Collection, Preparation, and Fixation of Specimens and Tissues
Nancy B. Simmons and Robert S. Voss
Department of Mammalogy, Division of Vertebrate Zoology,
American Museum of Natural History, New York, NY 10024
Email: simmons@amnh.org, voss@amnh.org
Citation: Simmons, N. B., and R. S. Voss. In press. Collection, preparation, and fixation
of specimens and tissues. In: Ecological and behavioral methods for the study of bats,
2nd Edition (T. H. Kunz and S. Parsons, eds.). Johns Hopkins University Press.
Introduction
In some parts of the developed world, researchers have access to reliable
published handbooks for taxonomic identification of captured bats. Good field guides
contain accurate illustrations, diagnostic external measurements, and authoritative
descriptions of key characters that often allow scientific names to be associated with
ecological, physiological, or behavioral data with a degree of certainty that is acceptable
for some research purposes. In many undeveloped regions, however, no reliable field
guides are available; in such situations, unvouchered identifications amount to more-orless educated guesswork. Especially in the tropics, local bat faunas include numerous
closely related and confusingly similar species, and new species are still being described
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at a surprisingly steady rate (Simmons, 2003). Even in western Europe and temperate
North America, however, thoughtful researchers working with state-of-the art field
guides may have reason to doubt the reliability of their identifications when
morphologically cryptic species are known to occur in sympatry (e.g., Pipistrellus
pipistrellus and P.s pygmaeus; Myotis yumanensis and M. lucifugus).
In the absence of unambiguously diagnostic field criteria, the best way to
document identifications of captured bats is by collecting and preserving voucher
specimens. The purpose of this chapter is to explain how to do so legally and in such a
way as to maximize the usefulness of preserved specimens as a source of data, not only
for the individual scientist who collects them, but also for the larger community of bat
researchers who may never have access to the same field opportunities. Properly
prepared and preserved in museums, scientific specimens are a permanent source of
crucial information for a wide range of biological research topics. Learning how to
collect and prepare voucher material takes time and effort, but it is an essential aspect of
field research that bat biologists ignore at their peril.
Like any other aspect of field research, collecting specimens requires advance
planning. Special materials are needed to optimally preserve and label specimens, and
certain skills (e.g., preparing study skins) are best learned and practiced well in advance
of the moment when they are needed. Also, it is advisable to arrange for a permanent
specimen repository before leaving for the field. All of these goals can be accomplished
by contacting a natural history museum that maintains a large research collection of bats
and is interested in acquiring new material from the vicinity of your field project. In
return for specimens, most research museums are willing to help train researchers in basic
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aspects of specimen preservation, and to provide crucial supplies (e.g., labels, archivalquality notebook paper) and advice (e.g., about negotiating import/export problems).
Because collected specimens may represent new species, important range extensions, or
other taxonomically significant discoveries, they can often lead to productive
collaborations (and publications) with museum scientists.
Permits and Regulations
Collecting Permits and Import/Export Permits
All countries have laws regulating the collection and export of biological
specimens. Even if such laws are not locally enforced, proper documentation that
specimens were legally obtained must accompany United States Fish and Wildlife
Service (USFWS) import declarations, and most museums will not accept specimens that
lack appropriate paperwork. Strict compliance with in-country regulations is therefore
necessary for scientific collecting.
Advance preparation is almost always needed to avoid costly delays in obtaining
collecting permits. Colleagues at in-country institutions (universities, museums, and
conservation NGOs) are certain to be familiar with application procedures, some or all of
which can often be initiated months in advance of the fieldwork for which they are
needed. Patience, goodwill, humor, and cultural sensitivity are usually helpful in
negotiating the sometimes-complicated bureaucracy that controls the permit-granting
process in many countries. Because collecting permits are a prerequisite for subsequent
applications to obtain export permits, and because both kinds of permits are usually
issued by the same office, no effort should be spared to make a good impression at the
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outset.
Careful inquiries should be made to determine whether only wildlife export
permits are required to take biological materials out of the country. In some countries,
airports require that biological specimens be certified as disease-free or “sanitary,” a
determination that is often made by different agencies (e.g., the agriculture ministry)
from those that deal with wildlife matters. Be prepared to deal with such complexities,
which can be difficult (or expensive) to resolve if encountered at the last minute.
Whatever physical method the researcher chooses to export specimens from the
country of origin (e.g., as personal luggage, through the mail, or by a commercial courier
service), they must be accompanied by appropriate documentation to be imported legally
to the United States and most other developed countries. For importation to the United
States, such paperwork must include USFWS form 3-177, otherwise known as the
“Declaration for Importation or Exportation of Fish or Wildlife” (available with
instructions at http://www.le.fws.gov/faqs.htm). Technically, this document can be filed
within 90 days after the specimens enter the country, but it is prudent for passengers
arriving with specimens in their luggage to have form 3-177 filled out on arrival. At
most airports no USFWS inspectors are usually on hand, so decisions about whether to
impound or release specimens are made by U.S. Customs Service personnel. Handing
over a filled-out copy of 3-177 with export permits attached may persuade otherwise
skeptical customs officials (“Preserved bats?”) that you know what you are doing.
However, be sure to retain multiple identical copies of all export and import documents,
which will be needed when the specimens are accessioned into a museum.
Obtain copies of form 3-177 before you leave the U.S. and familiarize yourself
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with the information that you will be required to provide on it. Among other items, the
USFWS requires “common” (English) names for each imported species. English
common names for most currently recognized species of bats can be found in Wilson and
Cole (2000) and Simmons (2005). Common sense will suggest how to fill out the form if
species identifications are in question. Obviously, identifications may need to be revised
in the light of subsequent museum study, but bear in mind that USFWS inspectors are
trained in law enforcement, not biology, and may be unfamiliar with the usual taxonomic
equivocations (“Artibeus sp.,” “unidentified phyllostomid,” etc.).
Exporting and importing specimens of endangered species require special permits.
The Convention on International Trade in Endangered Species (CITES) is based on lists
(“appendices”) of protected species belonging to several categories (see
http://www.cites.org/). Appendix I species are considered to be threatened with
extinction; Appendix II species are not necessarily considered to be at risk of extinction,
but international trade in specimens is controlled; and Appendix III species are protected
in certain countries. To date, only two species of Pteropus and seven species of
Acerodon are listed in Appendix I, but the remaining species of both genera are listed in
Appendix II. Appendix III contains only Platyrrhinus lineatus, which is protected in
Uruguay. Researchers who wish to collect voucher material of these taxa (including the
remains of animals eaten as food) should consult the CITES-licensing authorities of the
country of origin for special permits to do so.
In general, specimens can be imported to the United States only through
designated ports of entry. As of May 2005, the USFWS Office of Law Enforcement
website (www.fws.gov/le/Designated_Ports) lists 18 designated ports of entry including
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Anchorage, AK; Atlanta, GA; Baltimore, MD; Boston, MA; Chicago, IL; Dallas/Ft.
Worth, TX; Houston, TX; Honolulu, HI; Los Angeles, CA; Louisville, KY; Memphis,
TN; Miami, FL; Newark, NJ; New Orleans, LA; New York, NY; Portland, OR; San
Francisco, CA; and Seattle, WA. Researchers planning to enter the U.S. through any
other port should apply in advance for a Designated Port Exemption Permit (instructions
provided at www.fws.gov/le/DesignatedPort Exemption Permit).
DOT and IATA regulations
Investigators planning to ship specimens (vouchers or tissue samples) by air
should be prepared to comply with regulations of the United States Department of
Transportation (DOT) and International Air Transport Association (IATA). Ethanol (=
ethyl alcohol), isopropanol (= isopropyl alcohol), and formalin (formaldehyde solution)
are considered hazardous materials (DOT) or dangerous goods (IATA), and must be
handled and packed accordingly. There are strict limits on the amount of chemicals that
may be shipped, and specifications regarding packing material and labeling (IATA
regulations may require special labels that must be obtained in advance of shipping).
Because regulations change regularly, it is wise to consult both your carrier and IATA
directly before leaving for the field. For information on IATA regulations, see
http://www.iata.org/whatwedo/dangerous_goods; for DOT regulations, see
http://hazmat.dot.gov/.
Data
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Field Notes
Biological specimens without data are virtually worthless, so it is important that
fieldworkers record accurate information about where and when specimens are collected.
Such information becomes more valuable as the years go by and habitats are transformed
or destroyed by human activities. When more than a few specimens are to be collected in
the course of a field season or over an entire research career, it is useful to keep two
permanent (archival-quality) field records, a journal and a catalog. Both journal and
catalog should be written in waterproof carbon-based (“India”) ink on acid-free paper in a
standardized format. If carbon-based ink is not available, use a pencil (ball-point pen ink
is soluble in alcohol and should not be used for archival records). We prefer smallformat (ca. 16 × 22 cm) loose-leaf paper because it can be carried easily in a compact
ring binder for use in the field; later, the loose pages can be bound and archived at the
institution where specimens are finally deposited.
The journal is a continuous record of field activities, much like a diary, and
should include all noteworthy information that provides temporal, geographic, and
ecological context for collected specimens. Every page of the journal should record the
following items (see Figures 1 and 2): (1) the last name of the investigator, typically
written in block capitals in the upper right-hand corner; (2) the year, typically written
under the investigator’s name; (3) the locality, typically underscored with a long squiggle
on the top line; and (4) the day and month, typically in the left-hand margin. These items
are essential to preserve the correct sequence of pages if they are mixed up at any time.
Nothing should be written within about an inch of the left-hand edge of the paper to allow
for future binding
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Figure 1. Sample journal page from 1998 field notes by Voss, illustrating the format
and information typically recorded for an array of mist nets and a night of netting. It is
helpful to include illustrations of net placements if they are not linear. Complete lists of
species captured are an important supplement to catalog entries, which include more data
but take note only of specimens preserved as vouchers.
Figure 2. Sample journal page illustrating the format and data typically recorded for
roost surveys and collecting. To avoid confusion, we always number roosts
consecutively at any given locality, and include illustrations whenever a new or unusual
roost type is found. Specimens collected at each roost are cross-referenced with the
catalog. Individuals observed but not captured or collected should be described in journal
roost accounts.
Researchers should write in their journals every day, synthesizing a coherent
narrative of their activities and describing their collecting efforts. It is unwise to wait
several days to record journal entries because the days blur together and errors of
memory can cause serious confusion for others who may consult these documents in the
future. Because the journal is intended for other scientists to read, entries should be
written legibly, unusual abbreviations should be avoided, and sketches or maps should be
included to illustrate what is meant. The standard journal entry for a night of bat-netting
or -trapping should include: (1) where and in what habitat the nets or traps were placed;
(2) how many nets or traps were used; (3) the type and length of nets or kinds of traps
used; (4) the height(s) of the nets or traps; (5) the times when the nets or traps were
opened and when they were closed; and (6) the identification, sex, capture time, and
capture height (if necessary) of all captured bats. Searching for roosts is an important
supplementary method for bat inventory fieldwork because many species that are hard to
net or trap can easily be found in their diurnal refugia (Voss and Emmons, 1996). In
order to keep track of roosts found opportunistically, or as a result of deliberate surveys,
it is useful to number them and provide certain essential data: (1) nature of the roost (e.g.,
leaf-tent, exfoliating bark, cave, culvert, hollow tree); (2) where it was found; (3) what
species of bats occupied it; (4) the field catalog numbers of any specimens preserved as
vouchers; and (5) any other relevant observations (e.g., about the sexes, ages, or behavior
of roost residents). Examples of journal pages describing bat survey and inventory
methods are provided in Figures 1 and 2.
In contrast to the journal, the catalog is a of list individual animals prepared as
voucher specimens. Like the journal, each page should have the last name of the
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investigator, the year, and the locality written as headers. Catalog entries consist of a
standardized set of data recorded for each specimen. We record data for each specimen
on four sequential lines (see examples in Figures 3 and 4):
[first line]: field number, sex, and preparation type.
[second line]: identification and date.
[third line]: measurements and reproductive information.
[fourth and subsequent lines]: circumstances of capture and other notes.
Several simple conventions have been followed by many generations of mammalogical
collectors and should be maintained to facilitate clear transmission of essential
information:
The field number consists of the capitalized initials of the investigator (e.g., NBS)
followed by a unique whole (integer) number, one for each specimen; this numerical
series should begin with “1” for the first scientific specimen prepared by the investigator
and continue without interruption or duplication to the last he (she) will prepare at the
end of his (her) career. Other systems, regardless of intent, may lead to confusion; if you
have used another system in the past, start this system now and use it consistently
hereafter. The same series of numbers can be used for all collected material, whether
bugs, birds, frogs, plants, or mammals.
The preparation type describes the way in which the specimen is preserved.
Standard options include “skin and skull” (dry skin with skull removed for cleaning;
remaining carcass preserved in fluid); “fluid” (entire animal fixed in formalin and stored
in alcohol); “skeleton” (specimen skinned and gutted, skeleton retained for later
cleaning); “tissue” (tissue sample prepared). These may be combined as appropriate,
e.g., “fluid + tissue”, etc.
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Figure 3. Sample catalog page showing format and data recorded for bats captured in
mist nets (see text for explanation of format and abbreviations). Scientific names should
always be written in pencil to allow for corrections. All of the data to the right of the
vertical line were recorded in the field; the numbers to the left of the line are permanent
museum numbers which were added after the specimens were accessioned into museum
collections (in this case, the American Museum of Natural History [AMNH] and Museo
de Historia Natural del la Universidad Nacional Mayor de San Marcos in Lima, Peru
[MUSM]).
Figure 4. Sample catalog page showing format and data recorded for bats collected at
roosts. See text for discussion.
The identification should be the scientific name of the animal written in binomial
form. If the animal cannot be identified to species, the genus or other higher-level taxon
should be recorded (e.g., “Myotis sp.”, “Molossidae indet.”). Questionable identifications
can be indicated with a question mark (e.g., Artibeus ?jamaicensis). Identifications
should always be recorded in pencil to allow for subsequent corrections.
The date is the day the animal was collected, or the day following the night it was
collected (by our convention, bats caught at 9 pm on 6 October and at 3 am on the 7th,
will both have 7 October as the date of collection). Bats captured on the same night
should have the same collection date even if some are kept alive in camp for several
additional days before preparation. The collection date is important because researchers
may later be interested in knowing what other species were taken in the same net or trap
on the same night; the date when the specimen was prepared is unimportant.
The measurements (always in millimeters and grams) are recorded in a standard
sequence that (for bats, see below) consists of Total Length, Tail Length, Hind Foot
Length, Ear Length, and Forearm Length. Measurements in this sequence are always
separated by ×s, and the series is concluded by an equal sign (=) and the weight (mass).
Missing values (for whatever reason) should be indicated by a question mark in square
brackets. Thus, a small phyllostomid with both forearms broken (and therefore
unmeasurable) could have measurements recorded as 55 × 7 × 9 × 19 × [?] = 42 g.
Some investigators also measure the tragus, and use the notation “tr” to indicate
this value. It may be placed either in parentheses after ear length or noted separately
after the weight.
For most specimens, the circumstances of capture and other notes should include
(1) the habitat in which the animal was collected; (2) height above the ground (measured
or estimated); (3) the time (if known) when the animal was captured; (4) the method of
collection (trap type or net size and denier); and any other relevant data (e.g., roost
number, field numbers of other individuals captured in the same roost).
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Locality data
Although Global Positioning System (GPS) technology has revolutionized
geographic data collection, it is still important to record other locality data that help place
satellite triangulations in context. Minimally, locality records in both the journal and
catalog should include the country of origin, the next-highest administrative unit (state,
department, province, etc.), and the particular locality where material was collected.
These data are traditionally recorded in sequence, with the country name in capital letters
followed by a colon. Other punctuation conventions are optional but should be followed
consistently to avoid confusion. For example, specimens collected at the Matses Indian
village of Nuevo San Juan, which is on the Gálvez River in the Peruvian department of
Loreto could be recorded as from “PERU: Loreto, Río Gálvez, Nuevo San Juan
(73˚9'50"W, 5˚14'50"S), ca. 150 m”. Obviously, much more detailed geographic
information can be recorded in the journal (e.g., descriptions of where nets were actually
placed along trails leading from the village of Nuevo San Juan), but more succinct
formats are appropriate for the catalog and specimen tags.
Other useful locality data include distances by road (or trail) from well-known
landmarks, and whether specimens were collected on the right or left bank of a major
river (by convention, the observer is assumed to be facing downstream). These might
seem like superfluous details if GPS data are available, but many published maps lack
accurate coordinate grids, so museum researchers may be unable to correctly interpret the
geographic significance of collected specimens unaccompanied by traditional spatial
descriptors. It is also important that the journal contain an adequate description of local
habitats, especially if the landscape is a mosaic of two or more vegetation formations
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with significantly different faunas (e.g., gallery forest versus savanna, primary versus
secondary forest, etc.). Hand-drawn maps are frequently helpful for showing where
specimens were collected in relation to local landscape features.
Standard measurements for bats
As noted above, standard field measurements for bats include five linear external
measurements plus body weight. These should be recorded as soon as possible after
death to prevent inaccuracies caused by shrinkage and/or dehydration. Linear
measurements (always in millimeters) are best determined by using a transparent plastic
ruler trimmed so that one end corresponds exactly to the zero mark. The standard
methodology is as follows (also see Handley, 1988).
Total length: This is either the distance between the tip of the snout and the tip of
the last caudal vertebra (for bats with tails), or the distance between the tip of the snout
and the back of the pelvis (for tailless bats). Most bats should be measured by placing
them belly-down on a ruler with the snout at zero. Press down gently on the head and
back to flatten the animal against the rule, and grasp the terminal tail vertebrae with
forceps. Taking care not to displace the snout from the zero mark, gently but fully extend
the tail to read the total length of the animal. If the bat is pregnant and the fetus large
enough to distort the spine when the bat is on its belly, place the animal on its back to
measure total length.
Bats that lack an external tail (e.g., many phyllostomids) require a different
method to get an accurate measurement. The bat should be placed belly-down on a ruler,
and each leg folded anterolaterally so that both are positioned over the animal’s back.
12
Press down gently on the back and legs to fully extend the body; this will cause the ischia
of the pelvis to protrude as the posteriormost parts of the skeleton. Read the total length
from the tip of the snout to the back end of the pelvis.
Because the vertebral column is flexible, measurements of total length are
notoriously inaccurate and should be repeated several times for each individual to obtain
a consistent measurement. Total length should be recorded to the nearest whole
millimeter.
Tail length: This is the combined length of the caudal vertebral series from its
origin on the pelvis to the tip of the tail. Tail length is best measured by placing the bat
on its belly and holding the tail vertically with forceps so that the tail makes a right angle
with the pelvis. Place the trimmed (zero) end of the ruler at the base of the tail (i.e.,
where it joins the sacrum), gently extend the tail until it is straight, and read its length at
the tip. Do not hold the tail by the terminal vertebra, as doing so may make it hard to see
the tail tip properly and sometimes result in vertebral disarticulation. Like total length,
tail length should be recorded to the nearest whole millimeter.
Hind foot length: This is the distance from the anterior edge of the base of the
calcar (or the end of the calcaneum in bats that lack a calcar) to the tip of the claw of the
longest toe. Hind foot length is measured by flattening the foot on a ruler with the
anterior edge of the calcar at zero; the length is then read at the claw-tip of the longest toe
when all of the digits are fully extended. Like the previous two measurements, hind foot
length should be recorded to the nearest whole millimeter.
Ear length: This is the distance between the notch at the base of the ear and the
tip of the pinna. Place the zero end of the rule in the notch (posteroventral to the tragus)
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and gently extend the pinna along the ruler to its greatest extent; read the length at the
extreme margin of the pinna, being careful not to stretch the tissue while doing so.
Unlike the previous three measurements, ear length should be recorded to the nearest
half-millimeter.
Forearm length: This is the distance from the elbow (the tip of the ulnar
olecranon process) to the wrist. Forearm length is always measured with the wing folded.
Place the elbow on the tabletop (or a similar hard surface) and hold the forearm
vertically; placing the ruler with the zero mark at the elbow, read the length of the
forearm at the wrist. By convention, this measurement includes those carpals that are
exposed at the distal end of the forearm when the wing is folded. Like ear length,
forearm length should be recorded to the nearest half-millimeter when a ruler is used. If
calipers are available, however, forearm length can be accurately (repeatably) determined
and recorded to the nearest tenth of a millimeter.
Some investigators believe that forearm measurements made on fresh specimens
are not comparable with measurements made on dry specimens (e.g., skins). However, in
our experience the difference between such measurements is minimal if care is taken to
fully fold the wing of a fresh specimen before measuring the forearm.
Weight: This is the weight (mass) of the entire (unskinned) bat, recorded in
grams. Small spring-scales in a variety of sizes (e.g., 10 g, 30 g, 50 g, and 100 g
capacities; manufactured by Pesola and other ornithological suppliers) are ideal for
determining weights. The smallest-capacity scale appropriate for the size of the animal
should be used in order to insure accurate data; therefore, it is usually necessary to have
several scales of varying capacity at hand. Weights should be recorded to the smallest
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increment marked on the scale (usually to the nearest gram for large bats, to the nearest
tenth of a gram for small bats).
Some researchers take other measurements in the field (e.g., wingspread, length
of tragus; see Handley, 1988). However, we have found these to be of limited systematic
value. Additional measurements of taxonomic significance (e.g., the lengths of
metacarpals and phalanges) can usually be measured accurately on preserved specimens,
so we do not include them in our list of standard field measurements for bats.
Sex, reproductive status, and age
The determination of sex and reproductive status is straightforward for most bats
(see Racey, 1988, this volume). Males of most species have a naked, pendulous penis
that is obvious even in juveniles, and females have urogenital and rectal openings that are
separated by an area of smooth skin that may or may not be furred. Both males and
females of some taxa have well-developed mammae, so it is essential to examine
genitalia to determine the sex of an animal.
Members of two families, Emballonuridae and Noctilionidae, may present
difficulties in sex determination. The genitalia of both sexes of emballonurids are coneshaped in external appearance: males have a non-pendulous penis that is hidden within a
furred, conical fold of skin, and females have enlarged labia that are often furred and may
superficially resemble the male genitalia. In these bats, it is therefore necessary to
palpate the genitalia to determine if the conical structure consists of paired labia (as in
females) or has a small terminal opening (as in males). The situation in noctilionids is
somewhat different: males have a pendulous penis like most other bats, but the female
15
labia are sometimes elongated and may superficially resemble a penis. Again, palpation
of the external genitalia is sufficient to unambiguously determine the sex of an individual.
In addition to sex, it is useful to record other reproductive data. In male bats, the
presence or absence of external testes should be noted. If it is of interest from a research
perspective, testis length and width may also be recorded. Check females for pregnancy
by palpation, but be aware that early pregnancies may not be detected by this technique.
A more accurate field method is to open the abdomen and examine the reproductive tract,
but the time required for this (and potential damage to the specimen) may not be justified
unless reproductive data are important for the research goals of the fieldwork. If
dissections are done in the field, record the number and crown-rump length of each
embryo or fetus; this can often be done without removing them from the uterus. If a fetus
is completely removed from the mother, it should be given its own field number, and
field numbers for both fetus and mother should be cross-referenced in the field catalog.
Sex, crown-rump length, forearm length, and weight are routinely recorded for each
individually cataloged fetus.
In addition to checking for pregnancy, it is also important to note whether a
female is lactating. This can be done by examining the mammae (see Racey, 1988, this
volume; Hood et al., this volume). Tiny nipples surrounded by fur are common in
nuliparous females. If the nipples are large, keratinized, and surrounded by hairless
areolae, it is likely that the female is either lactating or has lactated in the recent past. In
lactating individuals, the white mammary glands can often be seen under the skin of the
chest and armpit. Gently palpate the glands and squeeze the nipple gently; if a drop of
milk can be ejected, the animal was lactating at the time of death. After lactation the
16
nipples generally retain their enlarged keratinized appearance, which is indicative of
parous females in most species of bats (Racey, 1988, this volume; Hood et al., this
volume).
Assessing the age of captured individuals is difficult under most field conditions
(Anthony, 1988; Rossinni and Wilkinson, this volume). Rather than introducing error by
trying to be too specific, we recommend using the following broad categories:
Fetus: This category is reserved for unborn young removed from the mother
during specimen preparation. As noted above, each fetus should receive its own unique
field number, which should be cross-referenced with that of the mother in the catalog.
Juvenile: We use this category for nursing offspring captured with their mothers
or in maternity roosts. Juvenile bats are young individuals that are smaller than
conspecific adults and are usually nonvolant. They frequently retain the deciduous
dentition (tiny hooklike teeth), have a juvenile pelage (often grayer and fuzzier than the
adult fur), and the epiphyses of their long bones are unfused (see below). Each juvenile
should receive its own unique field number, which should be cross-referenced with that
of the mother (if known) in the catalog.
Subadult: Subadult bats are volant individuals that are mature in size and
dentition but still have unfused epiphyses and may retain a juvenile pelage. Epiphyseal
fusion is most easily evaluated by examining the finger bones through the skin of the
wing. In adults, each metacarpal/phalangeal joint and interphalangeal joint appears
externally as a swollen knob that diverges sharply from the shafts of the participating
bones. In juvenile and subadult bats, these joints are characterized by more elongate
swellings that encompass cartilaginous epiphyseal (growth) plates can be seen on either
17
side of the actual point of articulation (Anthony, 1988). These plates are readily visible
when a bat wing is transilluminated because the cartilaginous areas appear lighter than
the more densely mineralized bone (Anthony, 1988). Anthony (1988: Figure 1) provided
excellent illustrations of internal and external morphology of wing joints that can be used
as a guide for identifying subadult bats.
Adult: Adult bats are characterized by fused epiphyses, and mature size,
dentition, and pelage. Clearly, some individuals may fall at the borderline of the
categories defined above. In such cases, use your judgment. As a general rule, the
distinction between adult and nonadult bats is the most important.
Labels and tags
All of the separate parts of morphological specimens must be labeled. Although
makeshift labels for haphazardly collected material can be improvised in the field, it is
important that they be durable and waterproof, and that the information recorded on them
be insoluble in formalin and alcohol. Traditionally, most museum collectors use
heavyweight archival-quality paper or cardboard labels on which data are recorded in
carbon-based (“India”) ink. Unfortunately, many inks traditionally used for this purpose
are no longer manufactured, and new ink formulas differ in their resistance to water and
other solvents. Therefore, it is prudent to test several ink samples for water- and alcoholresistance before purchasing a supply for the field. Labels of all kinds should be tied to
specimens with heavyweight buttonhole or crocheting thread, which should not be dyed.
All knots used to attach labels to specimens should be square knots that will not come
18
loose when specimens are packed together for shipment, so it is important to learn how to
tie these properly.
Skin labels: Although labeling conventions vary from museum to museum, skin
labels are typically longer than wide (about 80 × 20 × mm) and are perforated with two
holes at one end (Figure 5). A length of heavyweight (buttonhole) thread is passed
through both holes leaving a loop on one side; this loop is twisted once, both ends are
drawn through it, and the resulting hitch is drawn tight. A single overhand knot is tied
about 1 cm from the paper’s edge to allow the label to turn freely when tied to the
specimen. If possible, it is a good idea to thread a sufficient supply of labels in advance
to save time in the field. Data to record on skin tags include (1) locality, (2) date of
collection, (3) collector’s name, (4) field catalog number, (5) measurements and weight,
and (6) sex.
Skull/skeleton/fluid labels: These are generally smaller labels perforated by a
single hole through which a loop of heavyweight thread is passed, hitched over the free
ends, and knoted as are skin tags. Typically only the field catalog number is inked on
these labels, preferably on both sides. As noted above, it is important to test the
waterproof quality of the paper and the insolubility of the ink before leaving for the field
(where alternative materials may be unavailable). More than a few important specimens
have been lost because labels and/or ink did not perform as advertised.
Methods of Euthanasia
Field methods used to kill animals should be quick, as painless as possible, and
should not result in unnecessary damage to the specimen. Several humane methods for
19
euthanazing small mammals have been approved by the American Veterinary Medical
Association (1993) and by the American Society of Mammalogists Animal Care and Use
Committee (1998). These approved methods include the use of toxic inhalants, cervical
dislocation, and thoracic compression. Although lethal injection is another approved
method, it requires veterinary training and access to controlled substances (see American
Veterinary Medical Association [1993] for guidelines).
Inhalants acceptable for killing mammals include carbon dioxide, carbon
monoxide, halothane, enflurane, sevoflurane, methoxyflurane, desflurane, ether, and
chloroform (American Veterinary Medical Association, 1993; American Society of
Mammalogists Animal Care and Use Committee, 1998). However, some of these
substances pose risks to the investigator (e.g., chloroform is carcinogenic, ether is
explosively flammable), and others are impractical to use under field conditions (e.g.,
carbon dioxide). For some studies, however, the benefits of inhalants may be
considerable (e.g., because they kill ectoparasites), and outdoor use may minimize
associated risks (American Society of Mammalogists Animal Care and Use Committee,
1998). In general, inhalants should be used in well-ventilated areas by experienced
personnel wearing respirator masks (if appropriate). For detailed discussion of the
advantages, disadvantages, and proper use of various inhalants, see American Veterinary
Medical Association (1993).
Cervical dislocation is perhaps the quickest method for euthanizing small
mammals (American Veterinary Medical Association, 1993). For bats, cervical
dislocation is most easily accomplished by holding the animal with one forefinger across
its throat and the thumbnail of the same hand on the back of its neck (Handley, 1988).
20
With the other hand, the hind limbs are quickly pulled backward so that pressure from the
thumbnail causes separation of the cervical vertebrae. This is an effective method for
animals under about 60 g, but larger bats may require the use of pliers or wire-cutters
(with the jaws padded to avoid cutting the skin) to obtain a sufficiently firm grip on the
neck (Handley, 1988). Very large animals (over 200 g) can be killed by holding the head
in one hand and the hindquarters in the other hand, and rapidly hyperextending and
dorsally twisting the neck (American Veterinary Medical Association, 1993). The need
for technical competency is greatest when dealing with large animals because of their
stronger neck muscles (American Veterinary Medical Association, 1993).
Thoracic compression (= cardiopulmonary compression) is another means of
rapid and humane euthanasia (American Society of Mammalogists Animal Care and Use
Committee, 1998). For small bats (<50 g), thoracic compression can be accomplished
manually by quickly and forcefully compressing the chest between thumb and forefinger.
Compression should be forceful enough to expel all of the air from the lungs, and should
be maintained for at least two minutes after the heart stops beating. To euthanize larger
bats, we have successfully used large hemostats with a box-lock joint that can be engaged
to maintain sufficient pressure on the chest for several minutes. Suitable hemostats can
be purchased from medical and laboratory supply companies that deal in dissecting tools
and surgical instruments.
Storage of Specimens Prior to Preparation
Much of the scientific value of specimens---for both morphological and molecular
research---depends on timely preservation. Because tissue deterioration begins almost
21
immediately after death and may be very rapid in warm weather, it is generally advisable
to keep animals alive until just before they are to be prepared as specimens (for a
discussion of humane methods of restraint and transport, see Barnard, this volume; Kunz
et al., this volume). If specimens cannot be prepared immediately after death, they
should be labeled, stored in plastic bags (to reduce dehydration), and refrigerated or
frozen if possible. Freezing alters cell structure, however, so it should be avoided if the
specimen (or any part of it) is eventually intended for use in histological studies. If
refrigeration is unavailable, the specimen should be wrapped in a moist piece of cloth and
kept in the coolest place possible. Care should be taken at all times to protect specimens
from scavenging birds and mammals (chickens, cats, and dogs are common threats in
many field camps) and insects(ants, bees, and wasps), whose destructive potential should
never be underestimated in the tropics.
Choosing a Preparation Technique
Different preparation techniques result in specimens that are useful for different
purposes. The traditionally preferred “skin and skull” treatment---in which the skin is
stuffed and the skull preserved separately---has the advantage of preserving fur color and
pelage banding patterns, but it is laborious and results in irreversible damage to the
morphology of the face and ears (which frequently contain important taxonomic
information). Preservation as a “fluid” specimen – which consists of fixation of the
entire animal in formalin and subsequent storage in alcohol – is much less timeconsuming and preserves the morphology of the face and ears even if the skull is
removed. Fluid specimens are also the best preservation option if future dissection or
22
examination of internal organs is anticipated. However, fur color fades in fluid
preservatives, and it is difficult to determine pelage banding patterns on wet specimens.
Full skeletal preparations typically sacrifice the entire pelage and most internal organs
(although these can be saved in fluid), but has the advantage of allowing access to aspects
of skeletal morphology that are hidden in other preparation types. Obviously, it is
important to have a clear idea of the future use of the specimen before choosing a
preparation method.
We prefer preserving bats as fluid specimens under most circumstances. In our
experience, fluid preservation maximizes the usefulness of the specimen for future
systematic and anatomical studies while simultaneously being the least time-consuming
preparation method. Because skulls can be easily removed from fluid specimens after
preservation (leaving the face and ears relatively intact), this method does not preclude
examination of cleaned skulls. When only a single individual of a species is to be
prepared as a voucher, it should always be preserved as a fluid specimen unless it is
thought that pelage color or pattern may be critical for identification. On the other hand,
a dry skin should be prepared if the nature of the pelage does not agree with published
descriptions, or if it appears to differ significantly from that of other individuals of known
identity. When a series of specimens of the same species are to be collected at a single
locality, it may preferable to use a variety of different preparations techniques to
maximize usefulness of the series.
Preparing Fluid-Preserved Specimens
23
Preparation of fluid-preserved specimens consists of three steps after the animal
has been killed and measured: (1) preparation of the specimen for fixation, (2) fixation,
and (3) transfer to the storage medium. Transport from the field to the lab or museum
should be done only after fixation is complete.
Preparation and formalin fixation
After the animal has been killed, the first step in specimen preparation is to
measure it and record all relevant data in the field catalog. An appropriate fluid tag
should then be attached to the right leg of the animal. We prefer to tie the tag around the
leg midway between the ankle and the knee, inserting the tag string through the wing and
tail membranes with a large needle (like those sold for darning socks). The tag string
should project from the dorsal surface of the membranes, not the ventral surface. A short
length of string (ca. 1 cm) should be left between the tag and the animal to allow
manipulation of the tag without damage to the specimen. Never tie a tag around the
ankle of a bat, as this results in permanent distortion of the calcar and the membrane
attachments to the ankle and foot.
Once the animal has been tagged, open its mouth and insert a tightly-rolled ball of
cotton; this should be large enough to brace the mouth wide open so that the teeth and
other taxonomically informative oral structures will be visible for inspection after the
animal is fixed. Although it is much easier to insert the cotton ball before rigor mortis
sets in, the mouth can be opened after rigor is established if sufficient force is applied
(although care must be taken not to break any teeth). If cotton is not available, other
24
objects (e.g., a short stick) can be used to prop the mouth open, although these may be
hard to remove after fixation.
The next step is to position the bat in an appropriate posture for preservation.
This must be done before rigor sets in. The wings should be folded alongside the body
such that they do not overlap the abdomen or chin, the hind legs should be extended
posteriorly (away from the abdomen), and the neck should be extended anteriorly so that
the throat is exposed. It is often helpful to hook the claws of the hind feet over the edge
of a table, and stretch the body out belly-down on the tabletop to achieve this posture.
The value of preserving bats in an appropriate position cannot be overemphasized. In
addition to making it easier to get fluid-preserved specimens in and out of jars, proper
posture facilitates examination of taxonomically important aspects of the ventral pelage
and other external structures (e.g., nipples, genitalia, and throat glands) as well as
dissections of abdominal contents, thoracic musculature, and hyoid anatomy.
Proper fixation requires immersion in 10% buffered formalin (an aqueous solution
of formaldehyde gas; see Appendix 1). The length of time required for fixation depends
on a number of factors, including the size of the animal, whether or not the abdominal
cavity has been opened, whether or not the specimen has been injected, and whether or
not the fur has been sufficiently wetted prior to immersion. It also depends on the
volume of formalin and the number of specimens being fixed at the same time: as a
general rule, the volume of formalin should be at least three times the aggregate volume
of the specimens being fixed.
All bats should have the abdominal cavity opened prior to immersion in formalin.
The easiest and least destructive way to accomplish this is to use a pair of sharp scissors
25
to cut an incision in the ventral body wall along the posterior edge of the rib cage. Be
sure that the incision goes through both the skin and the muscular lining of the abdominal
cavity to allow formalin free access to the internal organs. For small bats, an incision of
about 1 cm should suffice; larger animals require larger incisions.
In addition to opening the abdominal cavity, it is also useful to inject formalin
into the thoracic cavity, throat region, and/or large muscle masses of the limbs if the
specimen is large (>100 g) or is intended for use in future dissections. Use glass syringes
with ground-glass plungers for injecting formalin (the rubber plunger gaskets of
disposable syringes may be attacked by formalin, rendering them useless after just a few
days of use). A syringe of 2 cc capacity fitted with a 22 gauge needle is ideal for small
bats; a 10 cc syringe fitted with an 18 gauge needle is more than adequate for most large
bats. Because needles are blunted with repeated use, carry a few spares or be prepared to
sharpen them on a whetstone. Use the same concentration of formalin for injection that
you plan to use subsequently for immersion. Take care to inject formalin slowly to avoid
overinflating body cavities and internal organs or causing formalin blisters to form under
the skin. It is usually better to make several small injections into a large muscle mass
than to inject the same total volume at a single site. Wear rubber gloves to protect your
hands from formalin throughout this procedure and those immediately following.
After the specimen has been prepared for immersion, the final step is to wet the
fur using water and a little dish soap. Fixation occurs more quickly and evenly if the fur
is wet because patches of unwetted fur prevent formalin from contacting the skin. After
using soapy water to wet the fur, rinse it thoroughly and then gently press out the excess
moisture before immersing the animal in preservative.
26
It is important to leave the specimen immersed in formalin long enough to
preserve all of its soft tissues. However, leaving a specimen in formalin too long may
harden muscles to the point that any manipulation of the limbs is impossible, and leaving
specimens in unbuffered formalin also tends to decalcify bones and teeth. . Most small
bats (<100 g, with the abdominal cavity opened as described above) are usually
sufficiently fixed after three or four days in formalin, but larger bats can take a week or
more. If batches of specimens are being prepared on a daily basis, it is wise to label a
series of jars with the dates when you expect specimens to be fully fixed, and to check the
state of fixation daily as specimens are nearing completion of the process.
To determine if a fluid specimen is adequately fixed, squeeze the abdomen and
larger muscle masses; they should feel firm to the touch (like well-done steak). If
specimens are overcrowded in the preservative, or if the formalin has been used
repeatedly to the point where it is significantly diluted, fixation may take longer than
expected. Because specimens can decompose in over-diluted formalin, it is best to
transfer problematic material to a freshly mixed batch of formalin. When fixation is
complete, remove the specimens from the preservative and store them for transportation
as described below.
Alternative field preservatives
Although fixation is the recommended first stage of fluid preservation, it can be
omitted if formalin is unavailable. In most field situations, the most frequently used
alternative preservative is alcohol. High-proof distilled liquor (e.g., rum, aguardiente,
gin) is readily available almost everywhere (except in Moslem countries) and can be used
27
to preserve specimens if necessary. “Denatured” alcohol (usually ethanol rendered
poisonous by admixture with methanol) or “rubbing” alcohol (usually isopropanol) are
other options. In order to insure that the animal does not rot before it is fully preserved,
open both the abdomen and the chest cavity and make sure that they are infused with
alcohol. Do not overcrowd specimens in storage containers, and make sure that there is a
high fluid/specimen volume ratio.
Storage and transport
After fixation, specimens may either be transferred to their permanent storage
medium, or they may be packed for shipment back to the laboratory or museum. The
only appropriate storage media are 70% ethanol (= ethyl alcohol) or 70% isopropanol
(isopropyl alcohol). Most museum prefer ethanol, but isopropanol is used in some
collections. If you plan on depositing your specimens in a particular museum, check in
advance to find out which kind of alcohol they use. Specimens should not be transferred
from one type of alcohol to another without soaking in a series of distilled water baths,
followed by transfer through a series of graded concentrations of the new alcohol. It is
much simpler to store specimens initially in the form of alcohol they will remain in
permanently. Before transferring a specimen from formalin to alcohol, the specimen
should be rinsed (not soaked) in water. After rinsed specimens have been immersed in
alcohol for about a month, they should be transferred to fresh alcohol to eliminate any
residual formalin that has leached from the tissues.
To safely store formalin-fixed specimens in the field, remove them from the
preservative and wrap them individually in cheesecloth that has been moistened in
28
formalin. Wrapped specimens should be placed together in a plastic bag to which enough
formalin has been added to wet all of the packing material. Pour off any fluid remaining
at the bottom of the bag, squeeze out as much air as possible, and seal the bag tightly.
Although “ziplock” bags may be used, larger bags made of heavier plastic that can be
sealed with heavy rubber bands hold more material. Double- or triple-bag each batch of
wrapped specimens to prevent leakage. Alternatively, wrapped specimens may be
packed and sealed in nalgene bottles or other waterproof containers. If you are packing
specimens for shipment by airplane (as cargo, baggage, or mail), be sure that your
packaging meets IATA regulations (see Permits and regulations above).
Preparing Dry Skins
If time permits or pelage variation seems important, preparation of dry skins may
be desirable. Skinning and stuffing bats is a complex procedure that is best learned by
working with someone who is already a proficient preparator. Practicing the whole
procedure several times on unimportant specimens of bats that commonly occur around
your home institution is highly recommended. When in doubt, it is probably always
better for a novice to preserve important voucher specimens in fluid rather than
attempting to skin them.
The initial steps of skin preparation are the same as those described above for
preserving specimens in fluid: the bat must be killed and measured, and all relevant data
should be recorded in the field catalog. A skin tag with all data recorded on it should be
prepared and set aside. Similarly, tags should be prepared for the skull and carcass,
which will preserved separately. The principal subsequent steps of preparing the
29
specimen are (1) skinning and stuffing the skin; (2) tagging the specimen; (4) pinning and
drying the skin; (5) preparing the skull and skinned carcass for preservation. In general,
skins should not be prepared during your last week in the field because thorough drying
can take several days in humid environments.
Skinning and stuffing
Skinning small mammals is a race against time. Premature drying is a major
problem, so it is important to have all your tools handy and to proceed quickly once you
begin the skinning process. With bats, drying of the fur-bearing part of the skin is only
part of the problem; wing membranes become brittle as they dry and are easily damaged.
Before making the first cut, therefore, make sure that you have completed measuring the
animal and recording necessary data in both the field catalog and on the skin tag. Prepare
a cotton filling from a thin triangular piece of cotton in which there are no lumps. Roll or
fold the cotton to form a conical form with roughly the same proportions as the body of
the bat (the tip of the cone will go in the nose of the skin, replacing the skull; the
remainder of the form will fill the neck and body cavity, filling the space previously
occupied by the rib cage and internal organs). The density of the form should imitate that
of the abdomen of the bat: do not make the form too squishy or it will not serve to
adequately support the skin during drying.
Very sharp scissors and a blunt probe are the only necessary dissecting tools. The
scissors are used when cuts are necessary; the probe when stripping skin or muscle from
the animal. Although a scalpel may be useful for large animals (e.g., pteropodids), it is
30
easier to damage the skin accidentally with a scalpel than with scissors. In many
situations, your fingers are more effective tools than any metal instruments.
Begin the skinning process by making a midventral incision in the skin beginning
just anterior to the genital region and extending anteriorly to the middle of the chest.
Take care to avoid cutting through the abdominal wall. Many collectors use sawdust as
an absorbent medium, but clumps of sawdust may cause the finished skins of small bats
(<50 g) to appear lumpy. As an alternative, paper towels can be used to absorb blood
during the skinning process.
Once the midventral incision has been made, run your fingers or the probe
between the skin and the underlying muscle to separate the skin from the body. Do not
pull on the skin, as it may tear. Posteriorly, separate the skin from the body around the
genital region to expose the sides of the anus and the base of the penis (males) or the
vaginal opening (females). Clip through these structures right at the body wall, making
sure to leave the penis or labia attached to the skin. Next, peel the skin from around the
thighs and the base of the tail. Clip through the base of the tail right next to the pelvis; be
sure to leave the tail vertebrae in the skin. Clip through the femora (upper leg bones)
near the pelvis, and peel the skin up around the back of the bat once both legs have been
separated from the body.
Because the legs of bats may contain considerable muscle tissue that can impede
the drying process (or decay if the specimen is not dried quickly enough), it is best to
remove most of the leg muscles. Peel the skin down the leg to the level of the ankle, and
clip most of the muscle away. Keep the tibia intact. Be careful not to cut through the
fibula, which may be very thin or incomplete, resembling a string of gristle more than a
31
bony element. If you are not sure where the fibula is located, it is best to leave the
muscles of the lower leg intact. After cleaning the legs, slide them back into their normal
position in the skin.
Next, roll the skin (with the legs attached) up over the thorax of the bat, carefully
separating the skin from the body on all sides as you go. Expose both sets of shoulder
muscles, and then concentrate on one wing. Separate the skin from the underlying
muscle to a point roughly halfway down the upper arm, and then cut through the humerus
just below the shoulder. Most of the humerus and all of the more distal skeletal elements
of the wing will remain in the skin. Repeat this process on the other side of the animal.
At this point the skin (including the legs and wings) will remain attached to the body
carcass only at the neck and head.
As with the legs, the mass of muscle tissue associated with the forelimbs should
be removed. Roll the skin down one wing to expose the muscles of the upper arm and
forearm, and clip away as much of the muscle as possible. Be careful not to cut the ulna,
which tapers to a thin shaft distally as it approaches the wrist. When sufficient muscle
tissue has been removed, wrap a small bit of cotton around the jagged end of the humerus
(to prevent it from cutting through the skin), and slide the skin back up over the proximal
wing elements before moving on to skin the head.
Grasp the body in one hand and the skin in the other, and gently pull the skin up
over the neck and head. The neck is short, so the base of the skull will appear rapidly.
Using your fingers or the probe, peel the skin away from the back of the skull. As the
fold of retracting skin approaches the ear region, you will see a stiff but flexible tube
connecting the ear region on each side of the skull to the adjacent skin (remember that the
32
external ears will not be visible since they are on the outside). Run the tip of your
scissors around the front of each ear tube, being careful to keep the blade pressed close to
the surface of the skull. Clip each ear tube as close to the skull as possible. Continue to
peel the skin away from the skull, taking care to look for the posterior edges of the eyes,
which usually appear as thin black lines.
Separating the skin from the skull around the eyes is perhaps the most difficult
part of the skinning process. Remember that the skin of the face is continuous with the
eyelid, which in turn is continuous with the mucosa surrounding the eyeball. Once you
have located the eye, pinch up the skin at the eyelid to create tension, and clip between
the eyelid and the skull, continuing around the eyeball until the skin is completely free of
the eye. It is best to leave the eyeball in the skull, but the eye can be removed if
necessary and subsequently excised from the skin. Be especially careful not to cut the
postorbital process during the skinning process; in some bats (e.g., emballonurids) it is a
long, slender process that projects just posterior to the eye.
After the skin is free of both eyes, continue separating it from the skull until you
come to the corners of the mouth. Clip through the skin at the corner of the mouth on
each side, and separate the skin from skull and jaw along the gum line. Depending on the
size of the animal, this can be done either with your fingernails or with scissors. At this
point, the skin should be attached to the skull only at the snout; it should be completely
free of the lower jaw. If there are any glands on the muzzle, carefully remove them from
skin. Finally, pull the skin gently away from the snout, and carefully clip through the
nasal cartilage to complete the skinning process. Set the carcass aside, preferably in a
plastic bag or other closed container to prevent it from drying out.
33
Before turning the skin right side out, it is important to sew the mouth closed.
Using a needle and thread, sew the edges of the skin together beginning at the corner of
the mouth on one side and continuing around to the other side. Use small, even stitches,
and be careful to avoid damage to the tubercles of the chin region, which should be left
well outside the mouth (so they can be examined on the dry skin when it is completed).
After the mouth has been sewn shut, check the skin carefully and remove any excess fat
that may be adhering to the inside of the skin. Fat deposits are common on the snout and
in the mammary and genital areas, and these must be removed before stuffing to prevent
the skin from becoming greasy after it dries.
To turn the skin right side out, reach in from the fur side of the skin with your
forceps, and carefully grasp the nose region of the skin. Pull gently on the nose while
pushing from the other side with your finger to invert the skin so that the fur side is out.
Examine the fur, and remove any crusted blood or other material with your fingers.
Never use a brush on the fur of a specimen; if the skin needs to be cleaned, use a cotton
ball moistened with water.
Lay the skin out on a flat surface, and take up the cotton body form prepared
earlier. Turn back the tip of the body form so that it forms a dense knob of cotton
approximately the same breadth as the rostrum of the bat. Grasp the knob with your
forceps, and gently slide the body form into the skin until the knob of cotton reaches the
nose region. Release the knob of cotton and, using your forceps, gently pack the head of
the skin with cotton from the body form. Take care to avoid stretching the skin; try to fill
it in such a way as to mimic the proportions of the living animal. Moving from anterior
to posterior, use your forceps to pack cotton into the skin. More cotton may be added to
34
the abdominal region if the form proves to be too small; cotton may be removed if the
form is too large. Make sure that the filling is symmetrical, particularly in the posterior
end of the body. When the skin is well filled with cotton, tuck the remaining fibers into
the abdomen and draw the edges of the midventral incision together. Sew the edges of
the incision together using small, even stitches beginning in the genital region and
working forward. If the animal is a male, make sure that the penis is on the outside of the
skin when the incision is closed.
Some of these instructions may seem unusual to researchers familiar with
preparing skins of nonvolant mammals or birds. Bat skins do not require any wires,
which are traditionally inserted in the skins of nonvolant mammals to support the legs
and tail. And unlike bird skins, in which the skull is left in place, the skull is always
removed from the skin before stuffing mammals (including bats). Bat skins are also
dried belly-down (like other mammals) rather than belly-up (like birds); see below.
Tagging
Tags should always be tied to the right leg of the animal in order to reduce
tangling when specimens are laid out side-by-side in a museum tray. Tie the tag around
the leg midway between the ankle and the knee, inserting the tag string through the wing
and tail membranes with a large needle (like those sold for darning socks). The tag string
should project from the dorsal surface of the membranes, not the ventral surface. A
length of string (at least 1 cm) should be left between the tag and the animal to allow
manipulation of the tag without damage to the specimen. Never tie a tag around the
35
ankle of a bat, as this results in permanent distortion of the calcar and the membrane
attachments to the ankle and foot.
Pinning and drying skins
Proper pinning is one of the keys to making a durable and scientifically useful bat
skin. Many substrates may be used for pinning, but the best seems to be sheets of stiff
corrugated cardboard. Whatever you chose as a substrate, make sure that it is not too
flexible, and that the surface allows relatively easy insertion of pins (most wood is too
dense). Once inserted, pins should stay in place without shifting or coming loose when
the substrate is moved. Long stainless-steel sewing pins with round plastic heads work
well. Narrow-gauge entomology pins bend too easily to be optimal, and sewing pins with
flat metal heads are too short and hard to handle. It is important that pins be stainless
steel, as the moisture in the skin may cause non-stainless pins to rust and damage the
skin.
To begin the pinning process, place the bat belly-down on the pinning board with
the wings folded. The forearms should be aligned approximately parallel to each other
with the elbows adjacent to (but not hidden underneath) the sides of the body and the
wrists not too far apart (see Figure 5 for position and locations of pins). Placement of the
first four pins will anchor the specimen and determine the positions of the wings and
body. Begin by putting the first pin (Pin 1) through the wing membrane just posterior to
one of the wrists. Pin 2 goes through the muscle of the arm just lateral to the elbow on
the same side of the bat. Readjust the body and the other forearm, and insert Pins 3 and 4
to anchor the other forearm. Next, stretch the body out lengthwise and pin the knees out
36
Figure 5. Diagram showing the placement and sequence of pins necessary to properly
pin a bat skin for drying; see text for discussion. The bat shown has a short tail, see inset
for how to pin a long-tailed bat.
to the sides with Pins 5 and 6. Make sure that legs are extended adequately to spread the
tail membrane, but are not pulled out of their natural position (it is possible to stretch the
skin into an unnatural shape if you pull too hard). Spread the tail membrane out flat by
moving the lower legs apart, and pin the feet next. Pins 7 and 8 should go through the
foot between the metatarsals. Make sure the toes are spread and the claws are straight
(not twisted to one side or the other). Next, roll the posterior edge of the tail membrane
out by extending the calcars posteriorly, and place Pins 9 and 10 near but not at the tips
of the calcars (this placement will allow accurate measurement of calcar length; placing
the pin right at the tip of the calcar can make it hard to measure). If the bat has a long
tail, extend the tail and place a pin on either side of the tip.
At this point, all that remains to pin are the distal wings. Be sure to arrange the
fingers so that every bone is visible for measurement from the dorsal side. To make sure
that the wings end up symmetrical after pinning, it is best to work with both wings at the
same time (rather than pinning out one wing completely before moving on to the other).
Begin by placing pins through the wing membranes near the tip of each wing (Pins 11
and 12). Take care not to overextend the wing; the third finger is naturally bowed in
most taxa. Next, draw the third metacarpals outward (away from the body) to spread the
metacarpals, and place Pins 13 and 14 through the membranes to hold them in place.
Pins 15 and 16 serve to hold the lateral edge of the wings in place; these are only
necessary if the wing has a tendency to fold up on the edge.
Finally, fold the thumbs
back against the lateral edge of the wing, and hold them in place with pins alongside (do
not try to put pins through the thumbs). If the bat has a noseleaf, a final pin (Pin 19)
should be placed in front of the nose to hold the noseleaf erect during drying. After
37
pinning is complete, you may need to readjust the ears to make sure that they are upright
and symmetrical.
The pinning instructions given above are guidelines; keep in mind that different
bat species present different challenges due to different body forms and wing shapes.
Adjust your pinning technique accordingly, keeping in mind that the goal is to produce a
durable skin in which all of the critical elements (e.g., metacarpals, phalanges, calcar,
tail) are undistorted and exposed for measurement.
Skins should be kept pinned until they are completely dry, which may take several
days even for small bats. If specimens dry too slowly, decay or fungal growth can occur,
particularly on those parts of the skin that are pressed to the pinning board. Dying skins
adequately may not be a problem in dry climates, but in humid areas it can be difficult.
Do not place skins uncovered in direct sunlight, as this may cause changes in fur color.
In the tropics we have successfully dried skins by placing our pinning boards in steel
footlockers in the sun; we have also used our car as a drying oven. If these methods are
not feasible, skins may be dried safely by placing pinning boards in the sun under double
layers of paper. Each pinning board should be elevated above the ground (to facilitate
airflow and discourage insect pests), and boards should be oriented so that the skins lie
head-first toward the sun (to prevent one side of the body from drying faster than the
other, perhaps leading to lateral distortion).
When specimens are dried in the field in humid areas, it is important that they be
covered at night and during rainy periods to prevent them from rehydrating in the higher
humidity. Pinning trays may be placed in large plastic bags (or metal or plastic
footlockers) at these times, and removed the following day to continue drying.
38
If controlled indoor space is available, skins may be dried by leaving them in an
air-conditioned room, a dry cabinet, or near a heat source such as a stove in a wellventilated area. Drying specimens very rapidly (e.g., in an oven) is not desirable because
it may cause shrinking and warping of the specimen. In all cases, it is critical to protect
skins from insect pests during the drying process. The order of drying is usually skin,
ears, wings, and feet. When the feet are dry and no longer flexible, the specimen can be
unpinned.
Carcass preservation
After the skin has been pinned and set aside to dry, the carcass and skull should
be prepared for preservation. The entire carcass may be prepared as a skeleton, or the
skull may be removed for cleaning and the carcass preserved in formalin. Regardless, a
tissue sample should be taken from the specimen before proceeding further (see
Collection Tissues below).
It is useful to tag both the skull (around the lower jaw) and the body (around the
clavicle) regardless of the preparation technique to insure that all parts of the animal
remain linked with their data. If the entire specimen is to be skeletonized, remove the
viscera and, in larger bats, the fleshy parts of the pectoralis muscles. The remaining
carcass may be either dried or stored in alcohol (not formalin) pending final preparation
back at the museum. In our experience, alcohol preservation works especially well in the
tropics as a means of preventing fungal growth and egg deposition by flies. Dermestid
beetles, maintained in most museums for use in cleaning skeletons, prefer alcoholpreserved specimens (that have been soaked out in water and dried after arriving in the
39
museum) to those that have been allowed to develop fungus growth in the field. If
alcohol is in short supply, the specimen may be soaked briefly in alcohol and then
removed for drying, which will greatly reduce fungal growth and discourage flies.
If the body and internal organs are to be preserved in formalin, the skull should be
removed. Separate the skull from the rest of the body by clipping through the vertebral
column near the skull with a pair of sharp scissors taking great care not to damage the
back of the skull. After removal, the skull can be air-dried or stored in alcohol. The
carcass should then be injected with formalin and treated as described above in the
section on fluid preservation. Because the skin has been removed, fixation of carcasses
takes less time than fixation of whole bats, so check frequently to insure that you do not
overfix the specimen.
Storage and transport
Storage and transport of skins can be tricky because you must protect the
specimens from both mechanical damage and insect pests. The easiest way to exclude
insect pests is to use a commercially available insecticide or repellant that comes in solid
form (e.g., moth balls). Make sure that there is enough to adequately protect the skins,
but make sure that the skins are not in physical contact with solid insecticide or repellant,
as contact with these chemicals may cause changes in fur color. If your prepared skins
are to be shipped or otherwise transported in some fashion that may cause them to shift
about, you may want to wrap the repellant in cheesecloth (or stuff it in a clean sock) to
insure that it does not accidentally come in direct contact with the skins.
40
Skins should ideally be packed in a small but stout wooden box between a series
of cotton layers separated by cardboard sheets. Spread a sheet of cotton in the bottom of
the box, and place skins on the cotton so that they are not in contact with each other or
with the sides of the box. Fold the tag under each specimen so it will not get tangled with
other specimens or tags. If the bat has large upright ears or a large noseleaf, you may
want to pack extra cotton around the head to protect them. When a layer of skins has been
completed, cover it with another sheet of cotton. Then place a sheet of cardboard over
the cotton. Repeat the process as many times as necessary to accommodate all of your
skins. If your box is larger than needed, top the last layer of cotton with a cardboard
sheet, and fill the rest of the box with crumpled newspaper or some other filler that will
keep the skins from shifting during transport. Never wrap skins in plastic or any other
impermeable material on which residual moisture might condense during shipment.
Skeleton Preparation
Another method of specimen preparation is to prepare the entire animal as a
skeleton. Skin the bat as described above, but do not clip through any of the bones.
Because the skin will be discarded, it may be cut anywhere to facilitate removal. When
skinning the hind limbs, be careful not to disarticulate the calcar. The tail should also be
skinned with care to avoid accidentally pulling off the terminal vertebra, which may be
attached to the skin with ligaments. Similarly, take care to keep the fingers of the
specimen fully articulated when skinning the wings.
It is possible to prepare a half-skeleton while simultaneously preparing a dry
study skin. In this procedure, the leg and wing bones from one side of the animal are left
41
in place in the skin, while the bones from the tail and the other side of the body are
extracted along with the carcass. This process results in a dry skin with one complete
wing and one wing without supporting bony elements, and a clean skull and half
skeleton. For additional instructions regarding this process, see Handley (1988).
Once the skin has been removed, take a tissue sample from the specimen if
desired, and then remove the internal organs including the heart and lungs (which
requires cutting through the diaphragm). Trim away as much muscle tissue as you can
from the pectoral region, shoulders, upper arm, forearm, and legs. Remember to take
care to avoid cutting through the ulna and fibula.
Attach one tag to the skull (through the lower jaw) and another to body (through
the clavicle or around the vertebral column in the lumbar region). The skull may be
removed or left articulated. To aid in storage and to prevent tangling with other
specimens, fold the hind legs up to the belly region, and fold the wings in their natural
fashion (which varies from species to species). Wrap a length of heavy (buttonhole)
thread several times around the skeleton to form a neat bundle.
Storage and transport
As noted above under Carcass Preservation, skeletons can be dried or stored in
alcohol before being subjected to further cleaning back in the museum or lab. Alcohol
preservation works best in the tropics as it prevents fungal growth and egg deposition by
flies. As noted earlier, skeletons stored in unbuffered formalin tend to decalcify, and
dermestid beetles do not much care for formalin-preserved flesh.
42
Storage and transport of skeletons follow the same procedures outlined above for
fluid-preserved specimens with alcohol substituted for formalin in the bagging process.
70% ethanol is ideal, but isopropanol or other concentrations of either form of alcohol
may be used if necessary.
Collecting Tissues
Tissue samples have become increasingly important in almost every field of
research (systematics, population genetics, biogeography) and should be collected
whenever possible. Because several publications already provide thorough descriptions
of methods for collecting, storing, and archiving tissue samples (e.g., Dessauer et al.,
1990, 1996; Longmire et al., 1997; Kilpatrick, 2002; Prendini et al., 2002), the following
review emphasizes aspects of tissue collection, storage, and transportation that are
maximally useful for bat researchers working in the field. Because DNA is difficult to
extract from specimens that have been fixed in formalin, it is important to save tissues
samples even when entire animals are to be kept as fluid-preserved vouchers.
Storage containers and labels
Proper storage containers and preservatives should be obtained before leaving for
the field. Flat-bottomed 1.8 ml plastic cryotubes with internal threading (e.g., NUNC
CryoTube Vials ™) are ideal; larger tubes are usually unnecessary and smaller tubes are
hard to label effectively. Avoid tubes with external threading, as the threads may break
off at low temperatures (e.g., if they are placed in liquid nitrogen or an ultracold freezer).
If field numbers are written on the outside of the tube, be sure to use tubes provided with
43
an appropriately textured writing surface (the smooth plastic of the rest of the tube will
not hold ink or pencil). Appropriate cryotubes can be easily obtained from any scientific
supply house, or from institutional tissue collections or biorepositories.
As with voucher specimens, it is critical that tissue samples be properly labeled in
the field. In order to prevent accidental mixups, tubes (or set of tubes) should be labeled
just prior to inserting the sample. Use pencil or ink that is demonstrably both waterproof
and alcohol-proof when dry, and record the field number of the specimen directly on the
tube. The number should be identical to that of the rest of the animal (skin, skull, fluid,
etc.) if there are anatomical vouchers associated. All samples taken from a given
individual should have the same number.
Some institutional tissue collections (e.g., The Ambrose Monell Cryo Collection
at the American Museum of Natural History; http://research.amnh.org/amcc/) provide
researchers with cryotubes prelabelled with unique barcodes and numbers printed on
thermal- and solvent-resistant material suitable for long-term storage at ultracold
temperatures. If you are using such tubes, it is essential that you record each tube number
under the appropriate entry in your field catalog. If the link between specimen and data
is broken, the sample will be worthless.
Preservatives
Preservation of tissue samples may be accomplished in one of several ways, with
the choice typically dependant on field conditions and the use intended for the samples.
The best results for molecular studies of vertebrates are typically obtained from fresh
samples frozen soon after collection using either dry ice or liquid nitrogen (Prendini et al.
44
2002). Sources of dry ice are available in most cities; liquid nitrogen is typically
available from medial and veterinary clinics, hospitals, and universities. New security
regulations now make it impossible to carry liquid nitrogen on airplanes, but “dry
shippers” (infused with liquid nitrogen that is subsequently poured off prior to shipping)
remain cold enough to preserve frozen samples for up to 10 days. If you plan to freeze
samples, it is important to plan ahead to make sure that you can keep your samples frozen
continuously until they are deposited in freezers in the lab or museum. Initial freezing
should be as rapid as possible; sample tubes can be snap-frozen by dropping them
directly into liquid nitrogen or a dry shipper, or by covering them with dry ice.
A less elaborate method of preserving tissue samples is to place them directly in
95-100% ethanol (Kilpatrick, 2002; Prendini et al., 2002). This technique yields samples
from which high molecular weight DNA can be isolated in quantities nearly comparable
to those obtained with freezing (Prendini et al., 2002), and it has the advantage of being
easy in remote field situations. Although ethanol-preserved tissues do not need to be
refrigerated in the field, they should eventually be transferred to an ultracold freezer for
long-term archival preservation.
Tissue samples can also be preserved in lysis buffer solution (e.g., 2% 2phenoxyethanol with glycerol or DMSO, 2-propanol with ethylene-diamine-tetraacetate
[EDTA] or sodium dodeyl sulphate [SDS]), a recipe for which is given in Appendix 2.
Under some circumstances, mammalian tissues preserved in this manner yield larger
quantities of PCR products than those preserved in ethanol (Kilpatrick, 2002). See
Longmire et al. (1997) and Kilpatrick (2002) for discussions of the pros and cons of
preservation in buffer solutions. In an emergency, even laundry detergent has been
45
shown to preserve vertebrate DNA effectively (Kuch et al., 1999). Many buffer solutions
can be used in the field without refrigeration or freezing (e.g., see Appendix 3), but others
require refrigeration or freezing after 24 hours to insure proper preservation (Prendini et
al., 2002). Be sure to check the requirements for storage of your chosen buffer solution
before leaving for the field.
Organ sampling
Tissue samples should be obtained as soon as possible after death. Solid tissues
(e.g., muscle, liver, kidney) destined for preservation in ethanol or lysis buffer should be
cut into small pieces (no more than 5 mm diamenter) to allow rapid penetration of the
preservative. Be careful to clean your dissecting tools carefully in between each tissuesampling episode to avoid contamination. Scissors can be effectively cleaned of any
foreign DNA-bearing tissue in a flame before collecting fresh tissue samples.
Blood
Small blood samples can be collected in heparinised hematocrit or
microcentrifuge tubes; larger samples are most efficiently collected using a heparinised
syringe (Dessauer et al., 1990, 1996; Prendini et al., 2002). If you plan to preserve blood
samples by freezing, be careful if you are using hematocrit or microcentrifuge tubes, as
these may shatter if snap frozen (Prendini et al., 2002). It is best to freeze them slowly
before subjecting them to ultracold temperatures.
Another method of collecting blood samples is to use Nobuto® blood filter strips
(available from most medical supply houses). These strips can be used to collect small
46
quantities of blood (0.1mL of blood or 0.04mL of serum) that is then dried. Typical
applications include storage of samples for testing for toxoplasmosis and other diseases.
Wing punches
A commonly used method for obtaining tissues samples from living bats is to take
wing punches using a commercially available biopsy punch. This method was described
in detail by Worthington-Wilmer and Barratt (1996), and has been used extensively since
that time. Biopsy punches come in variety of sizes (2mm -- 8mm) and can be chosen
based on the size of the study species. Wing punch tissue samples can be stored like any
other tissue. Once a wing punch has been removed, healing is rapid with complete
closure of the wound in 2–4 weeks.
If wing punches are taken from a sample of bats that will be released, it is highly
desirable to retain at least one individual as a voucher to allow future confirmation of the
specific identity of the sampled series should questions arise.
Storage and transport
Methods of tissue-sample storage and transport depend on the type of preservative
used. Be sure to check on the requirements of your chosen preservative and method
before leaving for the field. In general, it is safe to assume that all tissue samples are best
stored in a secure location away from light or heat sources. If traveling or shipping by
air, be sure to check airline regulations concerning transport of chemicals. If traveling
across international boundaries, be sure to check regulations for transport of biological
specimens (see Permits and Regulations above).
47
Literature Cited
American Society of Mammalogists Animal Care and Use Committee. 1998. Guidelines
for the capture, handling, and care of mammals as approved by the American
Society of Mammalogists. Journal of Mammalogy, 79: 1416-1431.
American Veterinary Medical Association. 1993. Report of the AVMA Panel on
Euthanasia. Journal of the American Veterinary Medical Association, 202: 229249.
Anthony, E.L.P. 1988. Age determination in bats. Pp. 47-58, In: Ecological and
Behavioral Methods for the Study of Bats (T. H. Kunz, ed.). Smithsonian
Institution Press, Washington, DC.
Dessauer, H.C., C.J. Cole, and M.S. Haffner. 1990. Collection and storage of tissues.
Pp. 25-41, In: Molecular Systematics (D. M. Hillis and C. Moritz, eds.). Sinauer,
Sunderland, MA.
Dessauer, H.C., C.J. Cole, and M.S. Haffner. 1996. Collection and storage of tissues.
Pp. 29-47, In: Molecular systematics, 2nd edition (D. M. Hillis, C. Moritz, and B.
K. Mable, eds.). Sinauer, Sunderland, MA.
Handley, C.O., Jr. 1988. Specimen preparation. Pp. 437-457, In: Ecological and
Behavioral Methods for the Study of Bats (T. H. Kunz, ed.). Smithsonian
Institution Press, Washington, DC.
Kilpatrick, C.W. 2002. Noncryogenic preservation of mammalian tissues for DNA
48
extraction: an assessment of storage methods. Biochemical Genetics 40: 53-62.
Kuch, U., M. Pfenninger, and A. Bahl. 1999. Laundry detergent effectively preserves
amphibian and reptile blood and tissues for DNA isolation. Herptetological
Review, 30: 80-82.
Longmire, J.L. 1997. Use of “lysis buffer” in DNA isolation and its implications for
museum collections. Occasional Papers, Museum of Texas Tech University 163:
1-3.
Nelson, D.W., and J. Sparks. 1999. Paraformaldehyde/Alconox problems. American
Society of Ichthyologists and Herpetologists Curation Newsletter 12: 1-2.
Prendini, L.,R., Hanner, and R. DeSalle. 2002. Obtaining, storing, and archiving
specimens and tissue samples for use in molecular studies. Pp. 176-248, In:
Techniques in Molecular Systematics and evolution (R. DeSalle, G. Giribet, and
W. Wheeler, eds). Birkhäser Verlag, Basel.
Racey, P.A. 1988. Reproductive assessment in bats. Pp. 31-45, In: Ecological and
Behavioral Methods for the Study of Bats (T. H. Kunz, ed.). Smithsonian
Institution Press, Washington, DC.
Simmons, N.B. 2003. Chiroptera in "Mammal Species of the World -- A Taxonomic
and Geographic Reference." Bat Research News, 43: 183-184.
Simmons, N.B. 2005. Order Chiroptera. In: Mammal Species of the World: a
Taxonomic and Geographic Reference, Third Edition (D. E. Wilson and D. M
Reeder, eds.). Johns Hopkins University Press.
Voss, R.S., and L.H. Emmons. 1996. Mammalian diversity in neotropical lowland
rainforests: a preliminary assessment. Bulletin of the American Museum of
49
Natural History 230: 1-115.
Wilson, D.E., and F.R. Cole. 2000. Common names of mammals of the World.
Smithsonian Institution Press, Washington, DC.
Worthington-Wilmer, J., and E. Barratt. 1996. A non-lethal method of tissue sampling
for genetic studies of chiropterans. Bat Research News, 37: 1-3.
50
Appendix 1: Preparing 10% Buffered Formalin
“Formalin” and “formaldehyde” are often used interchangeably to refer to the
commonest general-purpose fixative solution used by field biologists, but these terms are
not synonymous. Full-strength (100%) formalin is a saturated (37%) aqueous solution of
formaldehyde gas to which a small quantity of methanol is often added as a stabilizer.
Because formaldehyde solutions are naturally unstable and tend to acidify over time, fullstrength formalin should always be bought fresh from a reputable supplier just prior to
the field season when it will be used. Never use old formalin to preserve important
specimens.
To make 10% formalin, simply mix 1 part full-strength formalin with 9 parts
water (by volume). Wear rubber gloves to protect your hands from this solution, and do
the mixing in a well-ventilated area, preferably outdoors. For the short length of time
that bats are usually immersed in this fixative, buffering agents are seldom needed.
However, buffering is still a good insurance against the possibility that fixed specimens
must be left at a base camp for extended periods or end up warehoused while export
permit problems are worked out. In such situations, significant decalcification of bones
and teeth will occur.
Histologists buffer 10% formalin by adding 6.5 g of dibasic (anhydrous) sodium
phosphate and 4 g monobasic sodium phosphate per 1 liter. A tried-and-true substitute
for these chemicals that is often used by field biologists is commercial borax, which can
be added to 10% formalin in the proportion of about 3 ml per liter. If no other buffering
agent is available, blackboard chalk is better than nothing.
51
Formalin can also be prepared by dissolving paraformaldehyde powder in water.
However, paraformaldehyde does not dissolve easily and at least some commonly used
wetting agents (e.g., Alconox) that help paraformaldehyde dissolve in water may
compromise long-term specimen storage (Nelson and Sparks, 1999). Also, because
paraformaldehyde is a white powder that cannot be safely sniffed or tasted, it may be
hard to convince skeptical customs officials or local police that it is not a controlled
substance.
Appendix 2: Preparing Lysis Buffer
A liter of lysis buffer suitable for preserving bat tissues without refrigeration or
freezing in the field can be prepared according to the following recipe (Longmire et al.,
1997):
Mix the following in order:
50 ml of 2 M Tris-Hcl, ph. 8.0
200 ml of 0.5 M EDTA, ph. 8.0
2 ml of 5 M NaCl
Distilled water up to 975 ml
25 ml of 20% SDS
This solution should be protected from light and heat.
52