color of the testes. Usu-
ally only the left testis is measured (usually being the largest), but care
should be taken to observe both to be certain the bird is male. Mostavian
females have only one ovary (the left), but some (e.g., Falconiformes)
have two. Figure 5 represents a nonbreeding adult of each sex.
Sexing adults during the breeding season is easily done because the
gonads are usually very large. During the nonbreeding season, however,
gonads can often be difficult to find because they are at their smallest.
This is especially true in first year birds, whose gonadal tissues have often
not yet become fully developed. The ovaries of young females are often
not yet granulated with ova, and the testes of young males may not yet
be opaque. First year autumn Parulidae, for example, can be quite diffi-
cult to sex unless very fresh or frozen quickly after death. When gonads
are difficult to find, rinsing the body cavity in cool water (after skinning)
can help, as can soaking the carcass in a bowl of water in a refrigerator
for an hour or more. This usually causes the gonadal tissue to become a
different color than kidney and adrenal tissues, making them easier to
discern.
External indicators of reproductive condition should be noted with sex
in the catalogue and on the label. Presence and/or condition of cloacal
protuberances and incubation patches (sometimes inappropriately called
‘‘brood’’ patches, after their secondary function) are normally recorded.
McCabe (1943) and Foster and Cannell (1990, and references therein)
described avian reproductive conditions in greater detail.
Aging by skull ossification.
—Skull ossification is a useful aging charac-
teristic in most passerines and some other birds. As these birds get older
(usually in the first year), the cranium gradually changes from a single
bone layer to two layers. The two-layered cranium develops along a front,
and the layers are separated by small bony pillars. As the two layers be-
come fully separated and ossified along the developing front, the small
space between the layers fills with air (becomes pneumatized). Many au-
thors refer to this entire process as ‘‘pneumatization,’’ which is inappro-
priate. Ossification precedes pneumatization, and, moreover, natural
stresses to the crania of wild birds frequently cause unpneumatized re-
gions to develop in fully ossified adult birds (pers. obs.; see also figures
in Klem 1990a). In hand-held living birds it is the contrast causedbetween
pneumatized and unpneumatized regions of the skull (observed through
the skin) that enables the bander to determine the stage of ossification
and pneumatization. Ossified, pneumatized areas are pale; unossified re-
gions are darker because brain tissue and fluid lie directly against the
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inner side of the single layer of developing cranial bone. Unpneumatized
areas are also darker, because fluid lies against the outermost layer of the
cranium (whether there are one or two layers). The contrast between
ossified and unossified areas is also easy to see while skinning, and the
best time to record the condition of ossification is usually when the skin
is first peeled from the head.
Unossified regions of the skull usually occur in bilateral symmetry,and
are readily discernible when large. Unossified areas are always unpneu-
matized, but the opposite is not true. When asymmetrical unpneumatized
areas are seen, look very closely to determine the actual state of ossifi-
cation: observe the distribution of the small points caused by the little
bony pillars separating the two cranial layers. Small, roughly symmetric
unpneumatized areas (often termed ‘‘windows’’) should also be carefully
examined for bone growth; they may represent the final stages of both
processes. Good light and a hand lens or stereo microscope can be help-
ful in close examination. The state of ossification should be recorded as
an estimate of the percentage of the cranium in which bone growth is
complete, stating whether the skull is ossified, or, if not, approximately
how much is ossified (e.g., 5%, 10%, 25%, 50%, 75%, 90%, or 95% oss.).
The degree of ossification is important, particularly in species where the
entire process takes more than a year. Ossification pattern is often
sketched in catalogues and on labels, but this information is not readily
computerized and is retrievable from the partial skeleton.
One advantage of the preparation method outlined here is that the
skull is available for later examination. In birds that have died of head
trauma (and in fluid preserved specimens) it can be impossible to deter-
mine stage of ossification until the skeleton has been fully prepared. Care-
ful study of skulls to verify age following skeletal preparation has also
proven very important in my own studies in which knowing the age of a
bird is critical, especially in many species of Tyrannidae and Turdidae,
wherein complete ossification is often not attained in the animal’s first
year of life.
Determining the age of a bird, whether from plumage stage or from
the degree of ossification, often requires experience. During and imme-
diately after the breeding season ‘‘ad.’’ for adult, ‘‘juv.’’ for individuals in
juvenal plumage, and ‘‘im.’’ for individuals in first basic plumage with
unossified crania are sufficiently simple, generic descriptors that fit most
passerines. If in doubt, don’t write anything except skull and gonad con-
ditions, which can help subsequent investigators determine age more ac-
curately. I use some bird banding age categories because they are more
refined than traditional museum aging descriptors (Department of the
Interior 1981, 1991), but they require some care in their use. Briefly, after
‘‘nst’’ for nestling and ‘‘juv’’ for individuals in juvenal plumage, the age
categories I regularly use from banding literature are: ‘‘HY’’ (hatch year)
for individuals still in the calendar year of their hatching; ‘‘SY’’ for indi-
viduals in their second calendar year; and ‘‘TY’’ for individuals in their
third calendar year. In many species and individuals it is not possible to
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determine age after the HY period, and in these cases they are denoted
as ‘‘AHY’’ for after hatch year. After second year (ASY) and after third
year (ATY) categories are also possibilities. The peer reviewed literature
should be consulted for detailed aging criteria on a species-by-species
basis.
The bursa and its measurement.
—In most nonpasseriformes skull ossifi-
cation is not particularly useful for aging. In some groups, such as Anse-
riformes, Galliformes, and Charadriiformes, the size of the bursa of Fa-
bricius can be useful in distinguishing first-year from older birds (Hoch-
baum 1942, Kirkpatrick 1944, McNeil and Burton 1972). The bursa is an
ephemeral, pouch-like organ that lies dorsally above the cloaca inside the
body cavity (see Proctor and Lynch 1993). In game birds it opens into
the cloaca and its depth is measurable in the living bird, but in shorebirds
it is not so accessible. The length and width of the bursa is readily mea-
sured in preparation, however, and outside of game species it is an un-
derutilized method of aging. Care must be taken in these taxa when cut-
ting the spine above the tail not to damage the bursal region. Do not
confuse the bursa with the caeca, which are paired and occur more prox-
imally along the intestine.
Fat.
—The amount of fat that a bird is carrying is important information
best recorded after the bird is skinned. A six-level scale is generally used:
none, very light, light, moderate, heavy, and very heavy. With experience,
these divisions, although representing a continuum, become fairly obvi-
ous. They are based on the amount of fat found in the feather tracts,
furculum, abdomen, intestines, and on the skin. Fat is assessed more thor-
oughly in skinned specimens than banded birds (contra Foster and Can-
nell 1990), although a six-level scale may be used in both. McCabe (1943)
and Foster and Cannell (1990) both described a six-level fat scale, but
their descriptions differ considerably, primarily because the latter was de-
veloped from passerine banding literature.
Briefly, ‘‘none’’ is recorded when little or no fat exists anywhere on the
bird (even a starving bird can retain tiny amounts of fat in the abdominal
cavity and on the dorsal tract). Very light fat is indicated by the presence
of some fat in the dorsal tract and a trace in the furcular area (usually
adheres to skin). Light fat adds depth to the former and some to the
abdominal cavity. Moderate fat continues adding depth to the dorsum
and furculum, and includes significant abdominal fat (inside and out) as
well as small plates or pads of fat on the sides and elsewhere on the skin.
Heavy fat is used when all of the feather tracts and much of the skin is
covered with heavy pads of fat and large deposits are present in the ab-
dominal cavity. At very heavy fat levels the body is entirely encased in
thick fat, as are the intestines; fleshing becomes difficult because the skin,
being stretched and greasy, is thinner and weaker.
Drying specimens.
—Exposure of the pinned or lightly wrapped skin to
air circulation is crucial until it is completely dry. A specimen is usually
dry when its toes are no longer flexible. In warm and dry environments,
drying specimens is usually not a problem; small birds can be fully dried
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in as little as a day. Drying is aided considerably by moving air, and an
electric fan is a superb tool for assisting the process, especially with large
birds. Drying specimens is often a challenge under humid, wet, or cold
conditions. Screen-bottomed racks over a stove set to give off a low heat
are very effective; warm, dry air passes over the specimens from below.
Care must be taken to keep heat levels low and to avoid exposure of
specimens to smoke or flames. Putting skins in direct sunlight should be
avoided; it causes plumage fading. However, using strong sunlight on a
shade under which the birds are placed with adequate ventilation is ef-
fective because it provides heat. A refrigerator will slowly dry a specimen,
and provides a good start on the drying process. It will also kill ants in
tropical environments. Rotate large birds periodically during drying.
Drying skeletons is as challenging as drying skins under poor condi-
tions, but the process can be assisted by immersing the thread-wrapped
and labelled skeleton in 95% alcohol for a short time, then hanging it to
dry. In all cases, dry skeletons slowly or the long bones may permanently
warp. This is a problem when artificial heat is used.
Pests, and care of prepared specimens.
—Arsenic, the effective poison and
old museum standby, is no longer considered safe for human exposure.
Even borax, a commonly used natural soap and mild insect repellent, has
been eliminated from use in bird skin preparation because it has been
implicated in accelerated foxing (color changes) of plumage (e.g., Phil-
lips 1991:xliii; but not without some disagreement, K. C. Parkes, pers.
comm.). Consequently, we use no chemicals when preparing bird skins.
Borax does remain effective, however, when dusted on drying skeletons
to aid drying and to provide a mild insect deterrent (D. Causey). Such
skeletons must be well rinsed before putting them in a dermestid colony.
Keeping insects from specimens as they dry and before they are safely
transported to a museum collection is an obligation and challenge. When
completely dried, unpinned or unwrapped, and labelled, skins should be
placed in a dark place where they are not susceptible to insect pests. Ant
repellent or poison regularly sprayed or dusted onto pinning boards (
not
onto the birds) helps during the drying process. Paradichlorobenzene
(PDB) crystals, a preferred form of ‘‘moth balls’’ still readily available
commercially, will temporarily prevent insect incursions following drying
and temporary packing when it is sprinkled liberally in a field case or
specimen packing box (cardboard boxes work well). Specimens should
be regularly inspected for insect activity. Personal exposure to the really
effective insect-deterring chemicals should be minimized. For reasons of
human health and skin specimen longevity, most museums now use no
chemicals on the skins themselves, and also minimize use of insecticides
and repellents.
Cryofumigation has become a popular method of ridding specimens
of pests. Freezing specimens at
40 C for at least 48 h appears to kill all
insect pests. Higher freezing temperatures (normal freezers rarely get be-
low
20 C) are not completely effective, but insect activity is usually com-
pletely retarded. High concentrations of PDB probably kill and at least
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severely retard the development of most insects. Together, two weeks at
20 C and a month in a case with a high concentration of PDB have
proven an effective and legal method of fumigation. Every specimen com-
ing into a collection or specimen case should be put through a fumigation
process first. Insect damage is irreversible, can be rapid, and can also
spread.
Skeletons.
—The method outlined above generates labelled, roughed-
out, partial skeletons. Upon drying these are ready to go to a dermestid
colony for cleaning by beetle larvae, and subsequently the cleaning may
be finished through cold water maceration. Matthiesen (1989) provided
excellent details of the complete skeletonization process. If the further
processing of dried, roughed-out skeletons into clean, museum-quality
skeletal specimens is not deemed cost effective or appropriate, be aware
that several museums will accept the donation of these specimens (prop-
erly labelled, of course) and complete the preparation process.
Speed.
—We want to produce a high quality specimen in as short a time
as possible. The quest for speed should never compromise the quality of
the final product, however. Useful specimens with complete and legible
data are, after all, the point. Blake (1949), noted for his preparation
speed, recommended maintaining an orderly work table, placing instru-
ments in a logical and consistent order, avoidinguncertain or unnecessary
movements by focusing on the job, and keeping close track of time while
practicing to increase speed and efficiency. Although most of his methods
are outdated, these suggestions remain effective. Keeping the number of
tools used to a minimum also helps.
With changes in preparation methods and data quality standards, the
time required to prepare specimens has increased. Coues (1874:71), us-
ing primitive methods that retained neither skeletal material nor tissues,
and which produced skins often damaged through even normal use, con-
sidered 15 min per skin in small birds to be good work. With practice,
the method outlined here requires 40–90 min per specimen in passerines,
everything included (each specimen being preserved in five pieces with
associated complete and partial labels). It is washing and drying that push-
es the time required to the upper limits. On average I can make up one
specimen per hour.
When in the field, collecting and preparing 10 specimens a day is ex-
cellent work. If preparation is delayed until return to the laboratory, how-
ever, 20–30 specimens or even more can be collected in a good field day.
Holes.
—When started, care must be taken not to enlarge a hole. Unless
very large, most holes do not need to be sewn closed. They can usually
be hidden by a final pull of the skin and arrangement of the plumage,
or sometimes by slightly understuffing the bird at the affected spot. When
it is necessary to sew a hole closed, one must be careful not to allow
stitches to pull feather bases awry, for it is difficult to get such feathers
to lie smoothly again.
Places students tend to make errors.
—Treating the plumage roughly dur-
ing skinning. Don’t brush feathers away against their grain; don’t blow
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indiscriminately to clear an area. Instead, move feathers gently and care-
fully away from incisions or exposed skin by pushing or brushing them
with the grain. The kinder you treat the plumage during skinning, the
easier it will be to finish with a good-looking skin.
Using the scalpel too much in skinning: you only need it to make the
initial incision, disarticulate, cut meat, and cut eyelids. Otherwise, use
thumbnails, pressing skin away from the carcass (do not pull the skin).
Using the scalpel elsewhere results in holes.
Duplicating the bird’s neck in building the cotton body, resulting inan
ugly skin with its neck stretched out too far. In theirnormal posture, most
birds carry the head close to the body by bending the neck rather se-
verely. In living passerines, for example, the furcular area is usually filled
with neck. Make the cotton body up close to the tip of the stick, leaving
only enough space (or just slightly more) for the stick to go into the
upper mandible (to the nostril) and the head to lie close to or against
the cotton body. All of the neck feathers should be visible, but the neck
should not be particularly outstretched. Long-necked species (e.g., many
Scolopacidae) and those in the Picidae and Anseriformes do require at
least some neck to achieve a decent looking skin.
Not using enough dust. Pour it on.
Feathers sticking out in the lower neck and shoulder areas. This is
caused by stretching the neck skin, which is usually folded with the neck.
Before the skin dries, bunch the neck skin as described in step 20 above.
Twisted feathers disrupt the ordinarily smooth surface of the feathers.
Grab the offending feathers before the skin dries and twist them back to
where they belong (try both directions).
Head: make eyes the same size as the original eyes or just slightly small-
er; make the head cotton the same size or slightly larger than theoriginal
skull minus the eyes. When putting in eyes, push the cotton well into the
head skin, stretching the eyes outwards on the skin.
Arrange leg so that it is oriented correctly and the claws do not pro-
trude beyond limits of tail. They tend to catch on packing cotton and
other specimens, leading to damage. Also arrange it so the tarsus (tar-
sometatarsus) is readily accessible to measurement: keep it in the plane
of the pinning board with the proximal end just exposed.
Tie label around both the leg and the stick together to increase the
strength and durability of the specimen. Leave one inch (2.5 cm) of dou-
bled thread between specimen and label. Less distance makes it difficult
to measure the tarsometatarsus, examine undertail coverts, or read both
sides of the label. Less distance also exacerbates problems of labels getting
tangled in claws. However, a longer thread between specimen and label
causes the label to become tangled with other labels and specimens.
Be sure to clean out the tail properly!
Body form: use the carcass as a model for the cotton body, and strive
for smoothness, symmetry, and proper form in the final specimen. The
skin usually dries rapidly after being put inside right. Once the cotton
body is inserted, the preparator often must work quickly to get everything
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into proper place while the feathers and skin are still manipulable. In
slowly drying birds one can inspect and rearrange things the next day.
Completely evert wings, so that skin around the ‘‘wrist’’ is not bunched
or wrinkled. Before adding any cotton, extend wings and arrange all wing
feathers, especially secondaries and tertials, into their proper order and
positions.
Labels properly tied (knots and distances) and written. Accurately fol-
lowing the directions given is not difficult, and the proper result is evi-
dence of professionalism.
VARIATIONS
Large heads.
—The heads of Picidae and Anatidae are generally too
large to be removed through the neck. In these species, an incision is
made either in the back of the head or in the neck and throat, usually
whichever side will not be facing upward in the final skin. This incision
is most easily made from the inside as one is skinning the head, and
should be sewn up from the inside as well to most easily keep feathers
on the outside.
Long necks.
—Birds with long necks should generally be made up with
the neck folded back along the body to save space in packing and in the
museum case. In these birds the neck can be made up only of cotton or
of cotton and wire. The head should be attached to the body with firm
stitches of heavy thread, leaving the bill accessible for measurement.
These stitches can be placed through any convenient part of the head
skin, or in some cases the nostrils.
Large birds.
—Large birds (e.g., large
Buteos
or
Larus
and up) usually
require more substantial stuffing than just cotton. Excelsior, also called
tow or wood wool and available from taxidermy supply houses, shouldbe
used at least as a body core, with a light cotton wrapping. To make ex-
celsior bodies very tight and compact, wet the excelsior with warm water
prior to forming and wrapping with thread, and let the body dry over-
night. Stuffing large birds also requires a lot of material, not always readily
at hand. I have had to resort to making rather flat specimens of gulls in
the field (using cardboard forms), and to using mosses, crushed leaves,
and accumulated cotton scraps. L. Stejneger once used his socks to stuff
a gull (D. Causey, pers. comm.), and K. Parkes (pers. comm.) has had to
use newspaper and shredded blueprints. Try to use archival quality ma-
terials, and, as usual, use the carcass as a model for size and shape.
In large birds it can occasionally be difficult to both tie the wings and
to get them to dry against the body without flopping down. In very long-
winged birds (e.g., Procellariiformes), I often find it easier to leave inter-
nal threads tied to each ‘‘wrist’’loose in the body cavity until finalstuffing,
and then tie them together within the bird, or foregoing internal thread
entirely and stitching the wrists to the skin in the proper position from
outside, using heavy thread. Whenever doing the latter, you must besure
not to place stitches so they inhibit measurement of individual flight
feathers (an abysmal old practice used by some preparators).
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Shmoos.
—When skeletal material is favored over a skin comparable to
historic material, more bones can be removed during preparation for
preservation with the skeleton—including the complete skull. Bill-less
skins are called ‘‘shmoos’’ or ‘‘muppets’’ in the museum community, and
provide researchers with an essentially complete skeleton and a skin use-
ful for plumage analyses. The biggest difference in preparation is to skin
the head completely, carefully cutting the skin away from the base of the
bill from the inside with a scalpel when that area is reached at the end
of the skinning process. In some species, much of the bill itself can be
skinned. The head of the skin must be reconstructed more carefully than
in the usual skin. Use the same amount of cotton for eyes and skull
replacement, but leave final arrangement until after the stick is in place
and the mouth sewn up. Insert the stick and cotton body as usual, but
make the stick project just a little beyond the mouth. (I usually do not
use a sharpened stick for shmoos). Using close, symmetrical stitches and
working the needle always from the inside out, sew the mouth shut
around and to the stick. A shmoo can be made up with no bones at all,
but duplication of whole bones with the skeleton comes at great cost to
a useful skin and should rarely be worth it. Dickerman (1989) provided
more details on shmoo variations.
Open wings.
—In a traditional skin, the closed wing prevents detailed
examination of primaries, secondaries, and the inner wing surface. In
many species (e.g., Dendrocolaptidae, Scolopacidae,
Catharus
) important
plumage patterns are thus hidden from view. Although it is possible to
prepare a skin with one wing open, it makes the specimen fragile and
difficult to handle, pack, and keep in the museum tray. When separated
from the skin, however, open wings are useful and easy to accommodate.
Open wing collections may have begun as a means to save something
of the skin from specimens otherwise made up as complete skeletons, or
from hunter-taken gamebirds. They are easy to prepare, requiring no
fleshing after the radius and ulna have been removed. Whether preparing
open wings during complete skeletonization or as a by-product of the
method outlined above, the methods are the same. The wing is taken off
at the elbow by cutting the skin and disarticulating the bones. It is then
skinned to the distal end of the radius and ulna. In small birds these
bones are disarticulated and removed for the skeleton; in large birds skin
out the hand bones as well. The wing is then put inside right and flight
feathers and coverts carefully rearranged on the pinning board. The wing
should be pinned out in a naturally outstretched position; examine the
wing before removing it from the bird to see what this should be. Artists
often find these preparations to be some of the most useful in a museum
collection. Open wings are also helpful for studying molt, particularly if
the preparator had the sense to note whether any present was symmet-
rical.
To complete a skin preparation when one wing has been removed,
Spaw (1989) presented the useful ‘‘button-stick’’ method. Instead oftying
the outer wings together as in step 10 above, one uses a short stick as the
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second wing and uses the same distance (or a little less) when tying this
stick (at its center) to the remaining wing. This stick is then placed
through the hole where the second wing was removed, there serving as
a ‘‘button,’’ giving firmness and symmetry to the remaining wing and the
complete specimen. With practice, a skin made up without one wing can
have perfect form and be difficult to separate from an ordinary skin with-
out turning it over.
Flat skins.
—The traditional round skin is most comparable to historic
preparations, but may not always be desirable or possible to prepare. For
example, when processing a large number of specimens is necessary and
the job can’t be done effectively given the time required by the method
outlined above, much time can be saved by preparing a flat skin and a
partial skeleton instead. Similarly, a series of large birds takes up less space
and can be prepared more quickly and effectively as flat skins and skel-
etons.
The method I use is to make an incision on the left side of the bird
from the left side of the mandible, under the left wing, to the distal end
of the left tibiotarsus. The shoulder is disarticulated first, then the skin is
removed from the left leg and the bird is skinned nearly the same as
usual, except that the right leg stays with the skin (the cotton around the
right tibiotarsus provides strength to keep the leg attached). The right
wing is removed at the elbow after skinning, and a cotton eye is inserted
into the right eye. The bill is closed as usual with pin and thread, and
the bird is pinned out skin-side down with the right leg cocked along the
bottom of the specimen and the left wing, folded, pointed backward
along the top side of the flat skin. The wing that was removed is pinned
out as an open wing. A single label is threaded through the skin of the
elbow area of the open wing and attached to the flat skin in a loose loop
through the right wing hole. Storing these specimens in archival plastic
sheets (photo album pages for small birds) maintains their integrity and
prevents damage. All standard skin measurements can be taken, and col-
orimetric readings can be made as well, provided care is taken in plumage
arrangement.
In small birds this preparation can be completed in 30 min or even
less. It is particularly effective in processing partially freeze-dried speci-
mens, when an aesthetic round skin is unlikely anyway. Spaw (1989) and
Garrett (1989) presented other methods of producing flat skins, but, un-
like the method presented here, neither produces a specimen that pro-
vides all the data obtainable from a round skin.
FINAL DISPOSITION
Scientific specimens should be deposited in a collection at an institu-
tion having a demonstrated long term commitment to specimen care and
use. Such collections are most likely to be successful as long term repos-
itories, and the specimens there will be most accessible to present and
future researchers. Specimens are routinely loaned and borrowed among
institutions through the mails, so a specimen’s geographic location is of-
K. Winker
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ten not particularly important. When deciding where to deposit speci-
mens, considerations should include: whether the institution has perma-
nent staff dedicated to collection care and use; the permits held by the
institution (e.g., CITES, USFWS, USDA-APHIS); whether their collection
is computerized and accessible over the internet (enhances accessibility
to the research community); whether agreements are extended to re-
searchers if specimens are being deposited while still part of a research
project; proximity to the researcher if specimens are still needed; and
international accessibility (trustworthy postal services). Institutions cannot
accept illegally taken specimens except under very special circumstances.
There is no current directory of the world’s bird collections. However,
many museums (including the University of Alaska Museum) will accept
legal, prepared, data-bearing specimens for long term archiving, making
them accessible to the world research community in agreement with the
depositing individual. Getting specimens into proper long term archiving
facilities in a timely manner is very important. Damages caused by insects,
mold, light, etc. are considerable and irreversible.
Birds are packed for transportation or shipping in wooden or double-
walled cardboard boxes. Specimens are usually wrapped in tissue paper
to prevent cotton from catching on them, and are then packed densely
and completely surrounded by heavy cotton. Boxes are packed full and
with slight pressure so that there will be no settling or shifting of contents.
Shipping labels should state, at a minimum, that the contents are scien-
tific specimens of no commercial value. A complete inventory of the con-
tents and copies of associated permits should be placed in an envelope
on the outside of the package for any international shipping. Boxes of
specimens are usually sent insured against loss.
CONCLUSION
There are many ways to prepare bird specimens. The general method
outlined here has been developed from a lot of experience. It seems to
maximize the usefulness of every bird prepared while not overly compro-
mising the number of preparations one can perform. Skin-only or skel-
eton-only preparations can be finished faster, and may be preferable in
special cases. However, sample sizes and overall specimen quality are
greatly enhanced when specimens are prepared as combination preps. I
have not considered many types of specialty preparations used daily by
research specialists—fluid preservation (‘‘pickling’’) and the preparation
of eggs and nests, for example. Persons wishing to learn more of these
and other methods should begin with the excellent information assem-
bled by Rogers et al. (1989) and Rogers and Wood (1989).
ACKNOWLEDGMENTS
I have benefitted from the experiences of others as much as from my own preparation,
use, and repair of specimens. For ideas, observations, conversations, and lessons, I am in-
debted to Clyde Pasvogel, Dwain Warner, John Klicka, S. Peter Getman, Bruce Fall, Gary
Graves, Ralph Browning, Steve Rogers, Mark Robbins, James Dean, Brian Schmidt, Phil An-
gle, Chris Milensky, Travis Glenn, Steve Cardiff, J. Van Remsen, Daniel Gibson, Douglas
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Causey, and Robert Dickerman. For the bird illustrations I thank Saitra Perez. A series of
field workshops in Mexico sponsored by NSF (INT-9403053) and CONACyT provided the
opportunity to develop emphases in teaching students to prepare specimens as outlined. K.
C. Parkes, D. D. Gibson, and an anonymous reviewer gave helpful comments on the manu-
script.
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APPENDIX
Sources of Supplies and Services
These sources are not meant to be endorsements; they are simplyplaces
with which I have done business in recent years. All are in U.S.A., but
many deal internationally.
Skinning implements, cryovials.
—1) VWR Scientific Products; many lo-
cations. Tel. 1-800-932-5000. 2) Fisher Scientific, HeadquartersPittsburgh,
711 Forbes Avenue, Pittsburgh, Pennsylvania 15219-4785. Tel. 1-800-388-
8355.
Vernier calipers.
—Forestry Suppliers, Inc. P.O. Box 8397, Jackson, Mis-
sissippi 39284-8397. Tel. 1-800-647-5368.
Stomach content vials.
—Perfector Scientific, Inc., P.O. Box 91, Atasca-
dero, California 93423. Tel. 805-466-8497 (Securi-Vial, 7 ml, #2142).
Cotton.
—1) Henry Schein, Inc., 255 Vista Boulevard, Sparks, Nevada
89434 (KenVet KV 2287, non-sterile veterinary cotton in 1 lb. rolls). 2)
Custom Hospital Products, Inc., 2623 S.E. Raymond, Portland, Oregon
97202. Tel. 503-231-9663 (70% non-sterile, non-absorbent cotton, 30%
polyester fiber, ‘‘Custom Cotton with Polyester’’).
Label and heavy sewing thread.
—Coats & Clark, Consumer Services, P.O.
Box 12229, Greenville, South Carolina 29612. Tel. 1-800-648-1479 (#220,
color 1).
Cob flour (dust).
—Mt. Pulaski Products, Inc., P.O. Box 100, 904 N Vine,
Mt. Pulaski, Illinois 62548. Tel. 217-792-3211. (#6 corn cob flour)
Mist nets.
—1) AFO Mist Nets, Manomet Bird Observatory, Box 1770,
Manomet, Massachusetts 02345. 2) Avinet, Inc. P.O. Box 1103, Dryden,
New York 13053-1103. Tel. 1-800-340-6387.
Auxiliary barrels.
—Robinson’s Gun & Tackle, 855 Street Road, South-
ampton, Pennsylvania 18966. Tel. 215-357-7381.
12 shot.
—Murmur Corporation, P.O. Box 224566, Dallas, Texas 75222.
Tel. 214-630-5400.
Reloading supplies.
—1) Gamaliel Shooting Supply, 1525 Fountain Run
Road, P.O. Box 156, Gamaliel, Kentucky 42140. Tel. 502-457-2825 or 2830.
2) Graf & Sons, RR 3, Highway 54 South, Mexico, Missouri 65265. Tel.
573-581-2266.
High quality hand binding for notes and catalogues.
—The Mount Pleas-
ant Bookbinders, HC 71, Box 38-B, Augusta, West Virginia 26704.
Institutional binding for notes and catalogues.
—1) University of Minne-
sota Bindery, Room 180 PSB, 2818 Como Avenue SE, Minneapolis, Min-
nesota 55414. Tel. 612-626-1516. 2) Page Book Binding, P.O. Box 187,
1891 Trumansburg Rd., Jacksonville, New York 14854. Tel. 607-387-4387.
... Para garantizar registros más completos de la alimentación de estas especies, las aves fueron capturadas al menos una hora después de iniciada la actividad de forrajeo dentro del área. Los individuos fueron sacrificados siguiendo el método de compresión torácica (Winker 2000), más recientemente conocido como compresión cardíaca rápida (Engilis Jr et al. 2018). ...
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Caldasia 44(1):154-164 | Enero-abril 2022 http://www.revistas.unal.edu.co/index.php/cal ECOLOGÍA Citación: González A, Jiménez A, García-Lau I, Mugica L, Acosta M. 2022. Ecología trófica de Calidris minutilla, Calidris pusilla y Calidris mauri (Aves: Scolopacidae) en dos humedales naturales de Cuba. Caldasia 44(1):154-164. Ecología trófica de Calidris minutilla, Calidris pusilla y Calidris mauri (Aves: Scolopacidae) en dos humedales naturales de Cuba Trophic ecology of Calidris minutilla, Calidris pusilla, and Calidris mauri (Aves: Scolopacidae) in two natural wetlands of Cuba ABSTRACT Shorebirds are noted for their long migrations and their dependence on a wide variety of wetlands annually. Their distribution in foraging habitats is related to the availability of prey. In this work, we characterized the diet of three species of the genus Calidris, C. minutilla, C. pusilla, and C. mauri in the wetlands of Río Máximo (RM) and Tunas of Zaza (TZ) and studied their relationship with prey availability. The diet was characterized by an analysis of stomach contents of individuals collected after peak feeding times. Prey availability was evaluated by taking 20 sediment samples with a 50 CC syringe that was buried at a depth of 10 mm. Subsequently, samples were filtered with a 500-micron sieve to facilitate identification. Diet analysis confirmed a high use of Diptera larvae and pupae by all three species in RM. In TZ, Diptera was dominant in the diet of C. minutilla, but the other species made a more balanced use of other items. The similarity between the resources available in the sediment and those found in the diet was less in TZ than in RM. Tanaidaceae (Malacostraca) were the most abundant item in the TZ sediment; however, it was not identified in any stomach. In general, in RM the prey was consumed according to their availability, while in TZ there was more selectivity in the use of the food.
... Acid-free tissue paper should be placed between the specimens to reduce mechanical stress on the feathers. Individual feathers or wings are best stored in acid-free paper packets, which in turn are stored in acid-free cardboard boxes (Winker 2000). ...
Chapter
2.9.1 Microslides Microslides are relatively easy to store and may be kept at room temperature under normal conditions. Slides should be handled gently, kept in the dark and at cool temperatures, and stored on stable shelving, especially in the case of large and heavy slide collections (Carter and Walker 1999). Permanent mounts are best prepared with resin-based products, such as Canada balsam and Euparal for botanical and mycological specimens, (Carter and Walker 1999) and epoxy resins for fossils. As slide mounts will always remain fluid to some extent, they should be stored flat with the mount on the upper side. Slotted slide boxes, for instance, should be stored upright like books on a shelf and marked in a way that indicates how to open the box correctly, to prevent slides from falling out and breaking. Microslides may be stored together with the main collection or separately with other microslides. Fossil microslides In fossils, the size of the microslide should correspond to the nature of the fossil, whether it be a longitudinal or tangential cut or a cross section, a small or large fossil, or a cross-drilling sample from a large fossil (e.g. Chinsamy and Raath 1992, Stein and Sander 2009). Commonly used microscope slides for fossils, also referred to as palaeohistological slides, come in different sizes: common formats are 2.5 × 5 cm, 5 × 5 cm, 7.5 x 5 cm. Unusual formats may include anything up to self-cut window panes measuring tens of centimetres in length and width. Palaeohistological slides are usually thicker than standard microscopy slides used in biology labs. They can be up to several millimetres thick, partly to withstand strains on the slide that might occur during the preparation of the fossilised samples. Due to the wide range of sizes and thicknesses, palaeohistological slides are stored in a variety of different slide boxes. Labels for microslides Care must be taken with slide labels. Some commercially available slide labels are unsuitable for permanent preparations because of the poor quality of the paper or glue. If a self-adhesive label is used, ensure that it is of archival quality (with acid-free adhesives) (Carter and Walker 1999). Recommendations à standardise storage types if possible à use resin-based mounting for long-term storage, i.e. Canada balsam and Euparal in botanical and mycological preparations, and epoxy resins infossil preparations Examples to see one of the largest collections of palaeohistological slides in Switzerland, visit the PIM in Zürich to see preserved microscopic algae (diatoms) on microslides and stored in histologic preparation boxes, see the MHNG in Geneva for taxonomic studies, the type specimens represented in the cryptogamic collection of the CJBG in Geneva have been removed from the general collection and stored in separate fire-proof metal cabinets (Clerc et al. 2017) Supplier storage boxes can be bought from a broad range of histology lab suppliers like Carl Roth Laborbedarf (Arlesheim CH, www.carlroth.com/ch) Further reading for an extensive overview of storing, managing, and digitizing slide collections see Neuhaus et al. (2017)
... Acid-free tissue paper should be placed between the specimens to reduce mechanical stress on the feathers. Individual feathers or wings are best stored in acid-free paper packets, which in turn are stored in acid-free cardboard boxes (Winker 2000). ...
Chapter
Fossils can be completely petrified, include original material or organic matter, contain unstable mineral parts or belong to the so-called subfossils consisting mostly of original material. The latter often do not differ significantly from zoological objects and can usually be stored in the same way (see section 2.7). Completely petrified fossils are the least problematic and may be stored relatively easily under standard conditions, like simple hard rock specimens and most other stable minerals. Fossils with unstable components may require special rooms or storage conditions. Small, fragile fossils may need to be stored in closed containers, boxes, or glass vials to prevent specimens and labels from getting lost. Even in smaller fossils, the application of accession numbers directly onto the specimen can prevent misplacement if several containers or storage boxes are opened for comparative studies or for inventory purposes (see figure 2.8.4.a). Pyritised fossils are unstable Pyritised fossils, containing the iron sulphides pyrite or marcasite, are unstable and prone to decay under normal atmospheric conditions, a process known as ‘pyrite-mar - casite destabilisation’ or ‘pyrite disease’ (Larkin 2011). The combination of high relative humidity and atmos - pheric oxygen causes a reaction producing sulphuric acid that attacks affected specimens and which may also affect nearby drawers, labels, boxes and other neighbouring fossils. If stored in glass, affected specimens can expand and shatter their containers. Decay can be prevented if specimens are stored in an oxygen-free environment, i.e. in an inert gas compartment. For larger specimens or whole collections, however, this may not be feasible, given the costs associated with airtight storage cases or other such storage options. According to Larkin (2011), the neutralisation of sulphuric acid may be achieved through treatment with ammonium gas or ethanolamine thioglycolate. Important prevention measures include the identification and isolation of potentially affected pyritised fossils, a rel- ative humidity between 30 – 45% if possible but certainly below 60% and regular collection checks to detect the beginnings of pyrite decay, such as the presence of greyish-yellow dust smelling of sulphur. Oil shale fossils can easily fall apart Oil shale fossils can fragment if the mother rock is drying out. A short-term transfer into distilled water can save the rock from dehydration. If stored for a longer time in water (not recommended but potentially necessary in certain cases), an additive should be used to prevent the growth of mould. For mid- to long-term rescue, specimens can be stored in glycerine or permanently transferred to synthet- ic resins. In the latter technique, known as the ‘transfer method’, the synthetic resin becomes the new support for the fossil and the original, fragile sediment is removed. To perform this transfer, different 2-component epoxy resins are available, some of which are also transparent, such as Araldite, Biresin and Bakelite/Epikote. Embedding the fossil is a permanent measure. The application of transparent or non-transparent resin should therefore be care - fully considered prior to the start of preparation, and depends on the following questions: which side should be visible at the end? Should the backside of the fossil be still visible through the resin? Shall the specimen be dis- played in the future? Will indirect lighting of the fossil through the resin be used?
... Twelve birds (6 of each sex) of the wild Turkey Meleagris gallopavo were collected to perform this study. The collected birds were anesthetized with ether, then transferred to the anatomical lab and put on the anatomical plate to open its abdominal region from the middle with accurate scissors according to (WINKER, 2000). The collected samples must be without any alimentary tract abnormalities. ...
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Introduction: the present study aimed to identify the histological and histochemistry characterizations of the Jejunum in the wild Turkey Meleagris gallopavo microscopically by using several histological and histochemical stains. The results showed that the jejunum wall is composed of the four layers that make up the wall of the digestive tract. The mucosal layer had a numerous long villus that appeared in various forms. Simple columnar epithelial tissue covers the villi, and this tissue is also rich in goblet cells. The secretory units of Liebrkuhkn’s crypts had a spherical shape, some of which contained entroendocrine cells. The Jejunum also was containing Payer’s patches. Auerbach’s plexus also appeared in the muscularis layer. The histochemical results of the epithelial tissue and intestinal glands showed a positive response to AB pH 1, pH 2.5, and PAS techniques, which indicated the secretion of mucous substances. In contrast, the response was negative for BP and SB techniques, which suggests the non-secretion of protein and fatty substances. The study concluded that the jejunum structure is similar to the composition of the rest of the small intestine. Still, it characterized by its multiform villi, entroendocrin
... Acid-free tissue paper should be placed between the specimens to reduce mechanical stress on the feathers. Individual feathers or wings are best stored in acid-free paper packets, which in turn are stored in acid-free cardboard boxes (Winker 2000). ...
... Acid-free tissue paper should be placed between the specimens to reduce mechanical stress on the feathers. Individual feathers or wings are best stored in acid-free paper packets, which in turn are stored in acid-free cardboard boxes (Winker 2000). ...
... Scientific collecting remains an important tool for researchers studying geographic variation, specieslevel classification, anatomy and morphology, molt and plumage sequences, subspecies or population limits, vouchering geographical records, and biodiversity inventories (Winker et al. 1991;Remsen 1995;Winker 2000;McGuire et al. 2009;Howell 2010;Rocha et al. 2014;Pyle et al. 2015;Clark and Rankin 2020;Puga-Caballero et al. 2020). As mentioned previously, collection of any hummingbird is regulated through state and federal permitting processes, which requires specific scientific collecting permission from the USFWS Migratory Bird Permit Office in the region where the investigator lives. ...
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Research on hummingbirds over the decades has provided insights into their evolution, migration, physiology, and numerous other areas, including conservation biology. Their small size, energy demands, and high metabolic rates are some of the challenges researchers face when obtaining research samples and biologic materials from live hummingbirds. This manuscript summarizes the established literature dealing with basic methods that scientists have used when capturing, handling, and otherwise researching hummingbirds. Based on the authors’ experience, best practices for working with live hummingbirds are presented, including permitting requirements for studying live hummingbirds, trapping and marking, handling techniques, safe collection of tissue samples, first-aid measures, and euthanasia of hummingbirds, as well as processing of hummingbird specimens (e.g., necropsy and preservation).
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