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Special Publications
Museum of Texas Tech University
Number xx xx XXXX 2010
Special Publications
Museum of Texas Tech University
Number 76 19 January 2021
Hummingbird (Family Trochilidae) Research:
Welfare-conscious Study Techniques for
Live Hummingbirds and Processing of
Hummingbird Specimens
Lisa A. Tell, Jenny A. Hazlehurst, Ruta R. Bandivadekar, Jennifer C. Brown,
Austin R. Spence, Donald R. Powers, Dalen W. Agnew, Leslie W. Woods, and
Andrew Engilis, Jr.
Front cover: Photographic images illustrating various aspects of hummingbird research. Images provided courtesy of Don
M. Preisler with the exception of the top right image (courtesy of Dr. Lynda Go).
Dedications
To Sandra Ogletree, who was an exceptional friend and colleague. Her love for family, friends,
and birds inspired us all. May her smile and laughter leave a lasting impression of time spent with her
and an indelible footprint in our hearts.
To my parents, sister, husband, and children. Thank you for all of your love and unconditional
support.
To my friends and mentors, Drs. Mitchell Bush, Scott Citino, John Pascoe and Bill Lasley. Thank
you for your endless encouragement and for always believing in me.
~ Lisa A. Tell
SPECIAL PUBLICATIONS
Museum of Texas Tech University
Number 76
Hummingbird (Family Trochilidae) Research: Welfare-
conscious Study Techniques for Live Hummingbirds and
Processing of Hummingbird Specimens
Layout and Design: Lisa Bradley
Cover Design: Lisa A. Tell and Don M. Preisler
Production Editor: Lisa Bradley
Copyright 2021, Museum of Texas Tech University
This publication is available free of charge in PDF format from the website of the Natural Sciences Research
Laboratory, Museum of Texas Tech University (www.depts.ttu.edu/nsrl). The authors and the Museum of
Texas Tech University hereby grant permission to interested parties to download or print this publication for
personal or educational (not for prot) use. Re-publication of any part of this paper in other works is not
permitted without prior written permission of the Museum of Texas Tech University.
This book was set in Times New Roman and printed on acid-free paper that meets the guidelines for per-
manence and durability of the Committee on Production Guidelines for Book Longevity of the Council on
Library Resources.
Printed: 19 January 2021
Library of Congress Cataloging-in-Publication Data
Special Publications of the Museum of Texas Tech University, Number 76
Series Editor: Robert D. Bradley
Hummingbird (Family Trochilidae) Research: Welfare-conscious Study Techniques for Live
Hummingbirds and Processing of Hummingbird Specimens
Lisa A. Tell, Jenny A. Hazlehurst, Ruta R. Bandivadekar, Jennifer C. Brown, Austin R. Spence, Donald R.
Powers, Dalen W. Agnew, Leslie W. Woods, and Andrew Engilis, Jr.
ISSN 0149-1768
ISBN 1-929330-44-8
ISBN13 978-1-929330-44-7
Museum of Texas Tech University
Lubbock, TX 79409-3191 USA
(806)742-2442
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Abstract
Introduction
1. Laying the Groundwork: Study Approval and Permitting
1.1. Literature search
1.2. Institutional Animal Care and Use Committee and Protocols
1.3. Permit requirements
1.3.1. United States Geological Survey Bird Banding Laboratory (BBL) permits
1.3.2. United States Fish and Wildlife Service (USFWS) permit
1.3.3. State permits
1.3.4. Additional permit authorizations
2. Working with Live Hummingbirds
2.1. Hummingbird identication in the United States of America
2.2. Safe practices for capturing hummingbirds
2.3. Handling, restraining, and retaining hummingbirds
2.4. Clinical warning signs when handling and sampling live hummingbirds
2.4.1. Respiratory system
2.4.2. Reproductive system
2.4.3. Torpor
2.4.4. Hypoglycemia and other compromised conditions
2.5. On-site care of hummingbirds
3. Marking Hummingbirds
3.1. Trends and applications
3.2. Feather clipping
3.3. Dye, pigment, paint, and miscellaneous marking
3.4. Banding (color and aluminum bands)
3.5. Acetate plastic colored tags
3.6. Radio-frequency identication/passive integrated transponder tags
3.7. Very high frequency radio telemetry tagging
4. Obtaining and Storing Blood Samples from Hummingbirds
4.1. Trends and applications
4.2. Toenail clipping method for obtaining blood samples from hummingbirds
4.3. Blood specimen storage and analysis
5. Obtaining Feather Samples from Hummingbirds
5.1. Trends and applications
5.2. Methods for sampling feathers from hummingbirds
5.2.1. Appropriate timing for feather sampling: molt
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5.2.2 . Appropriate timing for feather sampling: other considerations
5.2.3. Methods for sampling contour feathers
5.2.4. Methods for sampling wing and tail feathers
5.2.5. Methods for sampling blood feathers
5.3. Methods for feather specimen storage and analysis
6. Obtaining Oral or Cloacal Swabs from Hummingbirds
6.1. Trends and applications
6.2. Methods for oral and cloacal swabbing
7. Obtaining Cloacal Excreta Samples from Hummingbirds
7.1. Trends and applications
7.2. Detailed methods for cloacal excreta sampling from hummingbirds
7.2.1. Cage liner method for excreta sampling
7.2.2. Manual restraint and capillary tube method for sampling cloacal excreta
7.2.3. Dish collection for excreta sampling
7.3. Cloacal excreta specimen storage and analysis
8. Measuring Metabolic Rates in Hummingbirds
8.1. Trends and applications
8.2. Overview of open-ow respirometry methods
8.3. Measuring basal and resting metabolism
8.4. Measuring torpor metabolic rate
8.5. Hovering metabolic and evaporation rates
8.6. Measuring eld metabolic rate
9. Organ Tissue Sampling from Hummingbird Specimens
9.1. Trends and applications
9.2. Detailed methods for sampling and storing organs harvested from hummingbird
specimens
10. Muscle Sampling from Hummingbird Specimens
10.1. Trends and applications
10.2. Detailed methods for muscle sampling from hummingbird specimens
10.2.1. Dissection
10.2.2. Punch biopsy
10.3. Muscle specimen storage and analysis
11. Assessing Birds for Rehabilitation or Euthanasia, Methods of Euthanasia, and Post-Mortem
Examination
11.1. Trends and applications
11.2. Assessing a hummingbird’s condition: administering rst aid
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11.3. Assessing hummingbird health: rehabilitation or euthanasia
11.4. Methods for euthanizing a hummingbird due to injury or illness
11.4.1. Overdose of inhaled anesthetic agent (Isourane)
11.4.2. Overdose of inhaled carbon dioxide (CO2)
11.4.3. Overdose of pentobarbital anesthetic
11.4.4. Rapid cardiac compression (RCC)
11.4.5. Cervical dislocation
11.4.6. Decapitation
11.5. Conrmation of death
11.5.1. Heartbeat
11.5.2. Pupillary light response (PLR)
11.6. Post-mortem examination to determine cause of death
12. Field Collecting Hummingbirds for Specimen-based Research and Museums
13. Balancing Human Safety and Bird Health
Conclusion
Acknowledgments
Disclaimer
Literature Cited
1
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’ experi-
ence, 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, rst-aid measures, and euthanasia of hummingbirds, as well as
processing of hummingbird specimens (e.g., necropsy and preservation).
Key words: bleeding, capturing, euthanasia, handling, marking, metabolic rates, museum
specimens, permitting, restraining, sampling
Numerous species of hummingbirds (Family Tro-
chilidae) have been the subjects of scientic investiga-
tions involving evolution, behavior, physiology, ight
mechanics, feather structure, diseases, and conservation
biology. In addition, due to an increasing interest in
pollinators, hummingbird research appears to be on
the rise. However, given their small body size and
high metabolism, obtaining samples from live hum-
mingbirds can pose challenges, especially for novice
investigators. Sampling techniques have been well
established for other avian species, but these methods
are not always transferable to hummingbirds. To con-
tinue advancing scientic discovery for this unique
family of birds, this manuscript provides guidelines
for obtaining study permits, as well as safe, ecient,
and ethical methods of sampling live hummingbirds
while maintaining high standards of animal care and
welfare. Recommendations for dealing with injured or
extremely ill birds are addressed, along with protocols
for euthanasia. Necropsy techniques, specimen prepa-
ration for museum vouchering, and organ and muscle
sampling also are described in detail.
The study of wild birds remains essential for or-
nithologists who are trying to understand, under natural
conditions, basic questions of ecology, physiology,
disease, and behavior. Capture, restraint, and sampling
of hummingbirds have led to a better understanding
of their physiology and ight mechanics (Chai et al.
1996; Warrick et al. 2005; Bakken and Sabat 2006;
Welch et al. 2007), evolution of biodiversity and mu-
tualisms (Chaves and Smith 2011; González-Gómez et
al. 2014a., b; Maglianesi et al. 2014a, b; Ornelas et al.
2014; Abrahamczyk and Renner 2015; Gonzalez and
Loiselle 2016), disease prevalence (Godoy et al. 2013,
2014; Backus et al. 2019; Magagna et al. 2019; Baek
et al. 2020), and taxonomy and systematics (McGuire
et al. 2009, 2014). Much of this research has been
achieved through obtaining biological samples such as
blood, muscle, organ tissues, excreta, or feathers from
live hummingbirds or preserved specimens.
Although there is much to be gained scientically
by studying hummingbirds, striving for a balance be-
tween sampling risk and scientic gain is paramount.
Risks can be minimized if researchers are vigilant about
the working environment, the hummingbirds’ condi-
tion, the amount of handling time, and employment of
welfare-oriented best practices. Most researchers focus
on the obvious small size and delicate skeletal frame
when working with hummingbirds. North American
hummingbirds range in weight from approximately 2.3
g (Calliope Hummingbird, Selasphorus calliope) to 8.6
g (Blue-throated Mountain-gem, Lampornis clemen-
ciae). However, when working with hummingbirds,
many other factors must be considered. One factor is
the high mass-specic metabolic rate of these species
compared to those of other vertebrates (Suarez 1992).
Given their energy demands, hummingbirds operate at
the limits of their metabolic needs. Therefore, when
handling and restraining hummingbirds, investigators
must be mindful of their caloric needs to ensure suc-
cessful release. Additionally, hummingbirds have the
lowest mass of feathers by weight of any bird. Their
lack of down feathers (King and McLelland 1984)
prohibits over-sampling of feathers, which may alter a
hummingbird’s ability to thermoregulate and increase
the risk of hypothermia (particularly for species inhabit-
ing high elevations). For some hummingbird species,
tail feathers also play an acoustic role during courtship
displays (Clark et al. 2018a; Clark and Mistick 2018a);
therefore, sampling certain feathers during the breeding
season may alter reproductive tness.
As studies incorporating live hummingbird
sampling become more common, investigators should
examine and consistently re-evaluate techniques to
maintain good practices for conducting high-quality
and ethical research. This manuscript summarizes the
established literature that details basic methods used by
banders and scientists who work with hummingbirds
and then suggests best practices for working with live
hummingbirds. In addition, guidelines are provided
for optimizing the utility of birds that have died or are
collected for specimen archiving.
There are numerous approaches for achieving
similar outcomes; therefore, the proposed techniques
should be viewed as based on experience and not as
denitive or exhaustive in nature. Resources and rec-
ommendations listed in this manuscript are intended
to provide new investigators with baseline information
and oer experienced researchers alternative options.
1.1. Literature Search
Performing a thorough literature search prior to
sample or specimen collection minimizes scientic
duplication and maximizes eciency so as to reduce
the potential impact on study subjects. A literature
review will be required by an Institutional Animal Care
and Use Committee (IACUC) if the investigator is em-
ployed by an institution that receives federal funding
for laboratory research and performs research on certain
animal species (see section 1.2). Scientists also should
supply background scientic information when apply-
ing for federal or state permits. Literature searches for
hummingbird-related publications can be performed
using conventional biological literature databases.
The Cornell Lab of Ornithology (http://www.birds.
cornell.edu/Page.aspx?pid=1478) has avian-related
resources available, and Partners in Flight (http://pif.
birdconservancy.org/#) has population estimates and
avian conservation assessment databases. Ornithol-
ogy Exchange has a listing of ornithological journals
(https://ornithologyexchange.org/resources/journals/
database/ornithological-journals/) and other journals of
interest to ornithologists (https://ornithologyexchange.
org/resources/journals/database/other-journals/).
For this manuscript, a reference library of publi-
cations was compiled using search terms “humming-
bird” OR “hummingbirds” OR “Trochilidae.” Searched
databases included SciELO, Web of Science, BIOSIS
Previews, CAB Abstracts, Scopus, EBSCOhost, and
PubMed. Google Scholar was not included because
the search algorithm is not transparent, and results
can contain more false drops (items found that are not
related to the search topic) than useful records. Once
the reference library was established, studies were
evaluated and selectively summarized in tables so that
readers could see the variety of sampling methods
that have been published in peer-reviewed articles.
Manuscript-reported numbers for these tables rep-
resent unique studies where individual publications
were counted only once but appeared multiple times in
one table. This library is available on-line at (https://
www.zotero.org/ucdhummingbirdhealthbibliography/
library). As of September 2020, the library contained
2,039 citations.
The following subsections of this manuscript
discuss pre-study requirements for virtually all types of
research on hummingbirds. Such prerequisites include
following IACUC standards and obtaining permits al-
lowing the capture and processing of hummingbirds.
Depending on the focus of the study, the researcher may
be required to obtain permits issued by the United States
Geological Survey (USGS) Bird Banding Laboratory
(BBL), the United States Fish and Wildlife Service
(USFWS), the state in which the study is performed,
and, potentially, additional authorities.
-
tee and Protocols
The Institutional Animal Care and Use Commit-
tee is appointed to oversee research activities ali-
ated with an institution in the United States (US) of
America. This committee ensures that all procedures
comply with the Animal Welfare Act (AWA 1966,
1970; AWAR 2020) and Public Health Service Policy
on Humane Care and Use of Laboratory Animals (PHS
2015). Investigators working at research institutions
that receive US federal funding must apply for IACUC
protocol approval for any proposed research project that
involves animals or animal sampling. Protocols are
evaluated for scientic merit, animal welfare, procedure
appropriateness, and animal care and maintenance.
For IACUCs that evaluate research protocols involv-
ing hummingbirds, this manuscript could be used as a
reference for best practices. Other helpful references
that summarize concepts and provide guidelines for us-
ing wild birds in research include Wild Bird Guidelines
for Research (Fair and Jones 2010) and the Report of
the Committee on Use of Wild Birds in Research (Or-
ing et al. 1988). In addition, Espin et al. (2014, 2020)
summarized fundamental sampling and sample storage
protocols for raptors, some of which could be applied
to hummingbird research.
Because the authors work and have the most
expertise in the United States of America, the permit
section of this manuscript is limited to this geographic
region. Permit requirements to work with live birds
dier for Canada, Mexico, and Central and South
America and have not been addressed herein. Import/
export permit requirements also vary worldwide. Per-
mitting requirements detailed herein reect policies
that were in place at the time of publication but are
subject to change. Researchers should routinely consult
the websites of federal and state agencies for current
permit requirements. Website addresses (URLs) for
federal agencies are referenced in the footnote section
of Table 1.
The protected status of the hummingbird spe-
cies that will be captured, restrained, banded, marked,
and/or sampled is a key component to the permitting
process. All hummingbird species are listed in Ap-
pendix II of the Convention on International Trade of
Endangered Species of Wild Fauna and Flora (CITES).
CITES is an international agreement that serves to pro-
tect wild animals and plants from international trade
that might threaten their survival. The checklist for
CITES species can be found at http://checklist.cites.
org/#/en. Some hummingbird species also are listed
under the Endangered Species Act (ESA). ESA spe-
cies listings can be found at the website of the USFWS
Environmental Conservation Online System (ECOS) at
https://ecos.fws.gov/ecp/. At the time this manuscript
was prepared, none of the hummingbird species that are
found in or migrate through North America were ESA
listed. Hummingbirds found in North America have
protected status under the Migratory Bird Treaty Act
(MBTA 2020). In addition, special status species may
require additional justication and permitting restric-
tions under permitting programs. Federally designated
“Birds of Conservation Concern” (USFWS Birds of
Conservation Concern: https://www.fws.gov/birds/
management/managed-species/birds-of-conservation-
concern.php) or state-designated “Species of Special
Concern” (i.e., California Species of Special Concern:
https://wildlife.ca.gov/Conservation/SSC or Nevada
Species of Conservation Priority: http://www.ndow.org/
Nevada_Wildlife/Conservation/Nevada_Wildlife_Ac-
tion_Plan/) are examples of special status species.
In the United States of America and its territo-
ries, activities associated with hummingbird research
require federal and often state permits (Table 1). Fed-
eral permits are required when working with live or
Table 1. Required federal (United States Fish and Wildlife Service [USFWS] or United States Geological Survey [USGS] Bird Banding Laboratory [BBL])
or state permits for conducting hummingbird research in California, United States of America.+
Process Method USFWS BBL State
Capturing and marking a hummingbird mist-net, drop-door, etc. N Y Y
Capturing and not marking a hummingbird mist-net, drop-door, etc. Y N Y
Holding a hummingbird in captivity for ≤ 24 hours (only for the safety and well-being of
an individual bird; not for sample collection or procedures relative to research)
any method N Y maybe
Holding a hummingbird in captivity for ≤ 24 hours (for research purposes, including but
not limited to sample collection)
any method Y N Y
Holding a hummingbird in captivity for > 24 hours any method Y N Y
Auxiliary marking of a hummingbird++ color marking, tagging N Y Y
Banding of a hummingbird banding N Y Y
Feather sampling and banding/marking any feather N Y Y
Feather sampling and no banding/marking any feather Y N Y
Blood sampling and banding/marking any method N Y Y
Blood sampling and no banding/marking any method Y N Y
Cloacal or oral swab sampling and banding/marking swab N Y Y
Cloacal or oral swab sampling and not banding/marking swab Y N Y
Fecal/urine collection that requires restraint or containment of the bird restraint or contained holding of
a bird
Y N Y
Fecal/urine collection after the sample has left the bird’s body and without restraining or
retaining the bird
fecal/urine sample collection
from an abandoned nest or under
a free ranging roosting area etc.
N N maybe
Euthanasia due to injuries or pre-existing conditions IACUC approved maybe* Y** Y
Lethal take of birds (healthy birds for study skins, invasive sample collection, disease
studies, etc)
IACUC approved Y N Y
Process Method USFWS BBL State
Treatment of injured bird (during processing or pre-existing condition) IACUC approved N N N
Salvage provisions for bird mortality occurring in association with banding/marking
activities
possession for a period up to 6
months post death
N Y maybe
Salvage activities unrelated to bird banding/marking possession length dependent on
details listed in permit
Y N maybe
+Hummingbirds are protected under the US Migratory Bird Treaty Act (MBTA 2020). At the time that this manuscript was written, no hummingbird species found
or traveling through North America appeared on the federal list of endangered and threatened species. Should a hummingbird species, subspecies, or population be
added to that list, then a researcher proposing to study that species must obtain a USFWS Endangered Species Section 10 Recovery permit in addition to the permits
listed in this table. The USFWS Endangered Species program should be consulted for these permitting requirements. The Endangered Species Recovery permit must
be obtained before the BBL will approve authorizations on a banding permit to capture, band, and/or mark any federally-listed species.
++Auxiliary marking refers to any color or identifying tag or device other than a federally-issued bird band and includes any color marking or tag, for example a very
high frequency or radio-frequency identication tag.
* If an individual has a USFWS Scientic Collecting permit, then they may only euthanize a bird if lethal take of that species is authorized by the permit.
**This is limited to an individual holding a BBL master permit or sub-permit being able to conduct euthanasia only when a hummingbird is injured during permit
approved activities and the permit holder determines that euthanasia is the only proper action given the extent of the bird’s injuries. The BBL permit does not provide
general authorization to euthanize birds and should not be used to euthanize a perfectly healthy bird. The BBL general permit conditions are: If a bird is found injured
or severely diseased, the bander must assess the situation and determine if treatment and rehabilitation would lead to the bird’s recovery. If it is likely that treatment
will allow the bird to recover, the bander should transport the injured bird to an avian rehabilitation facility. If the BBL permittee determines that recovery is not
likely given the extent of injuries or disease, they should euthanize the bird using techniques as advised by the American Veterinary Medical Association or American
Ornithological Council. BBL permittees operating under an institutional animal care and use committee (IACUC) approved protocol need to use IACUC authorized
euthanasia methods.
Note: For current regulations and guidelines, it is advised to routinely consult the United States Geological Survey Bird Banding Laboratory (https://www.usgs.gov/
centers/pwrc/science/bird-banding-laboratory?qt-science_center_objects=0#qt-science_center_objects) and the United States Fish and Wildlife Service (https://www.
fws.gov/birds/policies-and-regulations/permits/permit-policies-and-regulations.php) websites.
Table 1. (cont.)
dead hummingbirds, and nearly every state also has
a separate and unique permitting process for working
with migratory birds, including hummingbirds. The
Ornithological Council provides guidelines, consider-
ations, and links for state permitting at https://birdnet.
org/info-for-ornithologists/permits/states/. Obtaining
new or renewed permits can take considerable time
(sometimes months to years) depending on the agency;
therefore, it is important to plan accordingly. Due to
variation in federal, state, tribal, and local laws, regu-
lations, and policies, permit requirements from these
entities also can vary. If there is a discrepancy between
two separate permits, permittees should follow the most
restrictive permit requirements.
Hard copies or readily accessible digital les of
federal and state permits must accompany the investi-
gator, master bander, or sub-permittees working in the
eld. Permit copies also must accompany samples
during shipment or transport, including specimens
donated to museums or research institutions. Best
practice would be that prior to shipment, permits (if
applicable) are exchanged between individuals send-
ing and receiving the samples to ensure that proper
documentation is in place. Depending on the specic
situation, the researcher may require a permit from
the BBL, USFWS, the state, and/or another authority.
1.3.1. United States Geological Survey Bird
Banding Laboratory (BBL) permits.— Anyone in the
US who is banding, marking, and handling humming-
birds independently in the eld is required to possess
either a valid BBL master permit or sub-permit (50 CFR
§ 21.22). United States BBL permit authorizations are
normally restricted to activities occurring within the US
and its territories. Under very limited circumstances,
the BBL may issue authorizations to use BBL-issued
bands outside of the US on species covered under the
MBTA (MBTA 2020). Permits from the foreign country
approving the use of BBL-issued bands in that country
must rst have been obtained and submitted to the BBL
with a special circumstances request. An example
would be a researcher wanting to conduct a study of
Ruby-throated Hummingbirds (Archilochus colubris)
and band them while in their wintering range in Central
America. The researcher must rst receive permit ap-
proval from the country in Central America to use the
BBL-issued hummingbird bands, and then the BBL
would consider granting approval. Further details can
be found on the BBL website (https://www.usgs.gov/
centers/pwrc/science/banding-foreign-countries?qt-
science_center_objects=0#qt-science_center_objects).
The BBL issues master permits or sub-permits
depending on the study proposal(s), an investigator’s
level of experience, and requested activities. Permits
issued will specify: states in which the bander can work;
hummingbird species with which the bander can work;
blood, feather, and/or swab samples that are allowed to
be collected; and capture techniques allowed for ob-
taining hummingbirds. All of these specications will
depend on the experience and training of the individual.
When a eld crew is conducting hummingbird-related
activities, at least one crew member is required to have a
BBL master permit or sub-permit. Other team members
can legally capture, handle, band, or mark humming-
birds without an individual permit but only under the
direct supervision of a BBL-permitted bander. If eld
team members are operating independently for any
activities (e.g., capturing, handling, or banding), each
member will need to be issued a BBL sub-permit. To
obtain a master banding permit, a completed application
must be submitted to the BBL permit oce (https://
www.usgs.gov/centers/pwrc/science/permit-appli-
cation-instructions?qt-science_center_objects=0#qt-
science_center_objects). Requests for sub-permittees
need to be made by the master bander and submitted to
the BBL permit oce (https://www.usgs.gov/centers/
pwrc/science/requests-sub-permits?qt-science_cen-
ter_objects=0#qt-science_center_objects).
To be granted a BBL permit, an individual must
provide evidence of their knowledge and skills used
to capture, handle, band, and/or mark hummingbirds.
Individuals initially are trained under the guidance and
supervision of a BBL-permitted hummingbird bander.
Trainees must also complete and pass a training course
in hummingbird handling and banding techniques,
including the process used to properly create hum-
mingbird bands. Additional information can be found
at https://www.pwrc.usgs.gov/BBL/homepage/spe-
cies_auth.cfm. Such courses provide an independent
assessment of a bander’s knowledge and skills and
must be completed before a request is submitted to
the BBL to add hummingbird banding authorizations
to a pre-existing BBL permit for other avian species.
Increasingly, graduate students are conducting research
projects that require BBL permits; however, in some
cases, faculty advisers have very limited or no banding
experience. The BBL will normally recommend that
these students rst train under the supervision of an ex-
perienced and BBL-permitted passerine bander before
starting their independent work with hummingbirds
(B. Peterjohn, pers. comm., 28 June 2020). In addi-
tion, they are required to complete the aforementioned
training on hummingbird-specic banding before they
can begin their eld activities. Because most of these
training activities occur only during the summer, gradu-
ate students should plan accordingly. Once graduate
students can demonstrate competence and obtain ref-
erences from BBL-permitted banders, the BBL will
consider issuing master BBL permits or sub-permits
with “limited” authorization(s) tailored to the specic
activities of the research project(s). Individuals wishing
to attend a hummingbird banding training course should
contact the BBL for more information, recognizing that
these training activities are very limited.
Depending on the scenario, trapping, mist netting,
and/or marking hummingbirds in the US requires either
a BBL permit or a USFWS Migratory Bird Scientic
Collecting permit. In addition, nearly every state has
an agency with its own permitting requirements. Pro-
spective researchers must submit an application to
the appropriate federal and state wildlife agencies. In
most cases, the federal permit(s) need(s) to be obtained
before a state agency will consider a permit request.
A BBL permit is needed for banding or marking hum-
mingbirds, and authorized capture techniques would be
included. For capturing and handling hummingbirds
where banding or marking will not occur, a USFWS
permit is required. Requirements for obtaining USFWS
permits are discussed in the following subsection. A
BBL permit also is required for anyone placing a federal
bird band or auxiliary marker (e.g., a color band, radio-
frequency identication tags, or geolocators) on a bird
that will be released into their natural habitat. Investi-
gators marking a free-ranging bird (e.g., feather dying,
applying paint, or notching feathers) without placing a
federal bird band or an auxiliary marker should contact
the BBL for guidance on permit requirements.
Birds that are hatched in captivity or caught in
the wild with the intent of being permanently housed
in captivity should not be banded with a BBL-issued
band (B. Peterjohn, pers. comm., 28 June 2020). In
the case of a banded free-ranging bird that is converted
to being permanently held in captivity, the BBL band
should be removed if removal can be performed safely.
Otherwise, the BBL band can remain on the bird. If
a hummingbird is hatched, raised, and retained in
captivity and needs to be individually identied for
sampling purposes, BBL federally issued bands cannot
be used. One option is to contact the BBL and arrange
for bands to be made specically for the captive bird
population using a dierent letter prex than those used
for free-ranging birds (B. Peterjohn, pers. comm., 28
June 2020).
BBL permit authorization to band and sample
rehabilitated birds also is allowed. Similar to master
or sub-permittee requirements, experience identifying,
aging, and sexing the requested hummingbird species
must be documented. However, documenting capture
technique experience is not required because birds will
already be in captivity at the time of banding. Because
hummingbirds require specialized bands, the applicant
also must have experience with hummingbird banding
protocols. The BBL requires submission of a project
proposal that details the scientic or conservation
merit for banding and sampling rehabilitated birds
(https://www.usgs.gov/centers/pwrc/science/banding-
rehabilitated-birds?qt-science_center_objects=0#qt-
science_center_objects).
Individuals banding or marking hummingbirds
also can apply to collect blood, feather, and/or swab
samples, and such authorizations will be included on
their BBL permit. Anyone wishing to collect blood
samples from hummingbirds must receive training in
blood sampling techniques before the BBL will ap-
prove that authorization on a banding permit. Because
specialized techniques are required to obtain blood
samples from hummingbirds, experience obtaining
blood samples from other birds is not necessarily ap-
plicable to hummingbirds. Training in hummingbird
blood sampling techniques is currently very limited,
and the BBL should be contacted for additional infor-
mation (https://www.usgs.gov/centers/pwrc/science/
blood-sampling?qt-science_center_objects=0#qt-sci-
ence_center_objects and Bruce Peterjohn, pers. comm.,
28 June 2020). Although swab sampling authorizations
are issued by the BBL, such requests require specic
details, such as the size of the swab to be used and
the anatomic site to be swabbed. Because there are
no published methods for safely taking swab samples
from the oral cavity or cloaca of live hummingbirds,
swab authorization requests are likely to be viewed as
experimental procedures. Such requests must be ac-
companied with detailed descriptions of the proposed
sampling protocol(s) before the BBL will consider
approval (B. Peterjohn, pers. comm., 28 June 2020).
For a bird to be banded and/or marked and re-
strained and/or held in connement for collection of
any other type of biological sample (other than blood,
feather, and/or swab samples), both BBL and USFWS
Migratory Bird Scientic Collecting permits are re-
quired (see https://www.fws.gov/migratorybirds/pdf/
policies-and-regulations/3-200-7FAQ.pdf). If samples
authorized on a USFWS Migratory Bird Scientic
Collecting permit (e.g., urine, fecal, or lesion scrap-
ing samples) are collected, and the bird is banded and
trapped/released during the event that had the sole
purpose of urine, fecal, or lesion scraping sampling,
then the bird should be reported as banded under the
BBL permit and reported as trapped and released on
the USFWS scientic collecting annual report form
(3-202-1; https://www.fws.gov/forms/3-202-1.pdf) .
BBL permittees must adhere to the BBL requirement
that healthy birds be released immediately following
completion of banding, marking, and blood, feather,
and/or swab collection. Birds cannot be restrained,
held, or conned for any activities outside of band-
ing, marking, or obtaining blood, feather, and/or swab
samples under the BBL permit unless they have a
USFWS Migratory Bird Scientic Collecting permit
that allows further containment (50 CFR § 21.23; B.
Peterjohn, pers. comm., 28 June 2020). Cloacal excre-
ment samples can be collected opportunistically (e.g.,
from a bird holding bag) while a hummingbird is being
processed (i.e., banded, bled, or feather sampled) under
a BBL permit. However, hummingbird processing must
be performed in a timely fashion and birds retained for
reasonable lengths of time (i.e., no longer than 15 min).
Containment should not be extended to advantage op-
portunistic sampling of cloacal excrement.
In the case of sick or injured hummingbirds, a
BBL permit authorizes permittees to hold such birds
in captivity for up to 24 h, but only for the purpose of
ensuring the safety and well-being of the bird and long
enough for the bird to recover or be transported to a
wildlife rehabilitator (B. Peterjohn, pers. comm., 28
June 2020). This provision does not allow an investi-
gator to opportunistically collect samples (e.g., urine,
feces) during containment, because the focus should be
on supporting the bird and minimizing disturbance prior
to timely release or transfer to a wildlife rehabilitator.
If bird mortality occurs during banding, mark-
ing, and/or blood or feather sampling, the BBL permit
serves as a salvage permit allowing dead birds to be kept
up to six months. Salvage activities unrelated to bird
banding, marking, or authorized BBL sample collection
procedures must be covered under a USFWS Migratory
Bird Scientic Collecting permit (that includes salvage
authority) and likely under state permits as well.
All permitting requirements described herein re-
ect BBL policies at the time of publication; however,
these requirements could change. Thus, researchers
should always consult the BBL website for the most up-
to-date banding permit requirements. Further details
regarding BBL permit requirements can be found on the
general permit information section of the BBL website
(https://www.usgs.gov/centers/pwrc/science/general-
permit-information?qt-science_center_objects=0#qt-
science_center_objects).
A USFWS Migratory Bird Scien-
tic Collecting permit authorizes qualied individuals
to collect (live, lethal, and/or salvage), transport, or
possess migratory birds, their parts, nests, or eggs for
scientic research or educational purposes (50 CFR §
21.23). USFWS Migratory Bird Scientic Collecting
permits also may authorize importing or exporting
of birds and/or bird samples into or out of the United
States; however, an investigator must request to add
this condition to their permit (https://www.fws.gov/mi-
gratorybirds/pdf/policies-and-regulations/3-200-7FAQ.
pdf).
A scientic collecting permit from a USFWS
Migratory Bird Permit Oce (https://www.fws.gov/
forms/3-200-7.pdf) is required if live birds will be:
(1) captured, restrained and/or contained and released;
(2) sacriced (lethal take); or (3) captured from a
free-ranging environment then held in captivity. If a
hummingbird is captured for blood, feather, or cloa-
cal/oral swab sample collection and not banded and/
or marked, then a BBL permit is not required; rather, a
USFWS Migratory Bird Scientic Collecting permit is
required and likely a state permit as well. In addition, a
USFWS Migratory Bird Scientic Collecting permit is
necessary for the capture, restraint, and/or retention of
a hummingbird for the purposes of urine and/or fecal
sample collection. Even though collection of urine and
fecal material do not fall under the purview of either the
BBL or USFWS Migratory Bird permits, the capture,
restraint, and/or retention activities do. If urine and fe-
cal samples are collected after the urine or feces exit the
cloaca and the hummingbird producing the excrement
is not captured, restrained, or contained to obtain the
samples, BBL or USFWS Migratory Bird permits are
not required; however, a state wildlife agency permit
might be necessary.
When applying for a USFWS Migratory Bird
Scientic Collecting permit, the investigator needs
to declare where live birds, samples, or specimens
obtained by lethal collection will be housed or stored.
Carcasses or any remaining materials from a specimen
must be donated to a public scientic or educational
institution upon conclusion of the research project. It is
recommended that the investigator work with a regional
or state museum to establish vouchering protocols that
can be included in permit applications.
USFWS Migratory Bird Scientic Collecting
permits include salvage authorization. Salvaged
specimens can be a bird and/or bird components (i.e.,
feathers, wing, skeleton) opportunistically found in the
wild, birds that have died at rehabilitation centers, and/
or bird specimens and/or bird components that are being
donated by a researcher after completion of a scientic
project. Salvaging is dened as collecting a deceased
bird when the collecting individual has no involvement
in the death of the bird. Salvaged specimens can be
accepted by a curator or researcher at an accredited
museum or other authorized institution. Salvage for
other purposes may be authorized under a USFWS
Special Purpose – Salvage permit (50 CFR 21.27; Form
3-200-10a; https://www.fws.gov/forms/3-200-10a.pdf) .
A USFWS Migratory Bird Scientic Collecting, Spe-
cial Purpose – Salvage, or Special Purpose Possession
for Education permit is needed to salvage nests, eggs
(viable or non-viable), or dead birds that the researcher
had no part in euthanizing or lethally collecting. The
type of permit depends on the type and purpose of the
activity. Additional information about Special Purpose
Possession for Education permits (Live and/or Dead
Possession: Form 3-200-10c) and Special Purpose
Salvage permits (Form 3-200-10a) can be found in
the Frequently Asked Questions associated with the
permit applications on the USFWS Migratory Bird
Program website (https://www.fws.gov/birds/policies-
and-regulations/permits/need-a-permit.php).
Public scientic or educational institutions can
accept migratory bird specimens for research and
educational use that were lawfully collected, and
those institutions do not need a permit to possess these
specimens (50 CFR § 21.12(b)(1)). The denition
of “public” can be found in 50 CFR § 10.12 and is
as follows: “Public as used in referring to museums,
zoological parks, and scientic or educational institu-
tions, refers to such as are open to the general public
and are either established, maintained, and operated
as a governmental service or are privately endowed
and organized but not operated for prot.” Private
scientic and educational institutions also can accept
and possess migratory bird specimens that were law-
fully acquired, without themselves needing a USFWS
Migratory Bird Scientic Collecting permit; however,
a Special Purpose Possession for Education permit
would be necessary.
To obtain a USFWS Migratory Bird Scientic
Collecting permit, applications must be submitted to the
USFWS Migratory Bird Permit Oce that oversees ac-
tivity in the geographic area where the researcher lives
(https://www.fws.gov/birds/policies-and-regulations/
permits/regional-permit-contacts.php), not where the
research is to be performed. For USFWS permits, if
the permit renewal application is postmarked 30 days
before the current permit expires, permittees may
continue activities authorized by their permits until
the USFWS has acted on the renewal request (50 CFR
§ 13.22). New activities will require approval. If the
deadline is missed and the permit expires, the permittee
will be required to submit a new application rather than
submitting the signed renewal letter that the Migratory
10
Bird Permit Oces currently provides to permittees.
The USFWS is currently in the process of developing
an online permitting system that is expected to be avail-
able by the end of September 2020. This new online
system will allow permit applications to be submitted
electronically. Until the online permitting system is
available, the Migratory Bird Scientic Collecting
permit application (Form 3-200-7) is available on the
USFWS website (https://www.fws.gov/forms/3-200-7.
pdf) and may be printed and mailed to the USFWS, Mi-
gratory Bird Permit Oce. Table 2 provides expanded
explanations of required information needed for a Mi-
gratory Bird Scientic Collecting Permit Application
(Form 3-200-7; http://www.fws.gov/forms/3-200-7.
pdf) or for a renewal application for a Migratory Bird
Scientic Collecting Permit. These items are not an
exhaustive list of application components but provide
guidance for expectations.
Scientic collecting per-
mits or research permits are required by state agencies
regulating wildlife-related activities. The Ornithologi-
cal Council website has a section that summarizes state
permitting requirements and provides the names for the
appropriate contact in each state (https://birdnet.org/
info-for-ornithologists/permits/states/). State permits
must accompany the appropriate federal permit(s).
Researchers should understand local conservation
and protection laws governing hummingbirds at their
study sites.
Any
scientic, banding, and/or marking activities conducted
on federal, state, or public lands may require a permit
from the agency administering those lands. Examples
of federal lands that might include special use permits
are USFWS National Wildlife Refuges, US Forest
Service, or Bureau of Land Management specially
designated areas. Public properties administered by
county or regional authorities and land maintained by
non-governmental organizations (e.g., Nature Con-
servancy reserves) also may have special permitting
requirements. Each agency/land management orga-
nization should be contacted regarding their specic
permitting requirements. The Ornithological Council
provides information and links to federal agencies that
require permits when a researcher is working on feder-
ally owned lands (https://birdnet.org/info-for-ornithol-
ogists/permits-us-federal/). The authors are unaware of
a similar resource for permitting requirements by state
agencies. Any work conducted on private lands must
be with landowner permission, and written consent is
highly recommended.
States of America
Proper identication of the species, age, and sex
of a hummingbird is critical for scientic studies. The
following guides have been found to be most useful
for providing identication data:
to North American Birds Part 1 (Pyle 1997); A Field
Guide to Hummingbirds of North America (Williamson
2001); Hummingbirds of North America: The Photo-
graphic Guide (Howell 2002); and the North American
Banders Manual for Banding Hummingbirds (Russell
and Russell 2019). Because identication of hum-
mingbird species, age, and sex can be challenging, it
is helpful for researchers to carefully study reference
guides and practice with museum study skins before
working with live birds. This will help minimize
restraint time and live bird energy expenditure and is
especially important for novice investigators or those
unfamiliar with species in a newly studied geographic
area. Voucher specimens are most likely to be found
at natural history museums, where access policies vary.
Researchers undergoing training to become hum-
mingbird banders should learn traits of the research
species and/or traits of the suite of species in the geo-
graphic area where they will be working. Research-
ers also should become familiar with traits of species
within the same genus, even if these species are not
commonly found in the geographic area of interest,
so that unusual encounters can be properly identied.
BBL permittees working with unfamiliar hummingbird
species also should be encouraged to receive training
from permittees that are experienced with the species
Table 2. Required or optional study proposal components for a Migratory Bird Scientic Collecting permit. The items below are expanded explanations
of information required for a Migratory Bird Scientic Collecting permit (Migratory Bird Scientic Collecting Permit Application; Form 3-200-7 found
online at http://www.fws.gov/forms/3-200-7.pdf) and Migratory Bird Scientic Collecting permit renewal letters (mailed or emailed by the Migratory
Bird Permit Oce). These components also need to be provided with permit amendment requests. This is not an exhaustive list of required application
components, and investigators are encouraged to fully review Form 3-200-7 to evaluate which components might be necessary for their studies.
Component
Required
or Optional Description Additional Comments
Collecting Activity
Table
Required One line should be completed for each species
in each location. The state, county, season or
months of collection, and numbers for each type
of collection should also be specied. In addition,
information should include whether individuals will
be adult males, adult females, juveniles, nestlings,
and/or eggs.
It is strongly recommended to include a paragraph explaining the
information in the Collecting Activity Table and justication of
numbers, age classes, locations, and time of year.
Proposal and
justication
Required The proposal should include the following items:
Justication for the proposed research (background,
statement of problem, and hypotheses)
Study site selection and description
Species to be studied
Time period of collection
Field and laboratory methods
Expertise of researchers to be conducting this
research
Literature cited
This section should be several pages long. Investigators may copy and
paste from an existing grant proposal or may write a brief summary
and include an entire grant proposal as a supporting document.
Regardless, a Collecting Activity Table is still required. Statistical
justication for the numbers of individuals requested is strongly
encouraged. Additional specic justication must be submitted to
collect birds on the most recent Birds of Conservation Concern List,
which is available at http://www.fws.gov/migratorybirds/.
Description of
background
and expertise of
researchers in
conducting the
proposed activities
Required If this information is already contained in the
investigator’s permit le in the Migratory Bird
Permit Oce, this component is not necessary.
A curriculum vitae is helpful; however, a paragraph highlighting the
investigator’s expertise in the requested eld of scientic work and
eld/laboratory methods also is necessary.
Justication for
number of birds
requested
Optional It is preferable that this be a statistical justication
such as a power analysis demonstrating the sample
size needed for statistical signicance.
This is especially important for bird species of conservation concern if
the investigator is lethally taking individual birds, trapping and holding
individual birds for longer than 2–3 hours, or requesting a large
number of individual birds for trap and release.
12
Component
Required
or Optional Description Additional Comments
Explanation of
population level
eects
Optional This information should detail the population-
level eects (i.e., the proportion of the local
population to be removed and citation[s] supporting
the population estimate) and how the researcher
proposes to avoid/minimize impacts.
Examples of minimization measures include the following:
Utilize existing museum specimens
Collect dead specimens from wildlife rehabilitation centers
Distribute lethal-take of birds geographically and temporally
Collect specimens outside the breeding season or toward the end of the
breeding season when young of the year are independent
Avoid lethal take of breeding birds
Utilize blood rather than tissue for genetic analysis
Coordinate project with other research projects collecting the same or
similar species to share samples
Explanation of
conservation benet
Optional This explanation would include information
regarding the conservation benets of the proposed
project.
The explanation should indicate that collected specimens would be
deposited in a museum. Working with local researchers will facilitate
identication of those most suitable and willing to receive materials.
Table 2. (cont.)
in question. All of these eorts will help minimize
bird handling time, diculties with identication in
the eld, and ensure proper source metadata pertaining
to collected samples.
If a BBL permittee is uncertain of a bird’s iden-
tication, guidance from the BBL is not to band the
bird. The metadata for the sample(s) should reect
the uncertainty of the species, sex, and/or age of the
bird. If a BBL permittee is condent of the species
identication but not the bird’s age and/or sex, multiple
photographs of the bird need to be taken to record all
key morphologic features. Important characteristics
to photographically document include spread wing
feathers and tail feathers, beak morphology, and general
plumage. The BBL permittee should then consult with
experienced hummingbird banders to reach a consen-
sus. For US and Canadian BBL permittees (master
and sub-permittees), there is a listserv (Humband)
available for requesting expert opinions for challenging
identication cases.
Some biologists use auditory traits to help iden-
tify hummingbird species (Clark and Mistick 2018b;
Clark et al. 2018b), especially for male birds. However,
it is important to ensure that the auditory traits are
directly associated with the bird being captured, identi-
ed, and sampled. The Merlin Bird ID mobile device
app (Cornell Lab of Ornithology; https://merlin.al-
laboutbirds.org/download/) is a useful resource but does
not contain all hummingbird vocalizations or sonations
(non-vocal communicative sounds made from wing or
tail feathers; Feo and Clark 2010). Xeno-canto (https://
www.xeno-canto.org/), a website dedicated to sharing
bird sounds collected worldwide, is a useful resource
for recordings of vocalizations from temperate and
neotropical hummingbirds. Another useful reference
for North American species is Peterson’s Field Guide to
(Pieplow 2019).
A variety of traps have been used to capture hum-
mingbirds. The North American Bander’s Manual for
Banding Hummingbirds (Russell and Russell 2019) de-
scribes several types of traps. The use of traps to safely
capture birds depends on the individual’s experience
and skill level. When trapping, certain precautions must
be taken to ensure the safety not only of hummingbirds
but also of researchers and bystanders. Drop-curtain
and drop-door traps require researchers to sit remotely
while the curtain or trap door is typically controlled by
a monolament line. This line should be clearly marked
with a material, such as vinyl agging tape, to prevent
tripping or other hazardous conditions, especially when
working in a public location. When using traps, the
best practice is to wait until birds are feeding or at least
perched on the feeder within the trap before closing the
trap door or dropping the net. If a trap is set properly
(i.e., the trap is not raised too high and/or the feeder
is placed as far as possible from the door), chances of
harming hummingbirds are minimized when triggering
the trap or closing the door. Regardless of safe-trapping
practices, to further minimize bird injury, all compo-
nents of the trap should be lightweight and the trap
curtain or door should not close with excessive force.
When capturing and processing hummingbirds,
researchers need to consider the time of day, tem-
perature, wind speed, precipitation, and the number
of trained individuals available to extract and process
birds. In order to minimize situations that predispose
birds to hypoglycemia, hypothermia, and respiratory
and/or cardiac distress and/or arrest, researchers need
to establish written protocols dening conditions during
which netting and trapping should be avoided or cur-
tailed. Every set of circumstances is unique; therefore,
a bird’s condition must be constantly monitored and
activities reduced or aborted as necessary.
Special precautions should be taken when trap-
ping or mist netting and sampling at rst light, because
some hummingbirds may be ending a prolonged fast
and may require time to re-establish energy reserves.
Similarly, care should be taken prior to sunset when
some hummingbirds will be building up their fuel
resources prior to entering torpor. Disturbance prior
to sunset could negatively impact a bird’s ability to sur-
vive through the night. This is especially important dur-
ing winter months and in geographic locations where
hummingbirds face challenging weather conditions.
From a welfare perspective, one universal guideline is
that hummingbirds should not be captured or processed
within 30 min after sunrise or before sunset. This time
guidance might need to be extended depending on the
situation.
In general, a researcher can consider a bird caught
at a feeder to be seeking food and the bird might have
had a small meal immediately prior to capture. In con-
trast, if a bird is caught in a mist net, there is no way to
know when the bird last ate; therefore, the researcher
should assume the bird could be close to an energy
crisis. During the breeding season, hummingbirds may
be caring for dependent young and have high energy
needs; therefore, birds should be extracted from traps
and netting and processed as quickly as possible. The
BBL does not have specic guidelines for checking
mist nets, but under most circumstances the authors
and B. Peterjohn (pers. comm., 28 June 2020) recom-
mend checking at least every 15 min. However, in
temperatures above 27° C (80° F) or below 10° C (50°
F), mist net checks are recommended every 10 min or
more often. To ensure bird safety, regular monitoring
protocols should be developed for all trapping activi-
ties, even when traps are open. Open drop-net traps
can be problematic because even though the trap is
“open,” hummingbirds might y to the top of the trap
and not nd an escape route. Capture stations must be
prepared to treat birds compromised by health issues
or a hypoglycemic/hypothermic event, which includes
providing supplemental heat and small volumes of
sugar water.
Research protocols need to describe how many
feeders to cover and/or remove to concentrate hum-
mingbirds at a trapping station. When setting up a
hummingbird capture station, researchers should assess
food sources available within a 1.6–3.2 km radius of
the research site. For isolated feeding stations (i.e.,
those with no other feeders available within 3.2 km)
at a time of year when owers and insects are scarce
(e.g., California during the dry season), decreased en-
ergy sources and increased competition at remaining
feeders can result in birds becoming exhausted. In such
conditions, the trapping station should be operated for
shorter periods of time (approximately 20–40 min),
after which the original number of feeders should be
returned to their original hanging locations or uncov-
ered. This protocol can be followed intermittently as
needed. Researchers should estimate time intervals
for returning the feeders by closely observing feeding
activity and the status of birds at the banding table.
The less food availability, the shorter the continuous
trapping time should be. For circumstances where each
14
bird is being held for an extended period of time (e.g.,
for blood and feather sampling and/or tag placement),
feeders should be returned to their hanging locations
and/or uncovered while each bird is being processed
and then removed to recapture the next bird.
In situations where additional feeders surround
the research site (i.e., within a 1.6–3.2 km radius from
the trapping location) and/or natural foods are plentiful,
hummingbirds will go elsewhere to feed so that reduc-
ing the number of feeders to facilitate trapping should
not have a negative impact. For locations hosting large
numbers of hummingbirds (i.e., >100 birds), one to
two uncovered feeders can be left hanging in addition
to the feeder(s) at the trapping station(s). Because
competition at the uncovered feeders can be relatively
intense, some hummingbirds may look for feeders at
other locations; thus, fewer birds will be available to
enter the traps. If this scenario involving plentiful
feeders applies to a private residence research location,
property owners need to be reassured that birds will not
starve if the majority of feeders are removed/covered
for approximately 20 min.
Acceptable weather conditions for capturing
hummingbirds will depend on capture techniques. In
general, netting and trapping should be avoided during
inclement weather (especially involving precipitation)
or high wind conditions because such conditions can
compromise a bird’s ability to thermoregulate and will
increase energy demands. The range of cold tempera-
tures conducive to working with hummingbirds will
depend on other variables, such as humidity and wind.
Banding is possible during colder temperatures (-4° C
[25° F] to 4° C [39° F]) (B. Peterjohn pers. comm., 28
June 2020) if the bird is removed immediately from
the trap and processed within 2–4 min of being cap-
tured. Banding also can occur when temperatures are
below 0° C (32° F) if birds are brought into a warmer
environment for a short period of time. Capturing and
sampling hummingbirds during extreme heat also can
be problematic. Once captured, birds must be pro-
cessed quickly to avoid overheating. If captured birds
are brought indoors for processing, investigators should
work only in rooms without ceiling fans and with ceil-
ings low enough to allow escaped birds to be captured
with a net. A long-handled net is a necessary piece of
equipment when working indoors. The net should be
similar in weight and material to a buttery net to avoid
crushing the bird when capturing it. Prior to initiating
work indoors, covering windows and glass doors should
be considered to prevent an escaped bird from being
injured. Some investigators prefer uncovered windows
or glass doors to facilitate capture as the birds are at-
tracted to them as a possible escape route. Whether
to cover or not cover windows and doors depends on
the acceleration speed that a bird could attain before
hitting the structure. Opening a door that leads to the
outside might help with guiding a bird outdoors if the
indoor ceilings are not too high.
Hummingbirds
With all captured hummingbirds, the researcher
needs to minimize detaining and handling time; thus,
being ecient is essential. Individuals who have not
handled hummingbirds previously can initially learn
bird handling skills and gain condence using captive
birds of similar size and weight (e.g., zebra nches).
Another way of obtaining experience is to volunteer
and train at a passerine banding station. At the time
this manuscript was written, the general policy of the
BBL was that handling several hundred passerines
under the supervision of an experienced bander was
required before a request for a hummingbird band-
ing permit would be considered (B. Peterjohn, pers.
comm., 28 June 2020).
Obtaining samples from hummingbirds requires
the handler to pay attention to the sampling task at hand
while simultaneously monitoring the bird’s condition.
When restraining hummingbirds, handlers should mon-
itor birds for signs of compromise or stress. Compro-
mised hummingbirds may continually vocalize, gape,
repeatedly close their eyes, maintain their feathers in
an erect position, or spontaneously shed large numbers
of contour (body) or tail feathers. Birds experiencing
an energy crisis may appear to fall asleep or exhibit
minimal response to stimulation. If these conditions
occur, the bird should immediately be placed in a bird
bag or released, depending on the researcher’s experi-
ence level. When in doubt, the researcher should err on
the side of caution and release the bird. If the bird is not
released, restraint can be attempted again after the bird
has rested, but this should be done by the person on the
team with the most handling experience. Depending on
the bird’s condition, oering sugar water or warming
the bird prior to release (if hypothermia is of concern)
may be helpful. If a bird shows signs of having an
energy crisis during processing—and to minimize the
risk of this happening to other birds—feeders should
be restored to their original numbers and locations for
a period of time so that the remaining population of
hummingbirds can replenish their energy stores.
The most critical thing to be aware of when re-
straining any bird is to avoid putting pressure on the
thoracic region, which will compromise the animal’s
respiration. Birds do not have a diaphragm; instead,
they use their intercostal muscles to move the keel up
and down, which imitates a bellows and moves air in
and out of the respiratory tract (i.e., air sacs, airways,
and lungs). Pressure on the keel can prevent the chest
area from rising and falling and can result in suoca-
tion. Therefore, proper restraint entails holding a bird
with gentle pressure applied at its sides and not from
front to back.
When handling a hummingbird, it also is impor-
tant to keep the bird upright and not allow the head to
go below the plane of the tail. This positioning will
prevent uid in the abdominal region from ooding the
lungs if the bird has liver disease. For female birds,
upright positioning will minimize air sac compression
by reproductive organs or a developing egg. Avoiding
pressure in the crop region will minimize regurgitation
and aspiration (passage of ingesta into the respiratory
tract) of sugar water, especially if the bird was caught
at a feeder. If a bird starts to regurgitate uid, it should
be immediately placed on a at surface, restraint should
cease, and the bird allowed to clear the uid itself. In
severe cases, the bird might need to be placed in a eld
intensive care chamber (see section 2.5) to recover.
The handler should never attempt to clear uid or turn
the bird upside down to let the uid drain; a conscious,
unrestrained, and normally positioned hummingbird
will be better able to clear the uid on its own.
Several methods for handling and restraining
hummingbirds have been described previously (Russell
and Russell 2019), and best safe practices will depend
on the handler’s skill level and hand conformation and
size. Depending on the handler’s hand conrmation
and size, it is possible to handle larger hummingbird
species using published small passerine restraint tech-
niques (Russell and Russell 2019). For smaller-sized
hummingbird species (less than 6 g), the hummingbird
bander's hold can be used where the bird is restrained
in one hand, with hyperexion of the third nger into
an upside down ‘V’ shape which creates a crevice
that allows for cradling the bird’s head and neck (Fig.
1 A and B), and the remaining ngers cup the bird’s
body. The hummingbird bander's hold decreases the
chance for cervical dislocation that can occur with the
“traditional” bander’s grip used for passerines. With
the “traditional” bander’s grip used for passerines, the
second and third ngers are extended (straightened) and
used to restrain the head. However, if this grip is used
with small-sized hummingbirds (less than 6 g), exces-
sive head and neck traction can result in cervical dis-
location. Using the hummingbird bander's hold, where
the third nger is bent (hyperexed), the handler can
restrain the head without too much traction, visually/
tactilely monitor the bird’s respiratory/activity status,
observe the eyes for closure, and minimally compress
the chest region while creating an environment where
the bird feels enclosed and is less likely to struggle.
During restraint, the bird is positioned upright. The
bird’s safety and the optimal technique depends on the
handler’s hand size, nger conformation, nger dexter-
ity (i.e., how much the handler can hyperex the third
nger), and positioning of other ngers relative to the
base of the hand for cupping the bird. To maximize
dexterity, prospective handlers should practice third
nger hyperexion without a bird in hand.
For containment and transport, hummingbirds
can be placed in a soft, well-ventilated bag that pre-
vents excessive movement, which can deplete a bird’s
energy stores. Instructions for making safe, eective
restraint bags from seine material are found in the North
American Hummingbird Banders Manual (Russell and
Russell 2019). Holes in the seine mesh should be large
enough to provide adequate air ow but suciently
small to avoid catching the tips of a bird’s primary
feathers, which can result in shoulder dislocation or
damage to ight feathers. Using the seine material
bag, the researcher can see and position a bird to gain
access to a leg for banding or toenail for blood sampling
(see section 4.2).
16
A B
Figure 1. llustration of lateral (A) and dorsal (B) views of the hummingbird bander’s hold. Note that the third nger
is bent into a distinct “V” shape and the index nger is slightly exed and positioned behind the bird’s head, while the
fourth and fth ngers are curled into the base of the hand. The bird is contained in a “cupped hand”, avoiding front
to back pressure, so it can breathe. Hyperexion (bending) of the third nger cradles the head and reduces the chance
for cervical dislocation. The safety of using this restraint technique will depend on the handler’s nger dexterity and
conrmation and hand size.
Weather permitting, hummingbirds can be con-
tained in a bird bag. Best practice is to oer all birds
a limited meal before they are placed in a bird bag;
however, they should not be allowed to overfeed.
Depending on the situation (e.g., whether or not a
bird drank sugar water prior to being placed in the
bag, the amount of sugar water consumed, the ambi-
ent temperature, and the amount of movement in the
bag), birds should be oered sugar water every 15–45
min to prevent hypoglycemia. Bags containing hum-
mingbirds should be placed in a safe, shaded location
to prevent birds from becoming overheated, and the
bags should always be suspended and not laid on at
surfaces, including tables and oors. A bird in a bird
bag should never be placed on the ground. In addi-
tion to avoiding inadvertent harm, hanging bird bags
allows for visually monitoring birds between capture
and processing. Various stands are used for hanging
hummingbirds contained in holding bags. For sta-
tions where sampling is the major focus (as opposed
to banding/capture/recapture), a small stand made of
polyvinyl chloride (PVC) pipe and ttings can be used
(Fig. 2). The bag must be hung high enough so that the
bird is suspended and not allowed to come in contact
with a at surface where it will struggle and potentially
harm itself or damage its feathers. The damage that a
struggling bird on a at surface might inict on itself
may not be visible, but continuous struggle against a
rm surface can result in muscle injury. The authors
do not hang bags by the drawstring (especially when
using short stands) but rather use the holes in the seine
bag to hang on the hook.
If a researcher needs to retain a hummingbird
in captivity for an extended period of time (i.e., >30
min) for sampling purposes, the bird can be placed in a
cage with food available ad libitum (Russell and Rus-
sell 2019). Similar to recommendations for holding
birds in bags, enclosures should be placed in a shaded
area with no direct sunlight to prevent overheating
and should be visible by a team member at all times.
Commercial buttery enclosures with mesh on at least
one side aord good ventilation and oer one option
for retaining birds temporarily. This is particularly
Figure 2. Illustration of a bird bag holding stand made
of polyvinyl chloride pipe and ttings. This stand can be
used for suspending hummingbirds contained in holding
bags at the eld work-station until processed. Note that
a hole in the seine mesh (as opposed to the cord of the
drawstring) is used for suspending the bag from the hook
so that the entire holding bag is suspended.
helpful if the research activity requires that the bird
not be in a bird holding bag or if the bird needs to be
acclimated to captivity for a captive study. However,
buttery cages are not a replacement for bird bags,
because birds will y against the sides of the cage and
damage their feathers or continuously y and become
exhausted. Larger enclosures can be made with PVC
tubing and small-gauge netting. When caged, a bird
must be observed drinking from the oered food source
before it is left unobserved for any extended period. If
the bird is not seen drinking, it should be caught and
fed every 20–30 min until it learns to drink from the
food source.
When providing food to a hummingbird that
may be kept in captivity for an extended period, using
syringes instead of an outdoor hummingbird feeder is
recommended. Syringes are easy to ll and can be dis-
assembled for cleaning. In addition, volume markings
on syringes allow for tracking food consumption and
controlling the amount of hand feeding. Syringes with
a 15 mL or 30 mL capacity (e.g., Becton Dickinsons
oral syringes; https://www.bd.com/en-ca/oerings/
capabilities/syringes-and-needles/oral-and-enteral-
syringes/oral-syringes) are recommended. The syringe
can be suspended in the cage by a exible wire, which
is coiled around the syringe and extended up to form a
hook by which the syringe is secured to the top or side
of the cage. The syringe opening should be pointed at
an angle that allows the bird to easily gain access to
the food while preventing the sugar water from leak-
ing out of the syringe, as might happen if the opening
were pointed straight down. Hummingbirds should be
monitored regularly to ensure that they learn how to
drink from the syringes. To that end, multiple syringes
should be made available to a newly captive bird to
improve its chance of trying one of them. A perch can
also be placed close to one of the syringes to facili-
tate access. If a bird is not observed feeding, then it
should be captured and its beak placed into the syringe
opening. This procedure may be necessary multiple
times until the bird is observed feeding on its own. All
syringe feeders should be cleaned with an appropriate
cleaning solution between each use and changed daily.
For long-term housing of birds, an enhanced diet is
recommended instead of sugar solution alone. Nekton-
Nektar-Plus® nectar concentrate (Hoheneichstraße,
Germany) can be used for adult hummingbirds. The
manufacturer recommends making new formula twice
a day (morning and afternoon) because it spoils easily
in warm environments. The volume of food oered
should meet the energetic and nutritional demands of
the species, age, and sex of the captive bird being held.
For juvenile birds, additional factors should be consid-
ered in conjunction with veterinary advice.
Hummingbirds can be transported in bird bags
either by hand or in a car. As previously described,
the amount of time hummingbirds are held in bird bags
should be limited, the researcher must ensure that the
bird has used a feeding system before transport, and
access to a stable food delivery system throughout
transport is imperative. If a bird is transferred in a car,
the bird bag must be suspended to avoid inadvertent
injuries. Hummingbirds should not be left unattended
in cars in hot environments, and air conditioning should
be used to prevent overheating.
18
Sampling Live Hummingbirds
An understanding of hummingbird anatomy and
physiology is critical to ensuring welfare during han-
dling. For general guidelines, see the North American
Bander’s Manual for Banding Hummingbirds (Russell
and Russell 2019). More specic advice regarding
threats facing hummingbirds during handling, including
organ system specic issues and conditions of specic
concern, is provided below. Figures 3 and 4 illustrate
a hummingbird’s internal anatomy.
A hummingbird gap-
ing or panting during handling usually indicates that the
bird’s breathing is impaired. Gaping can be a result of
excess pressure being applied to the chest region, which
impedes movement of the bird’s keel and restricts gas
exchange and oxygen availability. Gaping or panting
is an emergency situation that can result in hypoxia,
cardiac arrest, and death within as little as 5 sec after
the onset of the clinical signs. Once a bird starts gaping
or panting, the handler must loosen their grip to relieve
compression on the chest region and release the bird
into a holding bag, a container, or the wild. Airway
compromise also can result from regurgitated uid that
is aspirated. Other reasons for gaping include air sac
compression by enlarged organs, fat deposition, and/or
an egg. To reduce the incidence of respiratory distress
during sampling, birds should be maintained in a head-
up position with just enough restraint to prevent escape,
and handling time should be minimized.
—During breeding
season, female birds (hens) must be handled with care.
At the onset of handling, the abdominal area should be
assessed for the presence of an egg. Enlargement of
the reproductive tract displaces other organs and can
compress the air sacs. When evaluating whether a
female bird is gravid, the bird should be restrained in
an upright position to minimize pressure of an egg on
the air sacs and lungs and the birds should be processed
quickly. Gravid birds or birds that appear to have laid
an egg recently (i.e., those with an extremely enlarged
and swollen cloaca) should be released and not sampled
to minimize the time the hen is away from the nest.
Many researchers who have
worked with hummingbirds are familiar with torpor, a
state of decreased metabolic activity, in which all non-
essential functions are reduced and body temperature
may drop to below 10° C (50º F) in North American
species (McKechnie and Lovegrove 2002) and as low
as 3º C (37º F) in Andean species (Wolf et al. 2020).
Torpor is a mechanism for hummingbirds to conserve
energy throughout the night (Ruf and Geiser 2015).
Along with a reduction in body temperature, torpor
is characterized by elevated feathers and lack of a re-
sponse to movement or touch. Many hummingbirds go
into torpor when energetically stressed during emergen-
cies, often caused by low food availability or sudden
change in environmental conditions (Hainsworth et
al. 1977). Increasing evidence suggests that several
hummingbird species use torpor regularly, even when
they have high energy reserves (Krüger et al. 1982).
However, this distinction may depend on the size of
the species, with smaller species employing torpor
more often than larger ones (Shankar et al. 2020b).
This is an important factor to consider when working
with smaller species, because smaller hummingbirds
may go into torpor despite having food available ad
libidum (Krüger et al. 1982), while larger species may
use torpor less but employ it during times of duress
(Hainsworth et al. 1977). When birds are coming out
of torpor, they will respond to touch and stimulation,
stretch their wings (waking behavior), clamp their
feet reexively, and vocalize with a squeaking pitch.
This “keening note” is a unique sound, similar to one
a mouse might make, and not one observed in active
hummingbirds except during extreme distress (Stiles
1982). Although torpor itself is not a life-threatening
condition, special consideration should be given to
these events relative to sampling. Because humming-
birds might undergo torpor at night and come out of
torpor early in the morning, researchers should never
take blood samples from hummingbirds for at least 30
min after sunrise or 30 min prior to dusk.
2.4.4. Hypoglycemia and other compromised
conditions.—Although hummingbirds can use torpor
to conserve energy, hummingbirds in a torpor state
predominately have been documented at night (Calder
1994; Ruf and Geiser 2015). In general, a healthy
Figure 3. Anatomic drawing (lateral view) of soft tissue organs in the coelom
of a hummingbird.
Figure 4. Anatomic drawing (ventro-dorsal
view) of soft tissue organs in the coelom of
a hummingbird.
20
hummingbird should not go into torpor during the day
and in response to handling and sampling. Therefore,
a bird that appears to be going into torpor during han-
dling, restraint, banding, or sampling during the day
is more likely to be hypoglycemic or hypothermic or
experiencing cardiac/respiratory arrest.
Metabolic requirements of hummingbirds limit
how long they can go without feeding; therefore, the
time between capture, sampling, and feeding must be
carefully monitored. Birds that become hypoglycemic,
hypothermic, or are entering another compromised
state may exhibit several warning signs, including
erect feathers and dull and/or drooping eyes, which
can gradually progress to overall unresponsiveness.
Thus, it is important to continuously monitor birds for
these signs during handling and especially after blood
sampling, because the bird’s status can deteriorate
rapidly. If a bird begins to show any of these signs,
processing should be immediately discontinued and
the bird’s condition assessed. If the bird is conscious,
sugar water should be oered. A bird that is extremely
hypoglycemic must be fed sugar water if it is going to
recover. In rare cases, the researcher might have to
carefully open the bird’s beak and deposit a tiny drop
(approximately 2 µl) of concentrated sugar water on the
tip of the tongue, which can be done using tuberculin
or insulin syringes with the needle removed. An eye
dropper can be used as a last resort but this technique
risks spilling sugar water on the bird’s feathers. If the
bird responds by extending and withdrawing its tongue
several times, it is ingesting the sugar water. If the bird
does move its tongue, the researcher should wait 2 min
and repeat the process. Hypoglycemic hummingbirds
can recover very rapidly and voluntarily ingest large
volumes of sugar water once their blood sugar concen-
trations are restored. If the bird recovers, it may be ap-
propriate to resume data collection. If the bird does not
feed or feeds but does not recover, it should be placed
in a eld intensive care chamber (see the following
section), provided supportive care (i.e., supplemental
heat and a quiet environment), and monitored closely.
When working with hummingbirds, it is best
practice to have an intensive care chamber and supple-
mental feeding sources available at study sites. Some
biologists place a compromised hummingbird inside
their clothing and use their own body heat to provide the
necessary supplemental heat. However, depending on
an individual’s clothing (e.g., the material and number
of cloth layers), this practice could reduce ventilation
and expose the bird to an accumulation of carbon
dioxide. In addition, the bird could be inadvertently
crushed. Placing a compromised hummingbird that
has been fed a sugar meal in an intensive care chamber
allows the bird to recover by minimizing exertion that
might occur with continued handling. The intensive
care chamber can be made from a small container with
air holes. A sherman’s bait bucket, which is typically
made of sturdy plastic and contains air holes in the
lid as well as a trap door for easy access to the inside
of the container, can be used for this purpose. A heat
source that does not emit fumes can be placed inside
the intensive care chamber; one recommendation is
a re-useable hand warmer (HotSnapZ®, La Porte,
Indiana), which uses sodium acetate crystallization to
generate exothermic heat. A tightly woven cloth that
will minimize the risk of a bird catching its nails can
then be placed between the bird and heat source. A
small tuberculin or insulin syringe without a needle can
be used to oer sugar water or a balanced electrolyte
solution to help restore energy reserves. As mentioned
previously, an eye dropper or a plastic transfer pipette
is not ideal because it risks spilling sugar water on the
bird’s feathers.
A literature review revealed 46 studies that used
some form of marking to address a specic research
objective. Of these, the majority used leg banding (16)
followed by color marking (19), colored acetate plastic
tags on the leg or back (3), radio-frequency identi-
cation tags (6), radio-telemetry tags (7), and plastic
colored bands (1). Table 3 summarizes these studies.
Note that for any marking methods discussed below,
federal and state permits are required. Depending on
the research objectives and corresponding biological
specimen sampling plan, it may be necessary to mark
hummingbirds to avoid resampling a bird on the same
day and location or re-sampling a bird if the recapture
interval is short. When marking hummingbirds, it is
important to consider the long-term eects on the birds
and to use methods that will have the least impact. If
long-term population monitoring and/or resampling
individual birds is not necessary, a minimally inva-
sive method, such as subtle paint marking, may be the
best option. However, the impact of various marking
methods must be considered carefully. For example,
during the breeding season, paint can compromise the
cryptic appearance of a female on the nest, or clipping
certain feathers might impair the sounds that a male
bird’s feathers make during courtship dives.
Feather clipping is an easy way to mark a bird
without banding and to ensure identication of recap-
tured birds that were sampled. In past studies, 2–3 mm
of the feather tip was clipped, or small V-shaped cuts
were made in the tail feathers or the distal portion of
secondary wing feathers. Cutting primary wing feath-
ers hinders ight and, for some species, might impact a
bird’s ability to make sounds; therefore, it may be more
appropriate to clip other feathers for marking purposes.
Scientic studies have shown that hummingbird tail
Table 3. Summary of methods used to mark hummingbirds as reported in published studies.a
Method Count Permanence Applications References
Banding 16 Years Population biology,
conservation, ecology
Calder III et al. 1983; Miller and Gass
1985; Inouye and McGuire 1991; Mulvihill
et al. 1992; Oniki 1996; Hilton Jr. and
Miller 2003; Wethington and Russell 2003;
Bassett and Cubie 2009; Temeles et al.
2009, 2013; Hurly et al. 2010; Cubie 2014;
Maglianesi et al. 2014b; Supp et al. 2015;
Zenzal and Moore 2016, 2019
Acetate plastic
colored tags on back,
leg or leg band
3 Weeks to years Population biology,
behavior ecology
Stiles and Wolf 1973; Waser and Calder
1975; Kapoor 2012
Plastic colored bands 1 Years Ecology, behavior Temeles and Bishop 2019
Color 19 Days to months Behavior, population
biology, conservation,
ecology
Wolf 1969; Baltosser 1978; Ewald and
Carpenter 1978; Goldsmith and Goldsmith
1979; Stiles and Wolf 1979; Trombulak
1983; Gill 1988; Stiles 1992; Hilton Jr.
and Miller 2003; Temeles et al. 2006;
Clark and Feo 2008, 2010; González and
Ornelas 2009; Feo and Clark 2010; Hurly
et al. 2010; Clark 2011b, 2014; Golo
and Burch 2012; Zenzal and Moore 2016;
Tello‐Ramos et al. 2019
RFID 6 Days (glue applica-
tion) to years (sub-
cutaneous injection)
Behavior, population
biology, conservation,
ecology
Brewer et al. 2011; Hou et al. 2015; Ibarra
et al. 2015; Zenzal and Moore 2016, 2019;
Bandivadekar et al. 2018
Telemetry 7 Days Behavior, conservation,
ecology
Hadley and Betts 2009; Zenzal et al. 2014,
2018; Zenzal and Moore 2016; Hazlehurst
and Karubian 2018; Céspedes et al. 2019;
Pavan et al. 2020
a Literature searched until September 2020.
22
feathers are important during courtship and for com-
munication (Clark et al. 2013a, 2013b, 2018a; Clark
and Mistick 2018a). For example, tail feathers R4 and
R5 (Fig. 5) play an important courtship role in males
of some hummingbird species, so these feathers should
not be clipped during breeding season. Hummingbirds
in the bee clade make sounds with their tail feathers
(Clark and Mistick 2018b), so clipping secondary wing
feathers is a more acceptable alternative. However,
there are exceptions even to clipping secondary feath-
ers, such as in the Archilochus species whose primary/
outer secondary feathers have been modied to produce
sounds. Whichever feather is clipped, one must be
aware that it will remain unchanged until the following
molt, the timing of which varies by species and location.
A variety of dyes, pigments, or paints have been
used to identify previously marked birds (in which case
a single uniform mark can be used) or individual birds
from a distance (by using a unique shape, orientation,
or color combination). This approach is ideal when
an investigator wants to avoid distressing individual
birds that have already been captured, or when the
goal is to observe individual behaviors of a species
with predictable, small-scale spatial movements (e.g.,
territorial or lekking hummingbirds). An advantage
of paint marking is that it can be used in conjunction
with trap cameras or videos because the markings are
Figure 5. Illustration of extended wing (numbered wing feathers; remiges) and ared tail (numbered
tail feathers; rectrices) feathers of a hummingbird in dorsal recumbency.
readily visible. Animal detection software, such as
Motion Meerkat (Weinstein 2015), is available for
scanning video footage for visitations and movement
patterns of individual hummingbirds. Motion Meerkat
also allows investigators to easily delete video footage
that does not contain hummingbirds.
Published paint marking studies have reported
the use of several body regions, including the crown
(Temeles et al. 2006), the neck or chest (Hilton, Jr. and
Miller 2003; Hurly et al. 2010), and the back (Stiles
1992). The North American Banding Council recom-
mends using the crown for marking birds and this site is
considered the best practice for hummingbirds (Russell
and Russell 2019).
Although a range of pigments have been used
and reported in the literature, non-toxic, water-based,
fabric paint is generally accepted as best practice and
can remain on a bird for weeks to several months
(Russell and Russell 2019). The potential impacts of
using dyes, pigments, or paint to mark a bird always
should be considered prior to application. It is crucial
to minimize the amount of material that is applied.
Primary harmful eects include ingestion, toxicity,
and increased visibility that can result in aggression or
predation. Secondary eects include reduced reproduc-
tive success, reduction in feather waterproong, and
inability to erect feathers completely when a bird is
trying to maximize insulation.
Banding remains the primary means for track-
ing hummingbird movements and longevity as well
as for gathering other important demographic data.
The most common banding method utilizes aluminum
bands. However, one study (Temeles et al. 2013) re-
ported identication of individual hummingbirds at a
distance using specialized colored Darvic tarsal bands
(Avinet Research supplies®, Portland, Maine). A major
constraint when using color bands for marking hum-
mingbirds is the limited visibility of bands aorded by
the extremely small hummingbird tarsi that are covered
with feathers. Because of a hummingbird’s small size,
color banding is typically limited to a single colored
metal band. For a detailed guide on how to band hum-
mingbirds, see the North American Banders’ Manual
for Banding Hummingbirds (Russell and Russell 2019).
For any banding event, particularly if recapture
is not highly likely, the investigator might consider
whether banding is necessary. Anecdotal experiences
suggest regional and species-specic leg swelling pat-
terns in breeding female hummingbirds. Suspected
band-associated leg injuries were reported in recap-
tured female Anna’s Hummingbirds (Calypte anna) in
California during the breeding season (Colwell 2011).
Until further studies are performed, researchers should
consider possible seasonal changes and the importance
of determining proper band sizes for breeding females.
Another consideration when banding hummingbirds
is the relatively high prevalence of pox lesions in the
metatarsal region. Banding a hummingbird with pox
lesions on any area of the body is not advised as the
lesions might progress to the metatarsal region, thus
eectively tightening the band and restricting blood
ow to the distal limb.
A few hummingbird studies have used tags
made from acetate plastic colored sheets (Stiles and
Wolf 1973; Waser and Calder 1975) or ethylene vinyl
acetate (EVA) plastic colored beads (PerlerTM beads,
Igdesigngroup, Atlanta, GA) (Kapoor 2012). To mark
individual hummingbirds, tags were attached to the
back, leg, and/or leg band (Stiles and Wolf 1973; Waser
and Calder 1975; Kapoor 2012). Acetate tags have
been reported to be eective for marking hummingbirds
(Stiles and Wolf 1973; Kapoor 2012); however, there
are concerns regarding their safety. One investiga-
tor used back tags constructed of fused color plastic
beads for identifying birds as an alternative to colored
plastic leg bands (Kapoor 2012). While this method
oers the advantage of short-term visual detection of
marked birds, it may have limitations, especially in the
case of breeding females who might draw predators
to nests. One report (Waser and Calder 1975) raised
concerns that acetate tags may impair hummingbird
nest construction, which could reduce breeding success.
Anecdotal experiences also suggest hummingbird leg-
related injuries associated with the use of acetate tags
attached to either the leg or leg band. Stiles and Wolf
(1973) warned that an acetate tag too tight around a
hummingbird’s leg will restrict circulation with even-
tual loss of the foot. Further studies are needed to
evaluate the safety of acetate tags before they are used
to mark hummingbirds.
24
Integrated Transponder Tags
Radio-frequency identication (RFID) technol-
ogy is a method used to track hummingbird movement
and presence (Brewer et al. 2011; Zenzal et al. 2014;
Hou et al. 2015; Ibarra et al. 2015; Zenzal and Moore
2016, 2019; Bandivadekar et al. 2018). RFID tags
have been miniaturized and do not require batteries;
instead, they are powered by electromagnetic induction.
In addition, RFID tracking antennas may be placed
at hummingbird feeders or around owers. When a
tagged bird passes through or near the antenna, the
unique identifying code is recorded along with the date
and time, thus providing both temporal and geographi-
cal documentation. Tag events are limited to locations
with antennas, and the maximum distance for a tag
read depends on the type of hardware (i.e., tags and
antennas) employed. There is an inverse relationship
between tag detection sensitivity and reading distance;
one study reported a 25.4–30.5 cm maximum tag
reading distance from the center of a circular antenna
(Bandivadekar et al. 2018).
A type of RFID tag called a passive integrated
transponder is most commonly used for hummingbirds
due to their small size and mass. Tags can either be
glued to the feathers (with eyelash or surgical glue) on
the back of a bird or injected under the skin (subcuta-
neously) (Brewer et al. 2011; Hou et al. 2015; Ibarra
et al. 2015; Bandivadekar et al. 2018). If the tag is
glued, extreme care must be taken to avoid inadvertent
gluing of the primary feathers, thereby rendering the
bird ightless. Using a viscous glue in small amounts
helps control glue application and minimizes feather
contamination. Most published studies report gluing
or injecting RFID tags in the intrascapular region (i.e.,
between the shoulder blades) of hummingbirds (Ibarra
et al. 2015; Zenzal and Moore 2016; Bandivadekar et
al. 2018). However, when using subcutaneous applica-
tion, the tag must be placed in the mid lumbar region to
avoid rupturing the cervicocephalic air sacs and caus-
ing generalized subcutaneous emphysema (collection
of air). If the tag is placed too far cranially, chances
of cervicocephalic air sac invasion are substantially
increased. After placing a tag subcutaneously, the en-
trance site can be closed with surgical glue or suture.
In the authors’ experience, suture used in a simple inter-
rupted pattern achieves the best closure and avoids glue
contamination of wing feathers. RFID tag placement on
hummingbirds appears to be successful; however, there
are only a few published studies reporting use of this
technology (Brewer et al. 2011; Hou et al. 2015; Ibarra
et al. 2015; Zenzal and Moore 2016; Bandivadekar et
al. 2018) and more work is needed to determine best
practices and long-term impacts.
Tagging
In recent years, very high frequency (VHF)
tags have been suciently miniaturized for use in
some larger hummingbird species. Whereas RFID
tags record information only at sensor locations, VHF
tags allow for detailed study of individual movement.
However, VHF tags require batteries and are therefore
signicantly larger and heavier than RFID tags. To
meet the standards of the BBL guidelines on auxiliary
marking, the total weight of any tag or any glued or
“backpack” style attachment device must be less than
3% of the bird’s body weight. From a welfare perspec-
tive, the lean (non-migratory) body weight should be
used for calculations. Additionally, given their small
size, the battery life of VHF tags generally is limited
to a few days to two weeks.
Two published studies reported using eyelash
glue and/or surgical glue to attach VHF tags to the in-
trascapular region of hummingbirds (Hadley and Betts
2009; Zenzal et al. 2018). As described with application
of passive integrated transponders, extreme caution is
necessary when using glue near wing feathers. It is also
helpful to trim the tag antenna length shorter than the
bird’s wing length to preclude interference with ight
and to decrease the risk of the bird removing the tag.
4.1. Trends and Applications
A literature review identified 28 published
manuscripts that specied blood collection from hum-
mingbirds, including brachial, ulnar, jugular, or tarsal
venipuncture (i.e., blood collection from a vein) and
toenail clipping (Table 4). Toenail clipping was the
most frequently reported blood sampling method and
the one that the authors consider safest. Eight studies
did not report the blood sampling method. One study
(Tiebout III 1992) disclosed bird mortality. In spite
of the many techniques reported for obtaining blood
samples from hummingbirds (Table 4), a comprehen-
sive research investigation studying the eects (e.g.,
on reproductive success, survivability, and migration)
of obtaining blood samples from hummingbirds still
needs to be performed.
Individuals planning to obtain blood samples
and mark hummingbirds should apply for authoriza-
tion on their BBL permit. Blood sampling should be
performed only by individuals who have expertise in
handling and working with hummingbirds, because the
proper handling and monitoring of birds during these
procedures are just as critical as the blood collection
itself. In addition, a hummingbird’s relatively small
total blood volume means there is little margin for er-
ror in sampling.
Although blood sampling by venipuncture has
been described in hummingbirds (e.g., by puncturing
or sampling from the brachial/ulnar, tarsal, or femoral
veins), the authors do not endorse this method due to
the risk of fatal hemorrhage or secondary injury (e.g.,
broken wing). In addition, the supercial location of the
jugular vein provides limited soft tissue support during
a hemorrhagic event, which can result in fatal blood
loss. Hemorrhage post-sampling is a form of blood
loss and should be avoided at all costs given humming-
birds’ relatively small blood volume. At the time this
manuscript was written, the BBL would authorize only
toenail clipping as a means of obtaining blood samples
from hummingbirds and would authorize blood collec-
tion only by individuals who had received hummingbird
blood sampling training from an approved trainer (B.
Peterjohn, pers. comm., 28 June 2020). Blood sampling
experience with other avian species is not considered
sucient to receive BBL authorization for taking blood
samples from hummingbirds. Because the number of
approved trainers and their time availability are limited,
advance planning is imperative to receive this training.
Wearing nitrile gloves while blood sampling
protects the user from hemostatic agents used to stop
post-sampling hemorrhage, and gloves can be changed
easily between sampling events to minimize spread of
infectious diseases. A standardized guideline used by
avian veterinarians and ornithologists for the amount of
blood that can be sampled safely from live healthy birds
is 1% of the bird’s total body weight in volume during
a single phlebotomy event (Lumeij 1987). However,
because hummingbirds have a very small margin of
safety for blood loss, this general limit does not allow
for an acceptable safety factor. Considerations must
include secondary post-sampling hemorrhage due
to the sampling method, limited investigator experi-
ence, and the health and migratory status of the bird.
Therefore, the authors recommend a more conservative
sampling limit of 0.5% to 1% total lean body weight
(in volume) for the individual hummingbird species.
The precise amount depends on the skill level of the
individual performing the sampling, the blood sampling
technique, the bird’s condition, and the environmental
conditions that might impact the bird’s physiologic and
immunologic status. The lean body weight used for
calculations should be reference values for the species,
because some hummingbird species accumulate consid-
erable adipose stores during migration. For example,
the acceptable range in blood sample volume for a 3.5
g hummingbird is 17.5 to 35 µL. The 35 µL volume
should be used only for bleeding methods after which
hemorrhage is expected to be minimal (i.e., a toenail
clip), if the bleeder has extensive experience, and if the
bird is in very good body condition. Note that hum-
mingbirds living at high altitudes may require special
consideration. Even though it has been shown that
hemoglobin has increased anity for oxygen in hum-
mingbirds inhabiting high altitudes (Projecto-Garcia et
al. 2013), it is unknown how blood sampling impacts
a live bird that is released after sampling. Therefore,
a more conservative sample volume limit is advised
and should be tested incrementally (e.g., starting with
26
0.25% of body weight followed by post-sampling
monitoring) before sampling numerous birds.
In addition to the amount of blood sampled, the
authors recommend considering the timing of blood
sampling. Researchers should avoid times when birds
are at increased risk of hypothermia or hypoglycemia
(early morning, evening, cold temperatures) or when
ambient temperatures limit blood distribution to the
extremities. The range of temperatures in which blood
sampling should be avoided also depends on other
variables, such as humidity or wind. In the author’s
experience (L. Tell), ambient temperatures of 50° F
(10° C) or lower, with no other variables, indicate that
blood sampling should either be postponed, performed
in a warmer environment, and/or the bird should rst
be slowly warmed with supplemental heat before
sampling. In cool ambient temperatures, peripheral
vasoconstriction may necessitate clipping a large dis-
tal portion of the toenail, thus risking amputation of
the phalangeal bone. Best practice suggests forgoing
sampling if blood does not ow from a conservative
toenail clip. Alternatively, a supplemental heat source
can be used to facilitate vasodilation in the extremi-
ties. However, the heat source must be at a relatively
low setting to avoid overheating and to prevent a large
temperature dierence between sampling and release
to the wild. Heat sources intended for young animals
(e.g., puppies or kittens) or those that can be placed
in an animal crate typically have low settings. If the
ambient temperature is cool in the early morning but
rises later in the day, it is very important discontinue
use of the external heat source. Birds should never be
left unattended while lying on a heat source. Birds that
are overheating will open-beak breathe (pant) or will
move around more than usual; however, birds showing
none of these signs may still be overheating. Therefore,
the bird, the ambient temperature, and the heat source
temperature must be monitored continuously.
Table 4. Summary of methods used to obtain hummingbird blood samples as reported in published studies.a
Bleeding method Count Volume per
bird Applications References
Brachial
venipuncture
6 1–200 µL Endocrinology, genetics,
pathology
Williams 1978; Hiebert et al. 2000b;
Gregory et al. 2009; Projecto-Garcia et al.
2013; Matta et al. 2014; González-Gómez et
al. 2015
Ulnar venipuncture 1 30-200 µL Genetics Projecto-Garcia et al. 2013
Femoral
venipuncture
1 15 µL Genetics Roy et al. 1998†
Jugular venipuncture 1 70-100 µL Energetics Weathers and Stiles 1989
Tarsal venipuncture 2 30 µL Endocrinology González-Gómez et al. 2014a, b
Toenail clip 12 10-50 µL Energetics, endocrinology,
isotope analysis, clinical
pathology, parasitology,
physiology
Bakken et al. 2004; Bakken and Sabat 2006;
Hardesty and Fraser 2010; Fernández et
al. 2011a, b; Matta et al. 2014; González-
Gómez et al. 2015; Hagadorn et al. 2016;
Bradshaw et al. 2017; Safra et al. 2018;
DeRogatis et al. 2020; Godwin et al. 2020
Unreported method 8 20-40 µL Energetics, genetics,
physiology
Tiebout and Nagy 1991; Tiebout III 1992‡;
Chaves et al. 2007, 2011; Parra et al. 2009;
Chaves and Smith 2011; Harrigan et al.
2014; Wright et al. 2014
a Literature searched until September 2020.
†Specically reported that no birds died during bleeding.
‡Reported bird deaths associated with blood sampling.
The small body size and limited total blood
volume of some hummingbird species also means
there are limits on repeated blood sampling, especially
within a short time period. A minimum of 7–10 d from
a previous bleeding event should pass before a bird is
resampled. This will give the hummingbird a chance
to recover and compensate for blood loss. Even though
permitting agencies do not require marking a bird that
has been blood sampled, it is recommended (L. Tell)
that any un-banded hummingbird being blood sampled
be marked in a subtle way, such as minimal clipping
of a feather. The marking method does not have to be
permanent (such as leg banding) for purposes of short-
term identication, especially if birds in the area are
not monitored on an ongoing basis.
Blood Samples from Hummingbirds
Toenail clipping is the predominant and safest
method for obtaining blood samples from live hum-
mingbirds, especially for researchers who are not
experienced with blood sampling. To obtain a blood
sample from the toenail, the authors recommend using
a mesh bag made of seine material (see section 2.3) to
help restrain the bird and allow for easy access to the
toenails. The bird is restrained within the mesh bag in
a supine position with both wings positioned against its
body. The researcher then folds the long sides of the
mesh bag underneath the bird so that the bird is conned
to a limited space (Fig. 6). The bird is then placed on
Figure 6. Illustration of the toenail clip blood-sampling method for obtaining a blood sample from a hummingbird
that is restrained in a seine mesh bag. An oblong “bean bag” is used to rest and steady the researcher’s left hand
and also minimize bird movement within the holding bag. Curved cuticle scissors (with scissors positioned
in a concave “up” position) allows clear view of the toenail to be clipped. Because the researcher’s left hand
is resting on the “bean bag”, the thumb and second ngers that are restraining the toenail are not placing any
pressure on the bird’s chest.
28
a at surface, and a weighted object is placed next to
the bird to constrict bird movement in the bag. So as
not to impair respiration, the weighted object should
not be placed on top of the bird or too tightly against
the bird’s body. In Figure 6, the weighted object is an
oblong beanbag constructed of a four-way stretchable
polyester fabric stued with small polystyrene beads.
The small beads are lightweight and conform easily
to the bird’s side. The weighted bag also provides a
structure on which to rest a hand while obtaining the
blood sample so that the hand does not lean on the
bird’s body.
Commercial stainless steel, curved-blade scissors
(i.e., Revlon® curved-blade cuticle scissors, Revlon,
Inc., New York, NY) are ideal for cutting hummingbird
toenails for blood sampling because the curved blade
facilitates visualization of the toenail tip. The stain-
less steel ensures that scissors are sharp, even after
continuous use, and helps achieve a clean cut. Before
sampling, scissors should be cleaned with a sanitizer
(i.e., Super Sani-Cloth® germicidal disposable wipe,
PDI, Woodcli Lake, NJ), rinsed with water, and dried.
Researchers can use their ngernails to gently grasp a
toenail and pull the foot out through an opening in the
bird bag. Caution should be used when grasping the
toenail, because excessive manipulation can avulse the
keratin sheath.
Once the foot is accessible, it is held at the tarsal
joint. The second, third, and fourth digits can be then
be spread out and controlled using the researcher’s
fingers—thus keeping them protected—while the
toenail of the third digit is being cut. The researcher
should be aware of hand placement so as not to put
pressure on the bird’s body and restrict respiration.
Based on experience (L. Tell), the toenail of the third
digit is recommended for blood sampling because it is
the longest nail. The distal aspect of the toenail is cut
perpendicular to the digit, removing a maximum of 5%
to 10 % of the total nail length. Although it is unknown
if the phalangeal bone is cut during this process, the
risk of amputating the phalangeal bone increases as
more toenail is cut. Amputation of the phalangeal bone
is painful for the hummingbird. Using magnication
during this procedure helps the researcher visualize
toenails of digits two and four while the toenail of
digit three is cut.
If blood does not ow after the toenail tip is cut,
one should ensure that blood ow is not being impeded
by nger restraint of the toe before cutting more of the
toenail. Even a slight amount of nger pressure when
holding a hummingbird’s foot or toe can impede blood
ow. With the ngers still in place around the toes,
very gently release and re-apply pressure to the toes
to facilitate blood ow.
As discussed above, cool ambient temperatures
also can impact blood ow from a cut toenail. Cold am-
bient temperatures result in peripheral vasoconstriction
of the extremities; therefore, attempting blood sample
collection via toenail clipping at temperatures below
10° C (50° F) is not advised. At low temperatures, more
of the toenail will need to be cut for blood sampling,
which increases the risk of phalangeal bone amputa-
tion. In the presence of cooler ambient temperatures,
a supplemental heat source can be helpful (see section
4.1). In contrast, warm ambient temperatures result in
vasodilation and blood will ow freely from the clipped
toenail. Therefore, it is imperative to have hemostatic
materials readily available after the toenail is clipped.
Until hemostasis is achieved, rm digital pressure on
the foot or toes also can help restrict blood ow.
Absorbent materials used for blood storage are
ideal for sample collection because they can be blotted
against the cut toenail tip to facilitate blood collec-
tion; examples include Whatman® Flinders Technol-
ogy Associates (FTA) sample cards (GE Healthcare,
Wauwatosa, WI) or lter paper strips (i.e., AdvantecTM
Nobuto blood lter strips, Cole-Parmer, Vernon Hills,
IL). If lysis buer is used, it is easiest to collect the
blood sample using a laboratory grade pipette (volume
capacity 0–20 µl) and ejecting the blood directly into
the center of the tube containing the buer. Another op-
tion for obtaining a blood sample is to place the lumen
of a capillary tube at the site of the cut toenail. This
works best with warm ambient temperatures because
blood ow is typically consistent. Blood smears can
be made by blotting a glass slide against the site of the
cut toenail or by letting a drop of blood fall onto the
coverslip.
Once the sample is collected, a hemostatic agent
such as silver nitrate or styptic powder is applied to
the cut toenail. Even if blood did not ow from the
cut toenail or if bleeding appears to have stopped, a
hemostatic agent always should be applied to prevent
unexpected bleeding post-release. Styptic powder does
not tend to adhere to the tip of a hummingbird’s cut
toenail (L. Tell). Therefore, the authors recommend
direct application of silver nitrate using a cautery stick.
A prescription from a veterinarian is needed for order-
ing silver nitrate cautery sticks. When using silver
nitrate sticks, the silver nitrate end needs to be held
directly and rmly against the toenail tip while the stick
is rotated. The researcher’s skin is protected by using
a nitrile-gloved hand to hold the toenail.
To assess whether the toenail can be cut without
amputating a portion of the phalangeal bone, the authors
used ve hummingbirds that were being euthanized
for other reasons. The tip of the third toenail was cut,
simulating toenail blood sampling in the eld. After
euthanasia, the foot was immediately xed in 10%
neutral formalin. The toes were processed through
alcohol and xylene solutions and embedded in paran,
ensuring that the digits were placed on their sides and
tamped down for straightening as the paran cooled.
Toes were cut in 4-micron serial sections for complete
microscopic examination of the entire distal phalanx.
Sections were stained with hematoxylin and eosin and
examined. Four of ve cut toenails did not have pha-
langeal bone imposition (Fig. 7A), whereas one toenail
had evidence of bone amputation (Fig. 7B). All capture,
handling, sampling, and euthanasia procedures for this
work was approved by the University of California-
Davis IACUC and were conducted under authorization
of federal and state scientic permits (LAT; USGS BBL
permit #23947, US Fish and Wildlife Scientic Collec-
tion permit MB55944B-2, and California Department
of Fish and Wildlife SC-013066).
Figure 7. A) Hummingbird’s (subject #1) distal phalanx of third toe after the keratin sheath of
the toenail was cut. Hematoxylin and eosin stain. B) Hummingbird’s (subject #2) distal phalanx
of third toe demonstrating the bone of the distal phalanx was cut. Hematoxylin and eosin stain.
A B
Advantages to the toenail clip method for obtain-
ing blood samples are that there is a low probability of
hematoma formation (swelling of blood within tissues
or under the skin) and a high probability of hemostasis
at the venipuncture site following cauterization. The
method is relatively safe compared to blood sampling
from soft tissue sites. In addition, if the bird is sampled
when the ambient temperature is at least 10° C (50°
F), the maximum blood volume allowed for a single
bleeding event is achievable. There are three disad-
vantages to the method: 1) with excessive and repeated
manipulation of the toenails, the keratin sheath of the
toenail might be mistakenly avulsed while trying to
extract the toenail from an opening in the mesh bag;
2) if the bird is cold, the blood ow rate might be less
than with soft tissue venipuncture, and excessive cut-
ting of the toenail might result in painful amputation
of the phalangeal bone; and 3) there is risk of human
skin exposure to silver nitrate if cautery sticks are being
used without a gloved hand.
A literature review revealed 18 manuscripts that
described seven methods for storing hummingbird
blood for later use (Table 5). The sample storage
method is highly dependent upon the research applica-
tion. For genetic analysis, some investigators prefer
using Whatman® FTA sample cards (GE Healthcare)
due to the ease collecting blood onto a card when using
a toenail clip bleeding method and the stability of the
sample at room temperature. For general blood storage
or testing for infectious diseases, blood lter strips work
well due to ease of blotting blood onto the strip when
using the toenail clipping blood collection technique
and the ability to distance blood spots along the strips.
For analysis of blood smears for estimated white
blood cell counts and the presence of hemoparasites,
the cover slip method for making blood smears is
useful when only a small volume of blood has been
obtained during a toenail clip. This technique puts
minimal traumatic force on the cells if the individual
making the slide has limited experience. The authors
found that using two medium-sized (24 mm x 50
mm) versus small-sized (20 mm x 20mm) cover slips
facilitates making a feathered monolayer blood smear
because there is additional space for holding the cover
slips as they are separated. Helpful eld guidelines
for making and staining blood smears for species with
nucleated red blood cells can be found at https://www.
uvm.edu/~jschall/techniques.html. A modied Wright's
stain (Camco Quik Stain®, Cambridge Diagnostic
Products, Inc., Fort Lauderdale, FL) is a commercially
available rapid stain that works well with hummingbird
blood smears (Safra et al. 2018). Figure 8 details a
protocol for staining hummingbird blood slides that is
modied from the manufacturer’s recommendations.
A review of the literature regarding feather
sampling from hummingbirds revealed 33 results and
revealed trends in feather type, quantity, and application
used (Table 6). More research is needed to compile
best practices for feather sampling from hummingbirds.
As with all avian species, hummingbird feathers have
critical behavioral, thermoregulatory, and physiologic
functions (McDonald and Grith 2011). Thermoregu-
lation especially is important for hummingbird survival
and should not be compromised. Studies evaluating the
impacts of molt on bird health and survival in other bird
species have shown signicant costs in ight maneuver-
ability after feather loss and energy expenditure during
feather regrowth (Swaddle and Witter 1997; Bridge
2004). One investigator documented that removing
all of a hummingbird’s tail feathers slightly reduced
its ight maneuverability (Clark 2011a). Feathers also
serve important communication functions; for example,
males of many hummingbird species can produce court-
ship sounds through aeroelastic tail utter during the
breeding season (Clark et al. 2018a; Clark and Mistick
2018b). Therefore, it is very important to be thorough
in evaluating which hummingbird feathers and how
many can be removed for sampling.
Table 5. Summary of methods used to store hummingbird blood or urine samples as reported in published studies.a
Sample
type Count Storage method Temperature Applications References
Blood 1 Nobuto blood strips Not reported Genetics Hagadorn et al. 2016
Blood 3 Whatman FTA Elute cards Not reported Phenology Correa-Lima et al. 2019; Baek et al. 2020; Godwin et al.
2020
Blood 5 Sealed capillary or
microhematocrit tubes
-80° C to 5° C Endocrinology,
energetics
Weathers and Stiles 1989; Tiebout III 1992; Fernández et al.
2011a, b; González-Gómez et al. 2014a
Blood 3Centrifuged immediately and
plasma extracted then frozen
-80° C Energetics,
physiology, genetics
Hiebert et al. 2000c; Bakken and Sabat 2006; Projecto-Garcia
et al. 2013
11.5-mL tube in 99% ethanol Not reported Isotope analysis Hardesty and Fraser 2010
Blood 1 SET buer -20° C to ambient
room temperature
Pathology Matta et al. 2014
Blood 5 Blood smear -20° C to ambient
room temperature
Genetics, pathology Williams 1978; Gregory et al. 2009; Matta et al. 2014; Wright
et al. 2014; González-Gómez et al. 2015
Urine 1 Frozen in 1.8-mL cryotube with
tris borate EDTA buer
20° C Pathology Henning et al. 2015
Urine 1 Cryotube frozen in liquid
nitrogen
-80° C Pathology Williams et al. 2012
Urine 2 Microcentrifuge tube,
centrifuged, then frozen
-20° C Endocrinology Hiebert et al. 2000a, b
Urine 2 Nalgene or other tube frozen -20° C Physiology López-Calleja and Bozinovic 2003; McWhorter et al. 2003
Urine 1 Dried for weight 60° C for 6 days Energetics López-Calleja and Bozinovic 2003
Urine 1 Frozen in 1-mL vials -5° C Toxicology Bishop et al. 2018
Urine 2 Flame-sealed capillary tube Not reported Energetics Barbachano-Guerrero et al. 2019; Shankar et al. 2019
a Literature searched until September 2020.
Figure 8. Suggested protocol for using a commercially available
modified Wright's stain to stain cells on blood smears from
hummingbirds.
Table 6. Summary of methods used to collect hummingbird feather samples as reported in published studies.a
Feather source Count Quantity Applications References
Tail
R4 2 1 feather Isotope analysis Brown et al. 2012; Moran et al. 2013
R5 32 feathers Genetics González and Francisco Ornelas 2014; Licona-Vera and Francisco
Ornelas 2014; Rodriguez-Gomez and Francisco Ornelas 2014
R4 and R5 4 2 feathers Physiology Clark 2011b; Clark et al. 2011a, b, 2012
R3 and R5 1 1 feather Genetics Malpica and Ornelas 2014
R1 2 1 feather Color spectrometry, viral DNA
detection
Meadows et al. 2012; Baek et al. 2020
Varied 2 1–2 feathers Trace element analysis, Tabletop
scanning electron microscopy
Mikoni et al. 2017; Yamasaki et al. 2018
Unspecied 6 2 feathers Genetics, isotope analysis Hardesty and Fraser 2010; Gonzalez et al. 2011; Rodríguez-Gómez et
al. 2013; Ornelas et al. 2016; Hernández-Soto et al. 2018; Rodríguez‐
Gómez and Ornelas 2018
Head/crown 5 5 feathers Color spectrometry Parra et al. 2010; Meadows et al. 2011, 2012; Dongen et al. 2013;
Eliason et al. 2020
Back 3 5 feathers Color spectrometry, genetics Parra et al. 2010; Dongen et al. 2013; Eliason et al. 2020
Chest 4 5 feathers Isotope analysis, genetics,
toxicology
Brown et al. 2012; Dongen et al. 2013; Hagadorn et al. 2016; Sierra-
Marquez et al. 2018
Chest 2 Multiple
feathers
Trace element analysis, viral DNA
detection
Mikoni et al. 2017; Baek et al. 2020
Gorget 4 5 feathers Color spectrometry Parra 2010; Meadows et al. 2011, 2012; Eliason et al. 2020
Back 2 5 feathers Color spectrometry, genetics Parra 2010; Dongen et al. 2013
Primaries and secondaries 1 2 feathers Mite study 96% ethanol Mironov et al. 2019
Unspecied feathers 7 2 feathers Genetics, irredescence Roy et al. 1998; Hardesty and Fraser 2010; González-Gómez et al.
2011, 2013; Gonzalez et al. 2011b; Ornelas et al. 2016; Sosa et al.
2020
a Literature search until September 2020.
Hummingbirds
5.2.1. Appropriate timing for feather sampling:
Migratory North American
hummingbirds exhibit a molting strategy that is simi-
lar across species. Most of these species undergo a
pre-basic molt on their wintering grounds beginning
in August to October and ending by January to March
(Pyle 1997). Within these dates, there is some varia-
tion among species. For example, some southwestern
species (e.g., , and
Lampornis) can have protracted molts that extend into
April or May. Exceptions to this molting strategy are
the Calypte and Amazilla species in which pre-basic
molt is primarily from June to September (Pyle 1997;
Pyle et al. 1997). Anna’s Hummingbirds (Calypte
anna) can have a protracted molt lasting from May
through January (Williamson 1956). Flight feather
replacement is sequential in all species, and sequen-
tial replacement of tail feathers begins sometime after
primaries are replaced. A recently discovered hatch
year, second pre-basic molt in winter has been found
in Ruby-throated (Archilochus colubris) and Rufous
Hummingbirds (Selasphorus rufus), but this should
not alter the timing of feather sampling (Dittmann and
Cardi 2009; Sieburth and Pyle 2018).
5.2.2. Appropriate timing for feather sampling:
Considerations regarding the
timing of feather sampling should minimize impacts
on the reproductive activities (e.g., courtship) and
thermoregulation of hummingbirds. Thus, sampling
should be synchronized to the expected breeding and
molting periods of their life cycle. For adult male and
female North American hummingbird species, feather
sampling is best limited to the period between the end of
nesting and the start of molting, which is between July
and September for most species. For example, removal
of an adult male’s iridescent head or gorget feathers
should be avoided during the breeding season. In ad-
dition, tail feathers and ight feathers from adult males
can be sampled after the breeding season and before
the winter molt without aecting courtship behavior.
Species-specic nesting periods can be ascertained by
using breeding season data in the
series (Cornell Lab of Ornithology) and molting dates
can be obtained (Pyle 1997; Pyle et al. 1997). Flight
feathers should not be sampled during migration, but
sampling can be considered during winter when birds
are molting. Sampling of primary feathers should be
limited to P1 and P2 (Fig. 5) unless birds are in primary
molt when older outer primaries can be sampled. When
considering sampling tail feathers, R2 and R3 (Fig. 5)
are recommended at all times.
Special consideration should be given to sampling
hatch year (HY) birds. Because these birds require
intact ight feathers prior to migration, they have
especially critical energy demands. In some species,
HY birds that arrive in wintering grounds undergo a
second pre-basic molt and replace all of their ight
feathers (Sieburth and Pyle 2018), thus following the
same molting schedule as adult birds. Sampling old
feathers at this time is acceptable. After molting in
the wintering grounds, HY birds attain adult plumage
and should not be sampled until after the breeding
season. The researcher should be familiar with the
dates of migration, courtship, and nesting of the hum-
mingbirds involved in their study in order to select the
least invasive time for feather sampling. In addition to
assessing the risks associated with removing a certain
type of feather from a given species, knowledge of the
proper technique for feather sampling is paramount.
Hummingbird skin is delicate. If feathers are pulled
too aggressively or in the wrong direction, the skin can
tear, necessitating closure with suture.
5.2.3. Methods for sampling contour feathers.—
When sampling contour feathers, the recommendation
is to collect a total of 20 to 30 feathers from four dif-
ferent quadrants and feather tracts in the chest region.
Feathers should be removed without creating patches
of exposed skin that could compromise thermoregula-
tion. This is especially important during winter months
or at high elevations when/where birds encounter low
temperatures. Ideally, feathers are removed at scattered
locations over the bird’s body so that normal preening
can cover the sampled areas. However, research is still
needed to evaluate the impacts of feather sampling on
hummingbirds. A hemostat or blunt-tipped forceps is
recommended for obtaining feathers. Both are ecient
for sampling contour feathers and minimize the chance
of inadvertently grasping skin during the sampling
process. During feather sampling, the bird is held in a
supine position in a cupped hand (Fig. 9). The bird’s
feet are secured so that they are not inadvertently
harmed during the sampling process. The instrument
used to pull feathers (e.g., forceps or a hemostat) is held
at the proximal end of the feather shaft (i.e., close to
where the feather attaches to the skin) with the instru-
ment tip positioned perpendicular to the long axis of
the bird (Fig. 9). Pulling feathers down and away from
feather follicles minimizes the risk of inadvertently
lacerating the skin. Before sampling, it is important
to visually conrm that no skin is contained within the
forceps or hemostat.
5.2.4. Methods for sampling wing and tail feath-
ers.—In general, protocols for sampling hummingbird
wing and tail feathers should be similar to guidelines
provided for sampling these feathers in passerines (Fair
and Jones 2010). An important exception involves the
possible sampling of tail feathers that are important to
hummingbird males during the breeding season. Re-
searchers also must consider the metabolic expense of
feather replacement for hummingbirds, a species that
already has signicant energy demands.
Figure 9. Illustration of sampling covert feathers from the pectoral region of a hummingbird. Note that the
bird is being restrained using the hummingbird bander’s hold with the fourth and fth ngers restraining the
feet. The forceps are held in a horizontal position relative to the long axis of the bird’s body, which helps
minimize the chances for inadvertently pinching the skin. When sampling, the feathers are grasped within
the opening of the forceps. The forceps are moved in a downward direction (in the direction of the arrow) to
minimize the chance of inadvertently tearing the skin when feather sampling. The feathers should be placed
in an acid free envelope to maximize the duration of sample usefulness for scientic studies.
Removal of a wing or tail feather can be accom-
plished without using an instrument. Secure the feather
base between the thumb and forenger and pull gently
but rmly while using the ngers from another hand
to stabilize the anatomic region where the feather is
attached to the body. For the safety of the bird and to
ensure proper removal, grasp the feather at the proximal
end (base) of the feather shaft during removal. If a
wing feather is sampled, the wing is stabilized with the
ngers of the opposite hand so that the bones are not ac-
cidentally fractured during the feather removal process.
As mentioned previously, knowledge of the acoustic
function that individual feathers play in courtship and
behavior is important in determining which feathers
to sample (Clark 2011b; Clark et al. 2018a; Clark and
Mistick 2018b). A best practice is to avoid sampling
wing feathers if an alternative feather type (such as a
tail feather) would suce; that is because wing feathers
are very important for ight and sometimes produce
sound (e.g., in Archilochus spp.). Of the ten primary
feathers, P10 (Fig. 5) contributes the most to production
of aerodynamic force. No research studies have been
conducted to determine if tail feather sampling aects
the overall reproductive tness of a hummingbird
population; however, sampling tail feathers other than
the acoustic tail feathers is best practice. If females
preferentially select a male based on the sounds made
by tail feathers, the change in tness for one male will
most likely result in a tness increase for another male.
Therefore, while there might not be a net change in
tness for the population as a result of sampling tail
feathers, tness for the individual sampled bird might
be altered due to human intervention.
5.2.5. Methods for sampling blood feathers.—
Compared to mature feathers, blood feathers—which
contain pulp tissue and blood—oer a more reliable
source of deoxyribonucleic acid (DNA). Sampling
blood feathers requires blood sample authorization
either on BBL or USFWS Migratory Bird Scientic
Collecting permits, depending on whether the bird is to
be banded or not. Because blood feathers have an active
blood supply, they must be removed in their entirety.
Removal of a blood feather should be performed only
by an individual with experience in removing blood
feathers. When a blood feather is removed, the feather
should be examined carefully to ensure that the entire
base of the feather shaft was removed; this will ensure
involution of the skin follicle’s vascular supply. If the
entire shaft of the blood feather is not removed, fatal
hemorrhage can occur. If any part of the shaft remains
and has sucient integrity, micro-hemostats can be
used to grasp and remove the remaining tissue. Mag-
nication is helpful during this process. Once the entire
blood feather has been removed, digital pressure can be
applied to the feather follicle. To avoid wing bone frac-
tures, special care must be taken when sampling wing
feathers. Digital pressure should be released if the bird
attempts to ap its wing; pressure can be reapplied once
the bird is calm. Because blood feather removal can
cause temporary discomfort and has inherent risk, the
researcher’s skill level and necessity/value of the blood
feather sample should be considered before sampling.
The total blood volume in a blood feather from
a hummingbird varies depending on the maturity and
size of the feather. Because early-developing wing and
tail blood feathers potentially have the greatest blood
volume, it is advisable for a researcher to sample only
one wing feather or tail feather from a hummingbird
during a single encounter. Further recommendation is
to wait at least 7–10 d before sampling another large
blood feather from that individual.
Analysis
Most studies reviewed did not specify how feath-
er samples were stored; however, those that did used
paper receptacles, such as coin envelopes. Acid-free
envelopes are recommended, and silica gel packs help
minimize moisture. A sample identication number can
be assigned to the envelope, which is cross-referenced
to the sampled bird by using a unique catalog number.
It is best to label envelopes in pencil and then place tape
over the writing to avoid smearing. Alternatively, feath-
ers can be stored taped to 3-in x 5-in archival cardstock,
which is sucient to hold the entire set of tail feathers
from a single hummingbird. The card is then slipped
inside a 3-in x 5-in archival glassine envelope, which
will help minimize specimen damage. These samples
can then be stored inside a 3-in x 5-in card box. Foil
or plastic bags also can be used to store feathers. Se-
lection of the storage system to use will depend on the
intended study (Espino-Espino et al. 2014). Although
it is possible to extract genetic material from feathers
maintained at room temperature, there is anecdotal
evidence (L. Tell) that freezing hummingbird feathers
at -20° C or -80° C may help minimize DNA degrada-
tion over time. If envelopes will be frozen or exposed
to moisture or humidity, the glue on the envelope ap
may not hold. Applying a small piece of adhesive
tape to the envelope seal will help prevent the loss or
mixing of samples.
6.1. Trends and Applications
Review of the literature found two published
studies mentioning oral and cloacal swab sampling
on hummingbirds (Williams et al. 2012; Barbachano-
Guerrero et al. 2019); however, the materials and meth-
ods were not specied. Oral cavity and cloacal swab-
bing are commonly performed in other avian species
to obtain samples for determining normal oral ora or
for pathogen (bacterial, yeast/fungal, parasitic, or viral)
testing. Swabbing the mouth or cloaca is not performed
routinely with hummingbirds due to the fragility of the
beak/tongue and the lack of a small enough swab tip
for insertion into the hummingbird cloaca.
Given the previously mentioned challenges,
methods for obtaining oral or cloacal swabs from
hummingbirds are not well-established. For obtain-
ing samples from the proximal gastrointestinal tract,
palpating the crop to induce uid regurgitation is not
recommended, because the bird could aspirate a small
amount of uid (that is not visually obvious) and the
health ramications would not manifest until 24–48 h
after the bird is released. An alternative to oral cav-
ity swabbing is to have a hummingbird drink from a
small vial lled with sterilized sugar water and then
sample from a small volume of sugar water that was
not ingested (Lee et al. 2019). Depending on the study
goals, this method might provide samples representing
the oral microbiota. An alternative for cloacal swab-
bing might be to use the tip of a cellulose eye spear
(i.e., Weck-Cel®cellulose eye spears, BVI Medical,
Waltham, MA). These sponges are made from highly
absorbent natural cellulose material and are designed
for use in delicate surgical areas.
A literature review yielded 21 results for cloacal
excreta sampling in hummingbirds as well as trends in
sampling techniques and applications (Table 7). Col-
lecting excreta (urine and/or feces passing through or
having exited the cloaca) from a conned but unre-
strained bird is a non-invasive method for urine and/
or fecal sampling. Urine or feces also can be collected
from a restrained bird.
When measuring analytes in urine samples from
birds, there are some fundamental dierences between
urine samples obtained directly from the ureters versus
the cloaca. Solute concentrations in cloacal urine can
dier substantially from ureteral uids released by the
kidney despite birds not having a bladder (that results
in urine pooling) nor substantial urine concentrating ca-
pabilities (Lotz and Martínez Del Rio 2004) compared
to mammals. The reason for the cloacal urine to dier
from the ureteral urine is attributed to dietary water
excreted from the gastrointestinal tract into the cloaca,
because avian gastrointestinal and urinary tracts join at
the cloaca. The uid volume that hummingbirds drink
and excrete through their gastrointestinal tract could
have a substantial impact on cloacal urine solute con-
centrations, as has been shown in sunbirds (McWhorter
et al. 2004). For mammalian urine samples, creatinine
typically is used as an endogenous marker to normalize
analyte concentrations and account for urine dilution or
concentration. However, birds have limited metabolism
of creatine to creatinine (Paton 1910), and therefore
creatinine is not useful for normalizing urine samples.
Creatine has been measured in bird urine samples by
modifying a creatinine analytical method (Wimsatt et
al. 2009); however, little is known about the use of
Table 7. Summary of methods used to collect hummingbird cloacal excreta samples as reported in published studies. a
Method Count Volume per bird Applications References
Uretal urine collected with a
close-ended cannula
2 Not reported Physiology Bakken et al. 2004; Bakken and Sabat 2006
Microcapillary tube held up to
cloaca
230–50 µL Endocrinology González-Gómez et al. 2014a, b
Micropipettes or microcapillary
tubes used to collect from surface
of a cage liner
10 50–1200 µL Energetics,
endocrinology,
physiology
Preest and Beuchat 1997; Hiebert et al. 2000a, c; McWhorter and
Del Rio 2000; López-Calleja and Bozinovic 2003; López-Calleja et
al. 2003; Lotz and Martínez Del Rio 2004; Bakken and Sabat 2006;
Golo and Burch 2012; Chavez-Zichinelli et al. 2014
Cloacal uid collected directly
from the cloacal opening using
100- µL micro-pipette
2 Anna’s Hummingbird
(30–60 µL)
Rufous Hummingbird
(5–30 µL)
Toxicology for
pesticides
Bishop et al. 2018, 2020
Cloacal swab 1 Not reported Pathology Williams et al. 2012
Fecal smears 1 Not reported Pathology Snowden et al. 2001
Oral and cloacal swabs (frozen
in viral transport media
supplemented with antibiotics
(stored at -80° C)
1 Not reported Prevalence of West
Nile virus
Barbachano-Guerrero et al. 2019†
Oral (sugar wáter in vials) 11 ml oered for
birds to drink.
Approximately 500-
900 µL of remaining
sugar water in vial
used for sampling.
Microbiome Lee et al. 2019
Cloacal pellets collected from
area around nest (stored at -20°
C)
1 Not reported Diet analysis Moran et al. 2019
a Literature search up to September 2020.
†Samples were collected from birds’ beaks by allowing birds to drink from 1.5-ml tubes.
creatine to normalize solute concentrations in avian
urine samples. Although specic gravity, as measured
by a handheld refractometer, has been used as a proxy
for normalizing solute concentrations in urine samples
from hummingbirds, George (2001) suggested that this
method should not be broadly applied to urine samples
from all animals. Multiple constituents in urine samples
can falsely elevate or decrease specic gravity values;
therefore, a validation study is necessary before using
specic gravity to normalize solutes in hummingbird
urine samples.
Sampling from Hummingbirds
Using this method, a captured hummingbird is placed
in a small enclosure with a perch, a feeder, and enough
space for the bird to comfortably feed. A soft mesh
buttery cage works well for this purpose. The bot-
tom of the enclosure is lined with a material that will
facilitate urine or fecal collection, such as plastic wrap
or plastic-coated paper. If urine collection is a goal,
micropipettes or microcapillary tubes can be used to lift
excreta samples from the oor liner. Filter paper can be
used to collect feces for DNA sampling, because it can
be easily frozen, cut, and immersed in uids for DNA
extraction. This reduces the need to collect excreta o
the paper. The cage liner method for collecting cloacal
excreta from hummingbirds is easy, does not require
specialized equipment, and involves very little risk to
the bird. However, this method is relatively inecient
because some sample volume may be lost during trans-
fer from paper to pipette or tube.
7.2.2. Manual restraint and capillary tube
The method of
collecting urine samples with capillary tubes, as rst
established in rats (Hayashi and Sakaguchi 1975), has
been applied to hummingbirds (Hiebert et al. 2000a;
González-Gómez et al. 2014a, b) for free-catch collec-
tion of naturally voided urine or feces. Only individuals
with training and experience with handling humming-
birds should use this approach. The hummingbird is
manually restrained and held in an upright position
while the opening of a capillary tube is placed near the
cloacal sphincter so that the voided urine or feces can
be directly sampled as excreta exits the cloaca (Fig. 10).
This handling technique for urine collection works for
smaller-sized hummingbirds (less than 5–6 g). The
hummingbird should be restrained immediately after
capture so it does not void urine/feces before sample
collection can occur. Fecal sample collection may be
more successful later in the day rather than rst thing
in the morning because birds will have had more time
to forage. While birds are being restrained, they should
be monitored continuously and oered sugar water as
needed. If urine/feces are not voided after 10–15 min
of restraint, the bird should be released. Advantages
to this method are that it is non-invasive, it minimizes
the time required for sampling, and it eliminates the
need to keep the bird in an enclosure. It is important to
note, however, that even if fecal material is not present
in the urine sample, the urine collected is not sterile
because it is not sampled directly from the lumen of the
ureters. A relatively small mass of fecal material also
can be collected using this method. When using this
method, the handler must be able to restrain the bird
without impairing its respiration while simultaneously
monitoring the bird’s status and collecting samples.
7.2.3. Dish collection for excreta sampling.—A
simple method for urine/fecal collection is to place
hummingbirds in a mesh bag and lay them in a sitting
position on a plastic, large-mesh grating over a dish.
Typically, hummingbirds will produce an ample urine
sample within 15–20 min, which can be collected in a
capillary tube from the bottom of the dish. Urine col-
lection is most eective when trapping periods occur
in the morning after the intense period of post-roost
feeding when hummingbirds are well hydrated.
Analysis
Most of the reviewed papers suggest storing
cloacal excreta samples in small cryotubes (1.5–1.8
mL) with or without a preservative buer (e.g., 0.9%
saline). This is followed by either urine/fecal sample
separation by centrifuge or storage in a freezer at -20°
C or -80° C (Table 5).
40
8.1. Trends and Applications
A literature search yielded 68 published studies
that used methods described in this section (Table 8).
Measuring metabolic rates in hummingbirds is of broad
interest to the scientic community for several reasons:
1) hummingbirds are among the smallest endotherms;
2) they have among the highest metabolic rates re-
corded in vertebrates (Lasiewski 1963a); 3) they are
capable of using deep torpor at night (Hainsworth and
Wolf 1976; Hiebert 1992; Powers et al. 2003); and 4)
they are the only birds that employ sustained hovering
ight (Warrick et al. 2005). Methods for measuring
metabolic rates in hummingbirds are similar to those
used for other animals. The biggest challenges result
from the birds’ small size and lack of long-term endog-
enous energy stores.
Figure 10. Illustration of urine/fecal collection from a hummingbird using a glass
capillary tube that is held in close proximity to the cloacal opening. The bird is
gently restrained in a cupped hand and the tail is reected cranially and dorsally.
Using this restraint method, the person sampling must constantly monitor for eye
closure and overall bird status since the head is not as easily visualized when the
bird is being restrained in this manner.
Table 8. Summary of methods used to measure hummingbird metabolic/evaporation rates as reported in published studies.a. Note that some studies listed
were conducted in a laboratory setting, but the methods can be applied to the eld.
Measurement Count Protocol Applications References
Basal metabolic rate 12 Open-ow,
positive pressure
Nighttime energy costs
(laboratory)
Lasiewski 1963a, b, c; Weymouth et al. 1964; Lasiewski and Lasiewski
1967; Lasiewski et al. 1967; Carpenter 1974; Opazo et al. 2005;F
ernández et al. 2011a, b; Fernandez and Suarez 2011; Shankar et al.
2020a
Resting metabolic rate 2 Open-ow,
positive pressure
BMR conditions during
daytime (laboratory)
Prinzinger et al. 1992; Bakken and Sabat 2007
Oxygen consumption and
carbon dioxide production
(including respiratory
exchange rate)
18 Open-ow,
positive pressure
Non-BMR conditions (eld
and laboratory)
Lasiewski 1963a, b; Lasiewski and Lasiewski 1967; Lasiewski et al.
1967; Wolf and Hainsworth 1972; Withers 1977a, b; Schuchmann
1979a, b; Schuchmann et al. 1979; Schuchmann and Schmidtmarloh
1979a, b; Powers 1991, 1992; Prinzinger et al. 1992; Chaui-Berlinck et
al. 2002; Powers et al. 2010; Dick et al. 2020
Oxygen consumption and
carbon dioxide production
(including respiratory
exchange rate)
1Open-ow,
negative pressure
Non-BMR conditions
(laboratory)
ChauiBerlinck and Bicudo 1995
Body temperature
measurement
1Wolf et al. 2020
Nighttime normothermic
metabolic rate
17 Open-ow,
positive pressure
Temperature response, torpor
patterns, energy storage
threshold, ventilation pattern
(eld and laboratory)
Bartholomew et al. 1957; Lasiewski 1963b; Lasiewski and Lasiewski
1967; Lasiewski et al. 1967; Hainsworth and Wolf 1970; Wolf and
Hainsworth 1972; Wolf et al. 1972; Hainsworth and Wolf 1978; Krüger
et al. 1982; Hiebert 1990, 1992;Hiebert 1993a;b; Bucher and Chappell
1997;Powers et al. 2003;Eberts et al. 2019;Shankar et al. 2020a
Total evaporation rate 2 Open-ow,
positive pressure
Water balance, humidity
eects (laboratory)
Powers 1992;Bakken and Sabat 2007
42
Measurement Count Protocol Applications References
Hovering metabolic rate 24 Open-ow,
negative pressure,
mask
Allometry, energy cost, energy
metabolism, metabolic fuel
use, thermogenesis (eld and
laboratory)
Berger and Hart 1972; Berger 1974a, b; Epting 1980; Bartholomew
and Lighton 1986; Suarez et al. 1991; Wells 1993a;b; Chai and Dudley
1996; Chai et al. 1996; Chai et al. 1998; Altshuler et al. 2001; Welch Jr
et al. 2006, 2007; Welch Jr and Suarez 2007; Evangelista et al. 2010;
Fernández et al. 2011a, b; Fernandez and Suarez 2011; Welch Jr 2011;
Chen and Welch Jr 2014; Kim et al. 2014; Groom et al. 2018; Shankar
et al. 2020a
Oxygen consumption and
carbon dioxide production
2Open-ow,
positive pressure,
metabolic chamber
Energy cost (laboratory) Lasiewski 1963b; Schuchmann and Schmidtmarloh 1979a
Respiratory evaporation
rate
2Open-ow,
negative pressure,
mask
Water loss, heat dissipation
(eld and laboratory)
Berger and Hart 1972; Powers et al. 2012
Field metabolic rate 5 Doubly labeled
water
Daily energy expenditure
(laboratory)
Powers and Nagy 1988; Weathers and Stiles 1989; Powers and Conley
1994; Shankar et al. 2019, 2020a
Daily energy intake 1 Measuring food
intake
Daily energy expenditure,
(laboratory)
Tiebout 1991; Tiebout III 1992
a Literature searched until September 2020.
Table 8. (cont.)
Standard open-ow respirometry can be used to
measure metabolic rates in captive hummingbirds in
eld settings (Lighton 2008). Open-ow respirometry
can be used to measure oxygen consumption and car-
bon dioxide production (both measures of metabolic
rate) as well as evaporative water loss. Specic open-
ow congurations that are appropriate for a variety
of metabolic rate measurements have been described
(Lighton 2008).
Open-ow respirometry systems can employ ei-
ther a positive pressure or negative pressure congura-
tion. In a positive pressure conguration, air is pushed
by a pump or compressed airline through a sealed
metabolism chamber containing the hummingbird. A
second airline that bypasses the metabolism chamber
is required for baseline measurements of airstream gas
composition. A single pump or compressed airline can
be divided to provide airow for baseline measurement
and the metabolism chamber, but the owrate must
be measured separately for each line. Baseline and
outgoing chamber airow then empties into a syringe
barrel where the ow can be sampled by the metabolism
system (see below).
Accurately measuring owrate is critical because
errors can lead to large errors in measurement of meta-
bolic rate. Using standard open-ow protocols, water
and carbon dioxide are removed from air owing into
metabolism chambers to keep gas components constant.
This can be accomplished by using absorbents (Dri-
erite®, W. A. Hammond DRIERITE Co. LTD, Xenia,
OH) to extract water and soda lime to extract carbon
dioxide. Some studies omit the removal of carbon
dioxide because its impact on owrate is extremely
low. Alternatively, if both the water and carbon diox-
ide content of the airstream are known, the owrate
can be corrected mathematically. To use this method,
airstream samples must be run through both a carbon
dioxide analyzer and a humidity sensor to measure the
proportional contribution of water and carbon dioxide.
Subsequently, the owrate can be corrected by subtrac-
tion, using published equations (Lighton 2008). This
is a useful approach when working in the eld because
it eliminates the need to transport and store absorbents.
Because the partial pressures of gases in the airstream
change with barometric pressure, owrate is corrected
to standard temperature and pressure (STP). This cor-
rection is done automatically if the owrate is measured
with a mass owmeter. Flowrate also can be measured
using a calibrated rotameter, but the STP correction will
need to be done manually.
In a negative pressure conguration, air is pulled
through a metabolism chamber or mask. One advantage
of this conguration is that a completely sealed cham-
ber is not required as long as the owrate is sucient to
collect all exhaled air from the hummingbird for mea-
surement of respiratory gas exchange (for metabolic
rate) and respiratory evaporative water loss. A com-
pressed airline cannot be used for a negative pressure
system. In addition, pumps must be completely sealed
because the metabolism system must sample air from
the outlet side of the pump. Further, separate pumps
must be used for baseline sampling and each chamber/
mask. All requirements for accurate measurement of
owrate described above for a positive pressure design
also apply to a negative pressure conguration.
Regardless of whether a positive or negative pres-
sure design is used, baseline and chamber outlet air is
sampled from the syringe barrel at the end of the open-
ow system. Air is pulled from the syringe barrel by a
sub-sampling pump. The owrate of the sub-sampling
pump must be less than the owrate into the syringe
barrel to prevent dilution of the sample with ambient
air. The sub-sampled air can then be pushed through
analyzers for measurement of the gas composition. If
water and carbon dioxide are being subtracted to correct
the owrate of the main pump, the system also must
include both a humidity sensor and a carbon dioxide
analyzer in addition to an oxygen analyzer.
Pumps and mass owmeters/controllers can be
purchased from a variety of sources. When purchasing
a pump, consider the maximum ow needs as well as
the altitude at which measurements will be made. In
most pumps, airow is generated by a diaphragm and
the maximum ow will be reduced by lower air density
at higher altitudes. The analyzer manufacturer most
commonly used for open-ow respirometry is Sable
Systems International (North Las Vegas, NV), which
sells analyzers in a variety of congurations, includ-
44
ing all-in-one systems that are easily transportable and
can run for extended periods on batteries. The Sable
Systems eld equipment is also user-serviceable such
that minor repairs can be done in the eld without
voiding the warranty.
Depending on the computer systems being used,
there are several options for data acquisition and analy-
sis software for open-ow respirometry. Commercial
software (Expedata Data Analysis Software, Sable
Systems International) integrates well with computers
that use Microsoft Windows as their operating system.
Warthog software, written by Dr. Mark Chappell at the
University of California, Riverside, and available at no
charge at warthog.ucr.edu, is well suited for Macintosh-
based systems.
Measurements of basal and resting metabolic
rates in captive hummingbirds can be made using the
open-ow respirometry methods described above. Few
studies have measured the true basal metabolic rate
(BMR)—i.e., the metabolic rate during the rest phase
(nighttime for hummingbirds), in the dark, within the
thermoneutral zone, and during the post-absorptive
phase (i.e., after digesting food) (Carpenter and Mac-
Millen 1976; López-Calleja and Bozinovic 1995;
Opazo et al. 2005; Fernández et al. 2011b). This is
likely due to the inherent diculties associated with
measurement of BMR (McKechnie and Wolf 2004),
which are further amplied by the small size of hum-
mingbirds.
Unlike BMR, resting metabolic rate (RMR) is ref-
erenced frequently in the literature. However, RMR has
many dierent meanings because it has been measured
over a wide range of temperatures, during the birds’ ac-
tive phase, and during periods of food digestion. Thus,
even though RMR measurements in hummingbirds
have been reported in a number of studies (Lasiewski
1963b; Lasiewski et al. 1967; Hainsworth and Wolf
1970; Krüger et al. 1982; Powers 1991; Powers et al.
2003), there is no standard methodology or terminol-
ogy for measuring RMR. Therefore, when measuring
and reporting RMR, it is important to clearly dene
what is being measured. Confusion sometimes can be
avoided by using terminology other than “RMR”. For
example, when measuring RMR in Anna’s Humming-
birds (Calypte anna), Pearson (1950) simply referred
to evaluating metabolic rates while birds were “asleep”
(Pearson 1950).
When measuring either BMR or RMR, metabolic
chamber size and owrate should be carefully consid-
ered. Because mass-specic metabolic rates are low
in resting hummingbirds, a lower owrate is necessary
to achieve oxygen/carbon dioxide measurements that
are suciently dierent from baseline to minimize
measurement error. A owrate of 500 mL/min appears
to work well for hummingbirds (Powers et al. 2003;
Opazo et al. 2005). Small chambers are necessary to
reduce the time required to reach an equilibrium value
and to better visualize short-term changes in metabolic
rate. Chamber volumes of 380–1000 mL have been
used successfully for hummingbird species with body
weights ranging from 3 to 18 g (Powers et al. 2003;
Opazo et al. 2005); however, appropriate chamber size
will vary with the body mass of the hummingbird spe-
cies being studied.
An important consideration when measuring
BMR is the requirement that it be measured at night.
One of the challenges of measuring BMR in humming-
birds is that they can safely be fasted for only up to a
couple of hours before being placed in a metabolism
chamber. The authors believe that the best approach is
to allow hummingbirds to ll their crops prior to being
placed in a metabolism chamber, thus providing them
with sucient energy to enter a resting state (Powers
et al. 2003). Furthermore, providing them with a sugar
meal will delay torpor (see section 8.4) and optimize
chances of recording accurate BMR measurements.
With this approach, it is useful to measure both oxygen
consumption and carbon dioxide production. This
allows for calculation of the respiratory exchange
ratio (RER; O2 consumption/CO2 production), which
explains what the hummingbirds are metabolizing for
energy (RER is 1.0 for sugar and approximately 0.7
for protein or fat). When hummingbirds transition to
metabolizing protein or fat, they are post-absorptive
and ready for measurement of BMR. This procedure
can take several hours. At the conclusion of BMR
measurement, the authors prefer not to disturb hum-
mingbirds during their resting phase but rather to
continue tracking metabolic rate for the remainder of
the night. If birds remain normothermic, they can be
returned to a holding cage. However, birds should be
allowed to feed for a period before being returned to
the dark. Alternatively, they can be left in place with
the lights on for the remainder of the night.
Approaches to measurement of RMR vary de-
pending on what the researcher is attempting to mea-
sure. For example, if the goal is to measure perching
metabolic rate, a perch is included inside the chamber
and measurements are made in the light. If the aim is
to measure the metabolic cost of being alert, measure-
ments can be made under BMR conditions but during
the day. Regardless of the measurement conditions,
hummingbirds must be given approximately 1–2 h to
acclimate to the chamber. The most common error in
measuring metabolic rate is impatience.
Measuring nighttime metabolic rate—including
torpor metabolism— in captive hummingbirds can be
accomplished by using the open-ow respirometry
methods described above. Protocol design depends
on the specic topic(s) being addressed in the study.
Classic studies investigating the eects of temperature
on metabolic rates during torpor have used standard
open-ow system congurations (primarily positive
pressure) similar to those described above (Hainsworth
and Wolf 1970, 1972, 1978; Wolf et al. 1972; Krüger
et al. 1982; Hiebert 1990; Bucher and Chappell 1992,
1997). Because the metabolic rate during torpor is ex-
tremely low, owrates as low as 150 mL/min have been
used successfully used to separate metabolic response
from background values (Bucher and Chappell 1997).
However, emerging evidence indicates that torpor
metabolism during scripted experiments may dier
from ways in which torpor is used by hummingbirds
subjected to natural light and temperature conditions
(Shankar et al. 2020b).
Several studies that have addressed the ecologi-
cal importance of torpor in energy management have
used protocols that measured metabolic parameters
under semi-natural conditions of temperature and
photoperiod (Hiebert 1991; Bech et al. 1997; Powers
et al. 2003; Shankar et al. 2020b). Some studies have
refrained from trapping hummingbirds until the end of
the day to allow for normal daytime energy acquisition
and endogenous energy storage and to allow birds to
ll their crops prior to measurement (Powers et al.
2003; Shankar et al. 2020b). To allow for completely
natural temperature and light cycles, Shanker et al.
(2020b) placed metabolism chambers outdoors and
used a negative pressure, open-ow design. Feeder
bases containing sucrose solution were placed in
metabolism chambers to allow hummingbirds to feed
either at the start or end of the metabolic trial. Inclu-
sion of the feeder base necessitated a larger chamber
(approximately 7 L) but the researchers used a owrate
of 500 mL/min, which allowed for measurement of
equilibrium metabolic values in under 30 min. This
approach appears to have yielded evidence of shallow
torpor use, a novel pattern of hypothermia for hum-
mingbirds, and is worth considering if appropriate for
the questions being asked.
Hovering metabolic rate (HMR) and respiratory
evaporative water loss (REWL) can be measured us-
ing a negative pressure system attached to a mask as
described above. Mask respirometry has been used in a
variety of studies involving measurement of HMR (Bar-
tholomew and Lighton 1986; Chai et al. 1998; Welch et
al. 2007; Welch and Altshuler 2008; Powers et al. 2012;
Groom et al. 2018). Mask-based respirometry has been
well-described in the literature (Welch, Jr. 2011). Mask
respirometry can be used to make metabolic measure-
ments both in the eld (Powers et al. 2012; Groom et
al. 2018) or in the laboratory, including during wind
tunnel studies (Clark and Dudley 2010; Powers et al.
2012, 2015; Sapir and Dudley 2012). Wind tunnel
studies allow metabolic measurements during both
forward and backward ight (Sapir and Dudley 2012).
With mask respirometry, the mask is attached to a
feeder so that the hummingbird must insert its head into
the mask to feed. While the bird is feeding, the negative
pressure system pulls respiratory gases exhaled by the
hummingbird from the mask, which are then directed to
open-ow analyzers. The mask should be suciently
large to allow a hummingbird to comfortably insert its
entire head while not obstructing shoulder movement.
If the mask is too long, hummingbirds will have to
overextend to reach the feeder and might attempt to
46
grab the edge of the mask with their feet. The mask
itself can be constructed of anything cylindrical and
transparent; syringe barrels work well. Syringes also
make good feeders for mask respirometry because they
are small and masks can be easily attached. The authors
recommend angling the mask down slightly, which
allows the hummingbird to easily extend its head into
the mask while hovering normally. Regardless of the
feeder/mask combination, it is important to prevent the
sugar solution from dripping into the mask, which can
cause substantial errors in measurement of respiratory
gases. Because HMR can be over 10 times that of
BMR (Suarez 1992; Suarez and Moyes 1992), higher
owrates can be used and are necessary to capture all
exhaled respiratory gases. The minimum recommend
owrate for measurement of HMR is 1500 mL/min in
relatively still air.
Unlike during measurement of BMR, RMR,
and torpor, respiratory gas measurements made dur-
ing mask respirometry are not equilibrium values and
therefore cannot be used to directly calculate HMR
(Bartholomew and Lighton 1986). Instead, total vol-
ume of oxygen consumed, carbon dioxide produced,
or respiratory water evaporated must be calculated by
integrating the area under the metabolic response curve.
Analyses should be performed only during feeding
bouts that are at least three seconds long. Once the
total volume of respiratory gas is calculated, it can be
divided by the time the hummingbird’s head was in
the mask to calculate HMR or REWL. If a feeding
bout involves multiple head insertions, the time of
each insertion must be summed. Two methods have
been used to do this (Bartholomew and Lighton 1986;
Welch and Suarez 2006). The method used most often
commonly employs photoresistors, which—when oc-
cluded—mark the presence of the hummingbird’s head
in the mask. Alternatively, video recordings can be used
to time feeding visits (Powers et al. 2012).
Field metabolic rate (FMR) is a measurement of
the energetic cost of animals living in the wild and is
often used to estimate daily energy expenditure (DEE).
The optimal method for measuring FMR is the doubly
labeled water (DLW) method, which allows direct
measurement of carbon dioxide production in free-
living animals (Nagy 1983; Speakman 1997). In the
DLW method, carbon dioxide production is measured
by marking an animal’s body water pool with isotopes.
A small amount of water containing a high amount of
both deuterium (2H) and oxygen-18 (18O) is injected
into the bird (i.e., the “body water pool”) and the rate
at which these isotopes are eliminated over time (the
turnover rate) is measured. During this process, 18O is
lost from the hummingbird’s body water pool as both
carbon dioxide and water, whereas 2H is lost primarily
as water. As such, the dierence in turnover rate be-
tween the two isotopes is assumed to be due to carbon
dioxide production. Both 18O and 2H are stable isotopes,
so no special permitting is required. A detailed descrip-
tion of the DLW method has been published (Speakman
1997). Understanding this basic theory is important,
particularly as described in Chapter 17 of the cited text.
Although the DLW method works very well for
measuring FMR, it is a challenging technique to use on
hummingbirds. The primary reason that isotope turn-
over rate is typically measured using blood samples and
hummingbirds have a limited blood volume that can be
safely sampled. To date, only three published studies
have used this method on hummingbirds (Powers and
Nagy 1988; Weathers and Stiles 1989; Powers and Con-
ley 1994). However, current methods for measuring
isotopic enrichment require very small samples of body
water so that use of the DLW method on hummingbirds
is now more feasible than it previously was.
To use the DLW method, hummingbirds are
trapped as described elsewhere in this review. Birds are
then weighed to the nearest 0.01 g, and a small amount
of isotopic water (approximately 2 µL/g) is injected into
the muscle. The precise volume may vary depending
on the isotope enrichment of the solution. Depending
on the species, the isotopic water should be injected
using a 50- or 100-µL precision glass syringe (Ham-
ilton Company, Reno, NV) that was tared previously.
Prior to injection, the syringe and its contents should
be weighed to the nearest 0.0001 g. This is important
because the isotopic solution will have a higher mole
mass than non-isotopic water, so conversion of injec-
tion volume to injection mass for calculation of carbon
dioxide production is not trivial. It is important not
to inject more than the indicated volume of isotopic
water. Today’s isotope analyzers are actually better at
detecting relatively lower versus higher isotope enrich-
ment. The current expense of 18O provides additional
motivation to conserve.
After injection, hummingbirds should be allowed
to rest quietly while the injected isotopic water equili-
brates with their body water pool. This usually requires
approximately 45 min (DRP laboratory) (Shankar et al.
2019). During this equilibration period, hummingbirds
are placed in a mesh bag and distanced from human
activity. After equilibration, the initial blood sample
should be collected in a heparinized microcapillary
tube as described elsewhere in this review (see section
4.2). A minimum of 15 µL of blood (approximately 10
µL of water) is generally sucient for use in the LGR
Liquid Water Isotope Analyzer ® (Los Gatos Research,
San Jose, CA), but the amount of blood required may
dier depending on the instrument used. After blood
collection, the capillary tube ideally should be ame-
sealed and refrigerated. A small butane jeweler’s torch
works well for sealing capillary tubes and is easily
used in the eld. In addition to study hummingbirds,
blood should be collected as described above from a
minimum of three injected hummingbirds for each
species being studied. These samples will serve as
background values required in the calculation of carbon
dioxide production. Following blood collection, the
researcher should ensure that the collection site has
stopped bleeding. The hummingbird is then hand fed
a 20–25% sucrose solution to provide energy prior to
release. The bird sampled also must be marked for
recapture identication. If work is being performed at
a site where hummingbirds are banded, band numbers
work well. Otherwise, color marking can be employed.
Use of newly inserted RFID tags is not recommended
because they may impact normal behavior during the
release period. Once released, hummingbirds can then
engage in their normal daytime/nighttime energy use
patterns.
One of the biggest challenges of the DLW method
is recapturing injected birds one day later. Water turn-
over rates are high enough in endothermic animals that
body water returns to background levels after two days
or less; therefore, recapture after being free-living for
24 h after release is important. However, the recap-
ture rate is only about 10% in most studies, although
this rate is highly variable. Hummingbirds appear to
have good memories, and when researchers are using
modied Russell traps, it is often dicult to trap the
same birds two days in a row. An alternative trapping
method, such as mist nets, can be used on the second
day. Once a hummingbird is recaptured, it must be re-
weighed and a second blood sample must be collected.
Afterwards, the hummingbird can be fed and released.
Because of the diculty in getting blood samples
from hummingbirds, it is important to understand that
the DLW method can work using any type of body
water sample just as long as the initial and nal samples
derive from the same source. DLW studies have suc-
cessfully used urine samples, which can be relatively
easy to obtain given the high-water turnover rate of
hummingbirds. Urine collection is best done in the
morning (see section 7). When collecting urine, the
sample should be clear and not grossly contaminated
with fecal material. Once collected, urine samples are
sealed and stored as described above for blood.
Although enrichment of a hummingbird’s body
water pool is always best done by injecting isotope,
feeding isotope in a sugar solution works reasonably
well when injection is not allowed. This is because
hummingbirds absorb and process most of the water
they consume through their kidneys (McWhorter and
Rio 1999). Isotope enrichment by feeding is much
more involved than by injection and requires longer
periods of handling; therefore, this method should only
be used as an option of last resort. When using this
approach, hummingbirds must be precisely weighed
before and after being fed to calculate the amount of
enriched nectar that was consumed. When creating the
enriched nectar, the sucrose and enriched water must
be added on a weight basis. This allows for subtraction
of sucrose from the consumed meal by multiplying the
weight of the consumed enriched nectar by the propor-
tion of the mass that is enriched water. It is important
not to over enrich the hummingbirds. The amount of
isotope added to water used to create the enriched nectar
will depend on how readily the species of interest takes
to hand feeding. The authors use syringes for feeding,
but any type of feeder will work. Note that when using
this method, it is much harder to control how much
isotope the hummingbirds receive because there is no
way to accurately measure consumption.
48
Literature review yielded 39 unique studies that
demonstrated trends in organs sampled, quantities
extracted, and applications (Table 9). Some of the
research applications for sampling tissues include phy-
logenetics (Gerwin and Zink 1998; Chaves and Smith
2011; McGuire et al. 2014; Weinstein et al. 2014), epi-
demiology (Magagna et al. 2019), toxicology (Godoy et
al. 2014; Filigenzi et al. 2019; Graves et al. 2019), and
physiology (Mathieu-Costello et al. 1992). Due to their
small body size and a paucity of techniques for ante-
mortem tissue sampling, carcasses are most commonly
utilized for tissue samples. Fresh tissue samples are
sometimes needed for certain types of modern genomic
testing, such as transcriptome and high-throughput
sequencing. However, many studies do not require
fresh specimens, so older frozen specimens can be
used instead of collecting new specimens. As stated
previously, museums are an ideal starting point for
obtaining samples because museum curators are often
associated with local communities that provide birds
for archiving purposes. Alternatively, academic institu-
tions or research programs that archive specimens for
future studies may be considered. Other options include
licensed wildlife rehabilitators or wildlife rehabilitation
centers, which may have birds that were euthanized or
did not survive the rehabilitation process.
Whether tissue is sampled from live birds or
harvested from deceased individuals, quantities of
available tissue can be less than optimal for analysis.
Therefore, analytic methods may require modication
to adjust for small sample volumes.
Assuming a whole bird has been procured, the
protocol for harvesting may vary depending on the or-
gans to be extracted. However, some general guidelines
can prove helpful. If the bird is to be vouchered as a
museum specimen, the following method for gaining
access to organs is necessary in order to preserve the
carcass. The specimen should be placed on its back
with the ventral surface exposed. The specimen weight,
measured in grams, should be recorded before the bird
is sampled. The breast feathers are parted and a #15 or
#10 scalpel blade is used to make an incision from the
mid-furculum caudally along the keel. The skin and
muscle overlying the abdominal area is then incised
to expose the coelomic cavity. Lateral to the keel on
either side, the ribs are transected from the level of the
abdomen cranial to the coracoid bones. The keel is
then elevated to expose the internal organs.
If the specimen is not going to be archived, the
following dissection technique can be used to obtain
access to the organs for sampling. The feathers are
parted and an incision is made with a scalpel from the
lateral commissure of the beak on either side, to the
ventral midline, at the junction of the neck and head,
then distally along the midline, over the keel, to the
cloaca. The skin can then be bluntly dissected and
pulled away from the keel and pectoral muscles bilater-
ally, exposing most of the pectoral muscles. Then an
incision can be made on each side of the keel, through
the ribs and through the clavicle, bilaterally, allowing
the entire breast plate (keel, pectoral muscles, and
ribs) to be elevated and moved to the side, exposing
the coelomic cavity.
Once the organs are accessible, tweezers or
small forceps can be used to isolate the desired organ
while small surgical scissors are used to dissect it from
the lining of the body cavity. The organ can then be
transferred to storage. Tools should be soaked for 5
min in a 1% sodium hypochlorite solution and rinsed
with distilled water before and between sample col-
lection for purposes of cleaning and minimizing DNA
contamination.
Given the wide range of organ functions and as-
sociated specic research applications, there may be
organ- or application-specic methods for storage and
analysis (Table 9). For example, for genomic banking,
all tissues of interest can be stored in the same cryotube.
However, other research designs might require that
no cross contamination of tissues occur, necessitating
storage of tissues in separate vials.
Table 9. Summary of usage and storage conditions for tissue samples collected from hummingbirds as reported in published studies.a
Tissue Count Amount Applications Storage†References
Brain 9 Not reported Pathology, physiology Formalin or cryoprotected in 30%
sucrose or immersion xed in 4%
paraformaldehyde in 0.1 M phosphate
buer or embedded in gelatin
Iwaniuk and Wylie 2007; Iwaniuk et al. 2009; Godoy
et al. 2013; Reiser et al. 2013; González-Gómez et al.
2014a, b; Gaede et al. 2019; Magagna et al. 2019; Diao
et al. 2020
Heart 9 26–31 mg Energetics, genetics,
pathology, physiology
Formalin or frozen in liquid nitrogen
and then at -70° C or dried at 60° C for
6 days
Gerwin and Zink 1989, 1998; López-Calleja and
Bozinovic 2003; López-Calleja et al. 2003; Cortés-
Rodríguez et al. 2008; Godoy et al. 2013; Reiser et al.
2013; Magagna et al. 2019; Diao et al. 2020
Intestine 821–24 mg Energetics, gut
bacteria, pathology,
physiology
Formalin or frozen in liquid nitrogen
and then at -80° C or dried at 60° C for
6 days
López-Calleja and Bozinovic 2003; López-Calleja
et al. 2003; Preest et al. 2003; Zamparo et al. 2003;
Williams et al. 2012; Godoy et al. 2013; Reiser et al.
2013; Magagna et al. 2019
Kidney 8 3–5 mg Energetics, pathology,
physiology
Formalin or dried at 60° C for 6 days Casotti et al. 1998; Beuchat et. al 1999; López-Calleja
and Bozinovic 2003; López-Calleja et al. 2003; Godoy
et al. 2013; Reiser et al. 2013; Mikoni et al. 2017;
Magagna et al. 2019
Liver 8Not reported Energetics, genetics,
pathology, physiology
Formalin or frozen in liquid nitrogen
and then at -70° C or dried at 60° C for
6 days
López-Calleja and Bozinovic 2003; López-Calleja et
al. 2003; Cortés-Rodríguez et al. 2008; Godoy et al.
2013; Mikoni et al. 2017; Lim et al. 2019; Magagna et
al. 2019; Diao et al. 2020
Lungs 4 16–20 mg
(dry)
Energetics, pathology,
physiology
Formalin or frozen in liquid nitrogen
and then at -80° C or dried at 60° C for
6 days
López-Calleja and Bozinovic 2003; Williams et al.
2012; Godoy et al. 2013; Magagna et al. 2019
Pectoral
muscle
10 Not reported Genetics, physiology Frozen or RNAlater or ethanol or salt
extraction of genomic DNA
Mathieu-Costello et al. 1992; Gerwin and Zink 1998;
Lance et al. 2009; Bailey et al. 2013; Reiser et al. 2013;
Benham and Witt 2016; Prosdocimi et al. 2016; Mikoni
et al. 2017; Magagna et al. 2019; Diao et al. 2020
Other muscle 13 Not reported Genetics, pathology,
physiology
Formalin or frozen in liquid nitrogen and
then at -80°C to -20°C or dried at 60°C
for 6 days
Gerwin and Zink 1989; Cortés-Rodríguez et al. 2008;
Welch et al. 2009; Fernández et al. 2011a, b; Oyler-
McCance et al. 2011; Donovan et al. 2013; Godoy et
al. 2013; Reiser et al. 2013; Velten and Welch Jr 2014;
Benham et al. 2015; Lim et al. 2019; Magagna et al.
2019
Tissue Count Amount Applications Storage†References
Carcass 3 Variable Toxicology, Micro CT
imaging
Frozen -80°C Filigenzi et al. 2019; Graves et al. 2019; Riede and
Olson 2020
Syrinx 1 Variable Micro CT imaging Fixed tissue - 0.1 M phosphate-buered
saline (PBS) in solution with 0.05%
sodium acid
Monte et al. 2020
Tissue (not
specied)
1 Not reported Genetics Museum specimens Hernandez-Banos et al. 2020
a Literature searched until September 2020.
†Tissues xed in formalin should be transferred to paran-embedded blocks as soon as possible (ideally within 2–3 days) for optimal preservation of RNA/DNA for
later molecular testing by immunohistochemistry, in situ hybridization, or polymerase chain reaction.
Table 9. (cont.)
10.1. Trends and Applications
A literature review yielded 23 records of hum-
mingbird muscle samples being collected from speci-
mens. The vast majority of these records reported
sampling of the pectoral muscle (Table 9).
Hummingbird Specimens
Using dissection techniques
and a size #15 or #10 scalpel blade, the entire deep and
supercial pectoral muscles can be harvested from both
sides of the keel.
The authors have found
that a 6-mm dermal biopsy punch (Miltex®, Inc.,
Princeton, New Jersey) is extremely useful for obtain-
ing pectoral muscle samples while limiting damage to
the surrounding tissue. This method is recommended
if the specimen is to be vouchered as a museum speci-
men. In addition to inicting minimal damage on the
specimen, this method allows for immediate harvesting
of tissue irrespective of the need to prepare a museum
specimen. Soaking the biopsy punch in 1% hypochlo-
rite solution for 5 min and rinsing with distilled water
is recommended before and between sampling. The
specimen also can be refrozen for later preparation as
a study skin.
To obtain a muscle sample, the breast feathers are
parted with water or 95% ethyl alcohol, exposing the
skin and underlying breast muscle. The biopsy punch
is pressed downward and rotated so that the circular
blade of the punch penetrates the pectoral muscle. The
instrument is then rolled back and forth between the
thumb and index nger to rotate the blade. This motion
can be continued until the metal tip of the biopsy punch
contacts the keel. A 6-mm diameter punch allows for
a substantial muscle sample to be obtained from hum-
mingbirds while still facilitating tissue extraction from
the instrument and leaving an acceptable defect size
for a museum specimen. Although a 4 mm diameter
punch still provides an adequate sample and produces
a smaller defect, extraction of the sample from the core
of the instrument is more challenging. If the carcass is
to be used as a study skin, a small plug of cotton can be
placed in the muscle defect to protect the feathers from
body uids. Note that the ecacy of dermal punch
muscle sampling depends, in part, on the degree of
carcass autolysis that has occurred prior to sampling.
Autolysis can soften tissue and complicate sample
extraction from the instrument.
The precise methods used to store muscle tissue
depend on the research application. The authors found
10 records in the literature that used pectoral muscle
samples for a range of purposes (Table 9). Most modern
museums store muscle samples in cryovials in a -80o C
freezer or liquid nitrogen tissue bank.
11.1. Trends and Applications
A literature review revealed 17 publications in
which hummingbirds were euthanized or lethal take
was a component of the study and the euthanasia
method was reported. Numerous manuscripts did not
report the method of euthanasia (n=28). In those that
did, the most frequently used method was carbon di-
oxide asphyxiation, followed by overdose of ketamine/
xylazine, along with reports of cervical dislocation and
anesthesia followed by either nitrogen asphyxiation or
ketamine/xylazine overdose (Table 10).
If a hummingbird is injured and has the potential
to recover, it should be taken to a permitted wildlife
rehabilitator, which can range from a private individual
to a center. When an injured hummingbird is deemed
non-releasable (most commonly because of a broken
wing) or if a bird is extremely ill, euthanasia may be
necessary to relieve pain and suering. Guidelines for
the humane euthanasia of animals have been estab-
lished by the American Veterinary Medical Association
(AVMA) (Leary et al. 2020). These guidelines are
required to be updated every 10 years but interim revi-
sions can be made. In order to ensure that researchers
are following the most current guidelines, the AVMA
resources web page should be consulted regularly
(https://www.avma.org/resources-tools/avma-policies/
avma-guidelines-euthanasia-animals).
Depending on the situation and individual bird,
an indirect method of euthanasia (e.g., carbon diox-
ide overdose) versus an active method (e.g., cervical
dislocation) might be less traumatic for the person
euthanizing the animal. Some IACUCs have minimum
requirements for basic training in euthanasia methods.
At minimum, individuals who are hummingbird re-
search site leaders must be trained in active euthanasia
methods if they will be employing them at their site.
Some institutional IACUCs also require training in
indirect methods of euthanasia. Special considerations
for euthanasia of wildlife, as opposed to captive ani-
mals, have been outlined previously (Paul et al. 2016;
Engilis, Jr. et al. 2018). A quick death is preferable to
inducing additional distress by capturing, manipulat-
ing, and transporting an animal to a facility where
euthanasia can be performed. Unacceptable methods
of euthanasia include freezing (ice crystallization of
tissues is very painful), suocation, vehicle exhaust, or
incorrect application of an approved method. Before
working in the eld, the investigator or lead individual
for the team should decide how euthanasia will be
conducted if such action is necessary. If the chosen
method requires special agents or equipment, those—in
addition to a stethoscope—should be essential parts of
the eld equipment. If the euthanasia method does not
require a chemical or gas agent and will be imposed
physically, the BBL permittee or sub-permittee must
be trained prior to being allowed to use the method in
the eld. Euthanizing a hummingbird can be traumatic
for the individual performing the euthanasia and/or for
team members; therefore, pre-emptive conversations
can be helpful.
After a bird has died, tissue samples can be col-
lected, the bird can be frozen and subsequently prepared
as a museum specimen, or the animal can be xed in
10% formalin and submitted for histopathology for dis-
ease assessment and/or cause of death. Most state and
federal permits require dead birds to be deposited in an
accredited museum to ensure long-term research value.
Table 10. Summary methods used for euthanasia or lethal take in published studies involving hummingbirds as study
subjects.a ,b
Method Count References
Anesthesia followed by nitrogen
asphyxiation
1Reiser et al. 2013
Anesthesia followed by ketamine/
xylazine overdose
1Donovan et al. 2013
CO2 asphyxiation 3 Casotti et al. 1998; Beuchat et. al 1999; Preest et al. 2003
Cervical dislocation 2 López-Calleja and Bozinovic 2003; López-Calleja et al. 2003
Sodium pentobarbital 1 Mathieu-Costello et al. 1992
Thoracic (cardiac) compression alone 3 Fernández et al. 2011a, b; Fernandez and Suarez 2011
Ketamine/xylazine overdose without
anesthesia
5Welch Jr and Altshuler 2009; Reiser et al. 2013; González-Gómez et
al. 2014a, b; Gaede et al. 2019
Isourane followed by asphyxiation 1Myrka and Welch 2017
a Literature searched until September 2020.
b Numerous references reported bird euthanasia or lethal take (n=28) without reporting the technique.
Administering First Aid
When assessing the need to conduct rst aid pro-
cedures, the welfare of the hummingbird takes priority.
Individuals working in the eld should learn basic rst
aid procedures to treat minor injuries, such as minor
lacerations that occasionally result from birds being
captured in mist nets. If BBL permittees have received
training in more complex rst aid treatments for birds,
they can conduct those procedures under a BBL permit
as long as the bird is either released or transferred to a
rehabilitator within 24 hr. If birds require rst aid pro-
cedures that are beyond the training of a BBL permittee,
the individual must transfer the bird to a veterinarian
or rehabilitator within 24 hr to receive appropriate
care. First aid treatment by USFWS Migratory Bird
Scientic Collecting permittees is not authorized un-
less the permittee is a veterinarian or permitted wildlife
rehabilitator or the permit includes authorization for
rst aid and treatment of minor injuries (50 CFR §
21.12, 50 CFR § 21.31). USFWS permit applicants
can request authorization to perform rst aid and treat
minor injuries by including a description of their train-
ing and experience along with their permit application.
If a scientist has both BBL and USFWS Migratory Bird
Scientic Collecting permits, administration of rst aid
procedures requires rst aid and minor injury treatment
authorization on at least one permit.
-
If a hummingbird is injured during banding or
tagging activities and the extent of the injury leads the
BBL permittee to believe that euthanasia is the only
appropriate option, euthanasia can be performed by the
individual holding a BBL master or sub-permit. Guide-
lines for assessing whether a bird should be euthanized
are outlined later in this section. The BBL permit does
not provide general authorization to euthanize birds
and should not be used to euthanize a healthy bird. If
a hummingbird is injured during banding activities and
has the potential for recovery, it should be transferred
to a licensed wildlife rehabilitator for treatment.
If a hummingbird has a pre-existing injury or
illness and is captured with the intent of banding, tag-
ging, or sampling the bird (i.e., birds that are captured
for banding and/or sampling but then are discovered
to be injured before the procedure), a BBL permit au-
thorizes permittees to evaluate the condition of the bird
and either euthanize it or take it to a licensed wildlife
rehabilitator or veterinarian for treatment. In cases
of captured birds with pre-existing injuries, the BBL
permittee must rst determine whether the bird should
be released in its current condition. If it appears to be
otherwise healthy and/or able to survive with the pre-
existing condition, the BBL permittee can release the
bird back to the wild with or without further treatment.
If the BBL permittee decides to release the bird, the
individual must decide whether or not to rst band the
bird. Banding birds with pox lesions anywhere on the
body is not advised because lesions can spread to the
distal legs and feet. The BBL does not have a stated
policy for banding birds with a single foot, and the deci-
sion is left up to the BBL permittee (B. Peterjohn, pers.
comm., 28 June 2020). From a welfare perspective,
birds with a single foot should not be banded on the re-
maining leg. Free ranging hummingbirds with one leg
have been known to survive; therefore, this pre-existing
condition might warrant releasing the bird versus tak-
ing it to a wildlife rehabilitator. Similarly, birds with
pre-existing beak injuries should be released unless the
bird’s condition warrants taking it to a veterinarian or
a wildlife rehabilitator authorized to determine if the
bird should be rehabilitated or euthanized (50 CFR §
21.12, 50 CFR § 21.31).
If an injured hummingbird is found by a BBL
permittee out of context of capturing the bird for
banding or if someone brings an injured hummingbird
to a BBL permittee, the BBL permittee is allowed to
conduct limited activities to promote the bird’s welfare.
An example would be allowing a window-strike bird to
recover in isolation. However, if the bird requires rst-
aid treatment before it can be released, it must be taken
to a permitted wildlife rehabilitator or veterinarian for
appropriate care (50 CFR § 21.12, 50 CFR § 21.31).
If a hummingbird is injured during research
activities authorized under a USFWS Migratory Bird
Scientic Collecting permit, the researcher should take
the bird to a licensed rehabilitator or veterinarian for
treatment (50 CFR § 21.12 and 50 CFR § 21.31) if
lethal take is not authorized on their permit. Alterna-
tively, a researcher could proactively request collection
authorization as part of their USFWS Migratory Bird
Scientic Collecting permit application (CFR § 21.23).
Demonstration of training and/or experience with lethal
take and/or euthanasia techniques will be required for
permit authorization. Specically, if a researcher only
possesses a USFWS Migratory Bird Scientic Collect-
ing permit and lethal take of the species is authorized
on the permit, the researcher can euthanize the bird.
However, if the researcher only has a USFWS Migra-
tory Bird Scientic Collecting permit and lethal take
of that species is not authorized on the permit, the
individual must take the bird to a licensed rehabilita-
tor or veterinarian for assessment in accordance with
USFWS Migratory Bird Rehabilitation regulations (50
CFR § 21.31). In that instance, the researcher is not al-
lowed to euthanize the bird. USFWS regulations allow
veterinarians to temporarily care for and stabilize wild
birds without a permit (50 CFR § 21.12). As mentioned
previously, rst aid or minor injury treatment is allowed
by a USFWS permittee such treatment is authorized
on the permit; however, medical treatment beyond the
training of the permittee is not authorized unless the
permittee is a veterinarian or permitted wildlife reha-
bilitator (50 CFR § 21.12, 50 CFR § 21.31).
If a hummingbird becomes moribund in the eld
while a bander and/or researcher is working with it, the
bird should be warmed and oered small amounts of
sugar water before euthanasia is considered. Because
hummingbirds live on the energetic edge, they may ap-
pear to be unrecoverable; however, they often recover
after receiving minimal supportive care. Euthanasia is
the most humane option for birds that sustain compound
wing fractures (i.e., injuries in which a fractured bone
penetrates the skin) or neck fractures during the course
of a banding or sampling event. Hummingbirds that
do not have a wing droop but cannot hover due to soft
tissue wing injuries should be considered for rehabilita-
tion. If given sucient time (sometimes months), these
birds may improve to the point of being releasable.
If a hummingbird has a signicant injury that will
prevent its release and the injury was pre-existing or
occurred during a banding/sampling event, placement
in a zoologic institution or an organization that accepts
non-releasable wildlife might be a consideration. Such
placement requires approval from the local USFWS
Regional Migratory Bird Permit Oce. If the animal is
non-releasable, there are no placement options, and/or
the bird would endure distress and discomfort while be-
ing held captive, euthanasia is the most humane option.
Wildlife rehabilitators or veterinarians are autho-
rized to treat injuries that occur during a banding or
sampling activity and are experienced in determining
whether a bird should be rehabilitated or euthanized
for pre-existing conditions (50 CFR § 21.12, 50 CFR §
21.31). Avian veterinarians are best equipped to work
with hummingbirds, and they can be found at https://
www.humanesociety.org/resources/how-nd-wildlife-
rehabilitator. In addition, wildlife rehabilitators can
be found by state at https://www.humanesociety.org/
resources/how-nd-wildlife-rehabilitator. The fol-
lowing USFWS regulations (50 CFR § 21.31, https://
www.ecfr.gov/cgi-bin/text-idx?SID=0e9fd773e2a078e
4c54d159ce4a9be80&node=50:9.0.1.1.4.3.1.11&rgn=
div8) stipulate which conditions necessitate a wildlife
rehabilitator to euthanize a bird:
You must euthanize any bird that cannot feed
itself, perch upright, or ambulate without
inicting additional injuries to itself where
medical and/or rehabilitative care will not
reverse such conditions. You must euthanize
any bird that is completely blind, and any
bird that has sustained injuries that would
require amputation of a leg, a foot, or a wing
at the elbow or above (humero-ulnar joint)
rather than performing such surgery, unless:
(A) A licensed veterinarian submits a written
recommendation that the bird should be kept
alive, including an analysis of why the bird is
not expected to experience the injuries and/
or ailments that typically occur in birds with
these injuries and a commitment (from the vet-
erinarian) to provide medical care for the bird
for the duration of its life, including complete
examinations at least once a year; (B) A place-
ment is available for the bird with a person or
facility authorized to possess it, where it will
receive the veterinary care described in para-
graph (e)(4)(iii)(A) of the USFWS regulations;
(C) The issuing oce specically authorizes
continued possession, medical treatment, and
rehabilitative care of the bird.
11.4.1. Overdose of inhaled anesthetic agent
Overdose of an inhaled anesthetic can be
the sole euthanasia method or the rst step in a two-step
process (anesthesia followed by euthanasia). For hum-
mingbirds, anesthesia overdose is best accomplished by
soaking a cotton ball with isourane liquid and placing
it in a small container with the bird. The bird should
be left in the container until respirations have ceased
for at least three continuous minutes. Disadvantages
of this method include the logistics required to possess
and maintain a Food and Drug Administration (FDA)
controlled substance and the risk of human exposure
to concentrations above 2 ppm within 1 hr (accord-
ing to the National Institute for Occupational Safety
and Health [NIOSH]). If individuals are transporting
anesthetic agents, such as isourane, to eld sites in a
vehicle, carrying very small volumes (such as two to
three isourane-soaked cotton balls) in a well-sealed
container will minimize accidental human exposure
during transport. A bottle of isourane, which holds
large volumes of the anesthetic, should never be trans-
ported inside the cabin of a vehicle where passengers
are present. Use of isourane is further complicated
if air or highway travel to the eld site is necessary.
Isourane is considered a liquid chemical and is not
allowed in carry-on or checked luggage. Furthermore,
isourane is designated by the US Department of
Transportation as a class 9 agent, and transport is sub-
ject to local, state, and federal regulations. Therefore,
despite its advantages in some situations, there may be
substantial limitations to use of an inhaled anesthetic
in the eld.
11.4.2. Overdose of inhaled carbon dioxide
(CO2).—When using an overdose of inhaled CO2 to
euthanize a hummingbird, a compressed gas canister
should be used to deliver CO2 and gradually ll the
bird’s containment chamber with gas. Gradual (versus
rapid) asphyxiation with CO2 is less likely to be painful
for animals, and the bird will ideally be rendered un-
conscious before gas accumulation causes discomfort.
A gradual ll method at a displacement rate 30–70%
of the chamber volume per minute is recommended
in the AVMA guidelines for euthanasia (Leary et al.
2020). In the eld, a small clear container can be
used as a gas chamber (Fig. 11). The container should
have a very small outlet for oxygen to escape as CO2
is being introduced. Small portable CO2 cartridges, a
regulator intended for delivery of CO2 for carbonating
beverages or sh tanks, and plastic tubing can be used
to deliver CO2. Dry ice should never be used to deliver
CO2 for euthanasia. Gas should be delivered slowly so
the temperature is not drastically reduced inside the
container. To determine the CO2 ow rate, multiply
the chamber length, width and height in inches, then
divide the product by 61 to determine the volume
in liters. Multiply the volume in liters by 30–70%
(0.3–0.7) to determine the proper ow rate (liters per
minute or LPM). For example, if a chamber is 12 in
length x 12 in width x 12 in height the volume is 1728
in or 61 L (1728/61). The proper CO2 ow rate would
be 8 LPM (61 x 0.3) to 20 LPM (61 x 0.7). The bird
should remain exposed to CO2 for at least 3 min after
respiration ceases.
Use of CO2 has many advantages: 1) it is readily
available; 2) it is not an FDA-controlled substance;
3) it is eective over a wide range of concentrations;
and 4) it poses minimal risk to the person euthanizing
a hummingbird when performed properly. Disadvan-
tages to this method are that it may: 1) cause pain; 2)
result in air hunger; 3) elicit a fear response if CO2 is
not administered properly; and/or 4) result in artifactual
histopathologic pulmonary lesions. In addition, CO2
accumulation may cause convulsions, which can result
in tissue damage and may be distressing to observers.
Neonatal hummingbirds may be relatively more toler-
ant of CO2 and may require increased concentrations
(Leary et al. 2020).
11.4.3. Overdose of pentobarbital anesthetic.—A
hummingbird has reduced muscle mass and very limit-
ed vascular access relative to the size of the hypodermic
needle necessary for delivering pentobarbital solution.
Therefore, hummingbirds should be anesthetized with
a gas anesthetic prior to injection with pentobarbital.
Pentobarbital can be injected into the jugular vein,
muscle, or coelomic cavity. Using a Luer lock syringe
and needle will minimize the risk of the needle disen-
gaging from the syringe during injection. Intracoelomic
delivery seems to be the least stressful method of injec-
tion, but the absorption rate is slower. Pentobarbital
should not be administered at too high a dose as this will
induce severe damage to the lungs, liver, heart, blood
vessels, and other tissues, thus interfering with post-
mortem examination. Although pentobarbital injection
is an eective way to euthanize a bird, pentobarbital is
a controlled substance (DEA Schedule II) that can only
be stored and distributed by complying with multiple
rules and regulations. Therefore, individuals must be
vigilant regarding inventory documentation, access
by authorized users, multiple individuals auditing the
inventory, and locking storage containers.
.—
Given the small size of a hummingbird, rapid cardiac
compression (RCC; previously referred to as thoracic or
cardiac/thoracic compression), if performed properly,
can quickly euthanize a hummingbird without the need
to anesthetize the bird rst. However, because RCC
is not included in the AVMA euthanasia guidelines as
a stand-alone method for euthanasia at the time of this
writing, investigators may rst be required to render the
bird unconscious with an AVMA-approved euthanasia
method, such as CO2 overdose. IACUCs at institutions
accredited by the Association for Assessment and Ac-
creditation of Laboratory Animal Care (AALAC) are
required to use AVMA-approved euthanasia methods.
However, investigators can request conditional ap-
proval for the use of RCC as a stand-alone euthanasia
method for hummingbirds citing work published by
Engilis (Engilis, Jr. et al. 2018). Before RCC is used
in the eld, individuals should be trained using a small
laboratory bird species and following Engilis’ published
methods (Engilis, Jr. et al. 2018). For investigators
who are not at AALAC-approved schools, RCC as a
primary method of euthanasia for hummingbirds could
be considered. However, proper training is essential to
minimize pain and suering while the hummingbird is
being euthanized.
Using RCC, an individual directly applies pres-
sure to the heart, which results in obstruction of venous
return and cessation of cardiac output. In many cases,
this results in rapid rupture of the thin-walled regions
of the vena cava or atrium and almost instantaneous
cessation of brain activity and pulse production (Paul-
Murphy et al. 2017). One study (Engilis, Jr. et al. 2018)
explains proper application of RCC for euthanasia of
birds weighing less than 500 g and describes external
cues as a bird progresses toward death. Advantages
of this method include: 1) rapid loss of conscious-
ness and death (Paul-Murphy et al. 2017); 2) yield of
Figure 11. Illustration of a self-made carbon dioxide chamber for
euthanizing a hummingbird in the eld. The paper towel separates
the hummingbird from the tubing that is piped into the container and
minimizes the bird’s access to the bottom of the container.
specimens in optimal condition for use as specimens
for research collections and other purposes; 3) no
requirement for drugs or chemicals; and 4) ready ap-
plication in any eld setting, particularly in situations
where controlled substances cannot be used (Engilis,
Jr. et al. 2018). However, given the small size of a
hummingbird, proper placement of ngers below the
wings from the dorsal aspect can be dicult, particu-
larly if the researcher has large ngers. As a result, the
cardiopulmonary region may not be properly accessed
during RCC. If improperly performed, rapid cardiac
compression may cause signicant suering before
death (Engilis, Jr. et al. 2018).
11.4.5. Cervical dislocation.—Cervical disloca-
tion is an AVMA-approved method of euthanasia (Paul-
Murphy et al. 2017) but should be performed only by
trained individuals. This method is best facilitated by
hanging the hummingbird’s head over a well-stabilized
edge and applying force between the rst cervical verte-
brae and the base of the skull to dislocate the head from
the neck. Like RCC, there is no need for specialized
equipment or anesthetic agent, and if performed prop-
erly, cervical dislocation results in instantaneous death.
However, this method can be dicult to perform if the
researcher is not properly trained. In addition, cervical
dislocation does not provide optimal specimens if intact
birds are needed. Visual aesthetics of this method can
also have signicant negative and emotional impacts
on observers and operators.
Decapitation is an
AVMA-approved method of euthanasia and is humane
if performed properly (Paul-Murphy et al. 2017). The
instrument used for decapitation needs to be sharp so
that the head is severed in one swift cut and the neck
is cut and not crushed. With the right instrument, de-
capitation will quickly induce loss of consciousness and
death. Disadvantages to this method include rendering
the specimen useless if the whole bird is needed and the
potential for signicant negative and emotional eects
on observers and operators. Similar to other methods,
individuals must be trained in this euthanasia method
prior to working in the eld.
For all euthanasia methods, death can be con-
firmed by cardiac auscultation and pupillary light
response. Lack of spontaneous breathing should not
be used as a sole criterion for conrming death. A bird
near death may exhibit shallow, irregular breathing
patterns, which could be interpreted as a lack of spon-
taneous breathing. In addition, the respiratory patterns
of a hummingbird in torpor might be similar to a bird
in a moribund state, which could make it dicult to
discern the two.
11.5.1. Heartbeat.—Cardiac auscultation is es-
sential to conrm death. Use of a pediatric stethoscope
is helpful due to the small size of the instrument’s
diaphragm, but any stethoscope is acceptable. A
stethoscope should be considered essential equipment
for individuals doing any work with hummingbirds.
Learning to perform cardiac auscultation on a live bird
is important prior to using a stethoscope on a dead bird.
This will ensure proper use of the stethoscope and pro-
vide a reference point. Dual-headed stethoscopes have
both bell and diaphragm components. The diaphragm
side of the stethoscope’s chest piece must be engaged
for cardiac auscultation. This is achieved by twisting
the chest piece 180 degrees and hearing a “click” as it
engages. The user can tap each head to see if the bell or
diaphragm component is engaged. Wrapping a thumb
under the stethoscope tubing will help keep it from
rubbing on other surfaces. This will reduce peripheral
noise and allow the user to be more condent that they
no longer hear a heartbeat.
—When
a very bright light is directed into the eye of a live
animal, the pupil will constrict (narrow), indicating a
neurologic response. It is essential to use a very bright
light (i.e., light emitting diode; LED) when testing the
PLR to ensure proper assessment. Upon death, the
pupils will not respond to light and instead will become
permanently dilated.
If a bird dies unexpectedly or has to be euthanized
due to illness or injury, a post-mortem examination
(necropsy) is necessary to determine the cause of death.
Performing a necropsy after a bird dies suddenly and
unexpectedly during handling by a researcher is an
important way of contributing to further knowledge.
The specimen should be submitted to a veterinarian or
veterinary diagnostic laboratory for necropsy. A list
of veterinary diagnostic laboratories and avian veteri-
narians can be found at https://www.aphis.usda.gov/
animal_health/nahln/downloads/all_nahln_lab_list.pdf
and https://www.aav.org/search/custom.asp?id=1803,
respectively. If an avian veterinarian cannot be found,
another veterinarian can be asked to preserve the bird
in formalin and submit it to a veterinary pathologist
who specializes in exotic birds.
Dead hummingbirds must be kept cold (but not
frozen) for up to 72 h after death to minimize autolysis
and optimize the post-mortem examination. Freezing
causes ice crystallization artifacts in the soft tissues,
thus compromising histopathologic evaluation. Ideally,
the specimens should be submitted within 24 h of death.
The feathers should be moistened with water prior to
refrigeration to facilitate rapid chilling of internal tis-
sue. Although autolysis occurs over time, necropsy of
a bird that died up to 72 h before submission may still
yield a cause of death.
As soon as the specimen is submitted for nec-
ropsy, samples for virology, bacteriology, and/or fresh
materials for research studies need to be collected.
Immediately following sample collection, the specimen
should be preserved in 10% buered formalin by the
veterinarian or diagnostic laboratory so that tissues can
be prepared for histopathology. Formalin is a carcino-
gen, and as such, its storage and handling must follow
specic requirements from the Occupational Safety and
Health Administration (OSHA). Personal protective
equipment (e.g., appropriate gloves, eye protection, and
face shield) and a well-ventilated space are required
when handling formalin. The xation process will
render the specimen unusable for skinning, and the
necropsy process signicantly damages the carcass,
making it less suitable as a museum specimen. If a
dead hummingbird is ultimately going to be placed in
a museum, necropsy should be avoided.
To properly x the specimen for necropsy, the
operator should wear laboratory grade gloves and spray
the bird’s external surface with detergent or a dilute dis-
infectant. This helps reduce feather waterproong and
allows the xative to penetrate the skin. After locating
the keel, a pair of clean scissors is used to incise the
skin at the base of the keel. Once the coelomic cavity
is entered, the skin is cut along the distal margin of the
keel. The skin is then peeled away from the pectoral
muscle on either side of the incision. On the lateral
aspect of the bird, the ribs are cut at their junction with
the keel and the keel is reected cranially. The clavicle
is also transected. Any hemorrhage in the body cavity
or gross lesions involving the organs are noted.
The skin on the back of the skull is then reected.
A small cut is made between the skull and the spinal
cord on the dorsal surface of the bird, so that the head is
only attached to the rest of the body by the front of the
skull, skin, and muscles/ligaments. The incision is con-
tinued around the skull, making a “skull cap” that can
be elevated forward to reveal the brain. Opening the
skull ensures that the brain will be properly xed, but
the brain itself should be left in place. The entire speci-
men should then be placed in a container of formalin,
making sure that all the tissue is covered by solution.
Placing a paper towel over the specimen helps to keep
it submerged. The container is then sealed and clearly
labeled. It is important to note that a deceased bird that
has been processed for a post-mortem examination will
be rendered useless for museum specimen vouchering.
Scientic collecting remains an important tool
for researchers studying geographic variation, species-
level classication, 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 previ-
ously, collection of any hummingbird is regulated
through state and federal permitting processes, which
requires specic scientic collecting permission from
the USFWS Migratory Bird Permit Oce in the region
where the investigator lives. Collecting (live or lethal)
permission can be incorporated into federal and state
scientic collecting and research permits that authorize
such activities, but it is not a permit condition granted
by the BBL. The BBL only authorizes salvage collec-
tion (not live or lethal collection) as a permit condition.
Permit limits are placed on the number of individual
specimens and the species authorized for collection
annually.
Two primary methods are used to collect birds:
mist nets and small-caliber shotguns. Hummingbird
traps described in Section 2.3 also can be used to obtain
specimens and might be a better option compared to
mist nets or shotguns, depending on the individual’s
skill level with the latter two options. The proper use
of mist nets for capturing live birds is summarized in
The North American Banders' Manual for Humming-
birds (Russell and Russell 2019). After capture in a
mist net, birds can either be immediately euthanized
in the net or removed and held in a cloth bird bag prior
to euthanasia. Lethal exposure to isourane (section
9.4.1) or rapid cardiac compression (section 9.4.4) can
be used for euthanasia (Engilis, Jr. et al. 2018). A .22
long rie or .410 shotgun are best for collecting small
birds. The Savage Model 24 .22LR/.410 combination
gun (or equivalent) is the preferred gun of choice by
eld researchers collecting small birds. Shot size is
critical to ensure that minimal damage is inicted on
the specimen. For hummingbirds, the .22 caliber long
rie bird shot (25 grain no. 12 shot) is preferred for birds
within a 10-m distance. The .410 shotgun should be
used for birds 20–30 m away. The .410 shotgun shell
should be loaded preferably with number 12 shot but
shot size up to number 9 can be used. Do not use the
.410 shotgun if the hummingbird is closer than 20 m as
the shot can severely damage the specimen.
Many eld researchers process their specimens
in the eld, but specimens also can be processed and
frozen for later work in a laboratory setting. How-
ever, care of any specimen should begin as soon as
it is collected. The collector should have cotton and
absorbent dust available. Cotton should be placed
inside the mouth, so body uids do contaminate the
feathers, especially if the animal was captured at or
near a feeder and the crop is likely to contain sugar
water. For specimens collected by gun, cotton also
should be used to obstruct the mouth so that blood or
ingesta does not contaminate the feathers. However,
if the study requires examination of the tongue or beak
structure, such as the elastic micropump mechanism of
the hummingbird’s feeding apparatus (Rico-Guevara
and Rubega 2011), the mouth should not be obstructed
with cotton. If the bird has been collected by gun, the
specimen should be examined for areas of potential
blood leakage. Absorbent dust should be placed on
those sites to prevent the spread of uids. After cotton
has been placed in the mouth and other uid leaks are
stopped, all feathers should be smoothed on the body,
wings, and tail.
Specimens of any type (including salvage) are
documented with a collecting data sheet that lists, at a
minimum, the location of collection (using descriptive
text as well as latitude and longitude data), the date
of collection, and the collector’s name and aliation.
Many researchers tie a specimen tag of archival paper
that contains this basic information on the leg of the
specimen. All specimens collected should be recorded
in a eld journal for reporting and archival purposes.
Pencil or archival ink pens should always be used to
record data. The Pigma Micron 01, 02, or 03 (Sakura®,
San Francisco, CA) are ideal for eldwork and are acid-
free and archival. Ballpoint pens should be avoided
because the ink can bleed and smear onto a specimen or
data sheet and render the writing illegible, particularly
if the specimen is placed in a freezer.
Once the specimen is cleaned and tagged, it
should be placed in a pre-made paper cone or rolled in
a small tube (Fig. 12A and 12B). Great care should be
taken with the beak so it is not damaged during storage.
Rolling a hummingbird specimen in a tube containing
the data sheet and then securing the tube ends with tape
best protects the fragile bill and specimen in a freezer
(Fig. 12B). The specimen should then be placed into a
sealed plastic bag from which the air has been removed.
For long-term storage (up to 6 months), hummingbirds
should be kept in a conventional freezer. Frost-free or
ultra-cold freezers (i.e., -20° C to -80o C), which cause
rapid desiccation of the specimen, should be avoided.
60
A B
When researchers work with live study subjects
and collect biological samples, standard operating
procedures should be used to avoid disease transmis-
sion between live birds and from birds to humans.
Researchers should avoid secondary transfer of topi-
cal treatments from humans to birds (e.g., sunscreen,
insecticide, or related products) and should be aware
that their shoes, clothing, and bodies (especially their
respiratory tract and hands) can be fomites. Even if all
birds in a work session appeared healthy, equipment
should be cleaned afterward. Recommended protocols
include cleaning equipment with a mild disinfectant,
washing bird holding bags with a dilute (10%) sodium
hypochlorite (bleach) solution, and thoroughly rinsing
any equipment and/or holding bags with water after
disinfection. Indirect transmission of avian pox can
occur whenever hummingbirds are held and sampled.
In addition, virus can be transferred by contaminated
materials through a break in the skin. Therefore, after
working with a bird potentially infected with pox, any
Figure 12. A) Cone method for specimen freezer storage – First recommended method for wrapping hummingbird
specimens for freezer archiving. This method will ensure feathers are not damaged and bill and tail protected. Data
sheets should always be facing away from bird in case ink is used for recording data on the paper tags. B) Tube method
for specimen freezer storage – Second recommended method for wrapping hummingbird specimens for storage in
freezer. This method will also ensure feathers are not damaged and bill and tail protected. It is easier to stack specimens
using this method compared to cone method. Data sheets should always be facing away from bird in case ink is used
for recording data on the paper tags.
A B
instruments or materials that came in contact with the
bird should be set aside for disinfection. Any scab
material shed from lesions on the bird should also be
cleaned from the work surface. Pox virus is highly
resistant to drying and can survive for months to years.
A 10% sodium hypochlorite solution prepared within 4
h before use is optimal for decontaminating surfaces,
instruments, and bird holding bags. Important consid-
erations for using sodium hypochlorite as a disinfectant
are detailed by the World Health Organization (WHO
2014). Sodium hypochlorite germicidal cleaners and
wipes are also made by various manufacturers. Al-
though they have not been tested for decreasing the
transmission of avian pox, they could potentially be
used to reduce contamination.
Zoonotic diseases (diseases that spread between
animals and humans) are a key consideration when
obtaining samples from hummingbirds. To the authors’
knowledge, zoonotic disease transmission has not been
documented between hummingbirds and humans. West
Nile virus has been detected in hummingbird tissue
samples but an insect vector is required for transmission
to humans. Regardless, prevention of disease transmis-
sion by conducting best practices should be a priority
for any eldwork. These practices include good hand
hygiene, no consumption of drinks/food in the work
area, no co-washing of feeders with human dishes,
no recapping of hypodermic needles, and disposing
of needles and/or syringes directly into a laboratory-
grade sharps container or a sharps container approved
for medical use. If a bird goes into respiratory arrest,
human mouth-to-beak assisted ventilation should not
be performed. Even though diseases such as avian
inuenza have not been reported in hummingbirds,
people should limit risks of exposure.
When working with hummingbirds, bees and
wasps are another concern due to their attraction to
sugar water. Therefore, team members may experience
stings, which—in some cases—may result in severe,
even life-threatening allergic reactions (anaphylaxis)
requiring emergency treatment. As a precaution, all
team members should have an antihistamine, such as
diphenhydramine, available. Although antihistamines
are available over the counter, individuals should be
responsible for bringing their own medications that
have been procured for their personal use. Knowing
the location of the nearest emergency hospital is also
integral to an emergency action plan.
This review is intended to serve as a guide for
scientists interested in pursuing or expanding research
involving hummingbirds, with an emphasis on welfare
issues that are unique to these birds. The authors pres-
ent general considerations and baseline practices that
are important when working with live hummingbirds as
study subjects. As research involving this unique fam-
ily of birds grows, guidelines for minimizing subject
risk and optimizing animal welfare while furthering
scientic methods should be constantly reviewed and
expanded.
The authors thank Dr. Bruce G. Peterjohn for
his careful and thorough review of this manuscript;
especially the permitting sections, Peter Pyle for his
input on best practice guidelines for feather sampling,
and Deanna L. Johnson (UC Davis librarian) for her
invaluable assistance with literature searches. We also
thank Drs. Eric Schroder and Connie Orcutt for their
meticulous copy editing. Line drawing illustrations
were provided by Kathy West Studios. Support was
provided by The Daniel and Susan Gottlieb Founda-
tion, Mr. and Mrs. Thomas Jeerson, and the UC
Davis Museum of Wildlife and Fish Biology. Special
recognition goes to the live bird study sites (TS Glide,
The Gottlieb Native Garden, UC Davis Arboretum),
Dr. Manfred Kusch, and UC Davis Hummingbird
Health and Conservation Program volunteers who
have graciously supported hummingbird research and
have helped rene these methods. Sta and students
62
of the Museum of Wildlife and Fish Biology provided
assistance, both in the eld and laboratory. Finally,
immense appreciation and gratitude is extended to Dr.
Holly Ernest who founded the UC Davis Hummingbird
Health and Conservation Program and inspired many
of us to care about hummingbirds and commit to their
scientic study.
The ndings and conclusions in this article are
those of the author(s) and do not represent the views
of the US Fish and Wildlife Service. References made
to commercial products, processes, or services by trade
name, trademark, manufacturer, or otherwise, do not
necessarily constitute or imply their endorsement,
recommendation, or favoring by the US government.
The views and opinions of authors expressed herein do
not necessarily state or reect those of the US govern-
ment and shall not be used for advertising or product
endorsement purposes.
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Addresses of authors:
School of Veterinary Medicine
University of California, Davis
Davis, CA 95616 USA
latell@ucdavis.edu
Biological Sciences Department
Hayward, CA 94542 USA
School of Veterinary Medicine
University of California, Davis
Davis, CA 95616 USA
rbandivadekar@ucdavis.edu
Sacramento, CA 95825 USA
University of Connecticut
Storrs, CT 06269 USA
austin.reid.spence@gmail.com
Biology & Chemistry Department
George Fox University
dpowers@georgefox.edu
Department of Pathobiology and Diagnostic
Investigation
Michigan State University Veterinary Diagnostic
Laboratory
College of Veterinary Medicine
Lansing, MI 48910 USA
agnewd@msu.edu
Department of Pathology, Microbiology and
Immunology
School of Veterinary Medicine
University of California, Davis
Davis, CA 95616 USA
and
California Animal health and Food Safety
Laboratory
University of California, Davis
Davis, CA 95616 USA
lwwoods@ucdavis.edu
Biology
University of California, Davis
Davis, CA 95616 USA
