Hematology at A Glance

Critical Care Management
for Laboratory
MICE
and RATS
F. Claire Hankenson
A Volume in The Laboratory Animal

Pocket Reference Series
A Volume in The Laboratory Animal Pocket Reference Series

for Laboratory
MICE
and RATS
The Laboratory Animal Pocket Reference Series
Series Editor
Mark A. Suckow, D.V.M.
Freimann Life Science Center
University of Notre Dame
South Bend, Indiana
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Critical Care Management for Laboratory Mice and Rats

The Laboratory Canine
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for Laboratory
MICE
and RATS
F. Claire Hankenson
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For Kurt and Shug, who provide me with
critical care, supplemental warmth, and
essential support (dosed daily, ad libitum)
contents
preface..........................................................................................xiii
acknowledgments.......................................................................... xv
about the author.......................................................................... xvii
1 general approaches for critical care.........................................1

overview..................................................................................1
obtaining a clinical history and medical records......................3
body condition scoring............................................................5
relocation for physical examinations........................................6
monitoring critically ill rodents................................................6
pain recognition and assessments......................................... 11
supportive care for surgical procedures.................................12
supplemental heat provision.................................................. 17
drug therapy.........................................................................18
Routes of Drug and Fluid Administration........................18
references.............................................................................. 21
2 critical care management for laboratory mice........................25

introduction..........................................................................25
overall assessments...............................................................25
general medical approaches to physical examination and
health assessments...............................................................26
Physical Examination......................................................27
Body Condition Scoring...................................................29
Clinical Assessments of Ill Health and Pain in Mice.........29
Monitoring Frequency...................................................... 31
Objective Scoring Systems............................................... 31
vii
viii contents
veterinary care measures......................................................33

Administration of Fluids..................................................33
Blood Sampling...............................................................35
Body Temperature Monitoring..........................................38
Endotracheal Intubation..................................................40
Injections and Oral Administration.................................. 41
Urine Sampling................................................................42
abnormal, critical, and emergent conditions..........................44
Abdominal Swelling.........................................................44
Abscessation....................................................................45
Cage Flooding with Subsequent Hypothermia..................47
Cannibalization...............................................................49
Conjunctivitis..................................................................50
Cross Fostering of Neonates/Mouse Pups......................... 51
Dystocia..........................................................................52
Fight Wounds..................................................................56
Fractures/Orthopedic Problems.......................................57
Hemorrhage.....................................................................59
Moribund, Weak, or Paralyzed Condition.........................59
Mortality (Sudden Death)................................................. 61
Ocular Lesions.................................................................62
Perineal Swelling.............................................................64
Poor Body Condition........................................................66
Rectal Prolapse................................................................68
Respiratory Distress........................................................69
Seizures...........................................................................71
Trauma............................................................................ 74
Ulcerative Dermatitis.......................................................76
Vaginal or Uterine Prolapse..............................................84
research-related medical issues.............................................85
Ascites Production...........................................................85
Experimental Autoimmune Encephalomyelitis Mouse
Models.............................................................................87
Hemophilic Mouse Models................................................88
Obese Mouse Models........................................................88
Opportunistic Infections in Immunodeficient Mouse
Models.............................................................................90
Radiation Exposure......................................................... 91
Streptozotocin Induction for Diabetic Models...................94
Tumor Burden in Mouse Models.......................................94
Urogenital Disease in Mouse Models................................98
contents ix
euthanasia............................................................................99
references..............................................................................99
3 critical care management for laboratory rats....................... 113
introduction........................................................................ 113
overall assessments............................................................. 114
general medical approaches to physical examination
and health assessments...................................................... 114
Physical Examination.................................................... 114
Body Condition Scoring................................................. 117
Clinical Assessments of Ill Health and Pain in Rats....... 117
Monitoring Frequency.................................................... 119
Objective Scoring Systems............................................. 119
veterinary care measures....................................................120
Administration of Fluids................................................120
Blood Sampling.............................................................122
Body Temperature Monitoring........................................125
Bone Marrow Access......................................................125
Endotracheal Intubation................................................125
Injections and Oral Administration................................128
Urine Sampling..............................................................129
abnormal, critical, and emergent conditions........................ 131
Burns............................................................................ 132
Catheter Infections........................................................ 133
Malocclusion (Incisors) and Caries.................................134
Moribund/Weak/Paralyzed............................................ 135
Ocular Lesions Secondary to Anesthesia....................... 137
Poor Body Condition...................................................... 137
Ringtail......................................................................... 139
Ulcerative Dermatitis..................................................... 139
Urolithiasis.................................................................... 141
research-related medical issues........................................... 142
Arthritis Models............................................................. 142
Cranial Implant Maintenance........................................ 143
Incontinence Secondary to Spinal Cord Injury Models...... 145
Middle Cerebral Artery Occlusion in Rat Models of
Stroke............................................................................ 145
Obese and Diabetic Rat Models...................................... 146
Opportunistic Infections in Immunodeficient
Rat Models..................................................................... 147
Pododermatitis............................................................... 147
x contents
Spontaneously Hypertensive Rat Models........................ 148

Tumor Burden in Rat Models......................................... 148
euthanasia.......................................................................... 150
references............................................................................ 151
4 special considerations for critical care management

in laboratory rodents........................................................... 159
introduction........................................................................ 159
aging animal model considerations...................................... 160
blood loss considerations..................................................... 162
chronic indwelling device considerations............................. 163
depilatory cream considerations.......................................... 163
equipment considerations for rodent surgery
and emergency procedures..................................................164
euthanasia considerations................................................... 165
experimental autoimmune encephalomyelitis
and demyelinating disease model considerations................. 168
EAE Scoring.................................................................. 168
Animal Care.................................................................. 169
fasting considerations.......................................................... 170
fluid therapy considerations................................................ 171
Crystalloids................................................................... 172
Colloids.......................................................................... 173
food and fluid regulation procedures................................... 173
humane or “clinical” endpoint considerations...................... 175
nutritional therapy considerations....................................... 178
perioperative care considerations......................................... 181
regulatory considerations.................................................... 186
restraint collar considerations............................................. 187
tracheostomy considerations............................................... 188
tumor development and monitoring considerations.............. 189
Evaluating Tumor Growth..............................................190
Tumor Ulceration........................................................... 191
Multiple Tumors............................................................. 191
Ascites Produced by Tumors.......................................... 191
wound management considerations..................................... 192
references............................................................................ 193
5 resources and additional information.................................. 199

introduction........................................................................ 199
organizations....................................................................... 199
contents xi
American Association for Laboratory Animal

Medicine (AALAS).......................................................... 199
American College of Laboratory Animal Medicine
(ACLAM)........................................................................200
American Society of Laboratory Animal Practitioners
(ASLAP).........................................................................200
Institute for Laboratory Animal Research (ILAR)............ 201
publications......................................................................... 201
Books............................................................................ 201
Periodicals.....................................................................203
electronic resources.............................................................203
AALAS Learning Library...............................................203
Animal Care Training Services (ACTS)..........................204
AVMA Guidelines for the Euthanasia of Animals:
2013 Edition..................................................................204
CompMedTM listserv.......................................................204
Drug Enforcement Agency (DEA), Office of Diversion
Control..........................................................................205
IACUC-Forum listserv....................................................205
International Council for Laboratory Animal
Science (ICLAS)..............................................................205
International Mouse Strain Resource (IMSR)..................207
Mouse Genome Database...............................................207
National Institutes of Health Office of Research
Infrastructure Programs (ORIP): Rodent Resources........208
Office of Laboratory Animal Welfare...............................208
Pubmed.........................................................................208
Rat Genome Database....................................................209
TechLink listserv...........................................................209
Veterinary Bioscience Institute (VBI)..............................209
Veterinary Emergency and Critical Care Society
(VECCS)........................................................................ 210
Veterinary Information Network (VIN)............................ 210
commercial resources.......................................................... 210
ALN Buyer’s Guide......................................................... 210
Lab Animal Buyer’s Guide............................................. 211
appendix A: glossary of acronyms and terms........................... 213
appendix B: s
uggested medical supplies for rodent
critical care........................................................... 215
appendix C: rodent formulary................................................... 217
preface
Progressive scientific and medical advances rely on appropriate
selection and study of animal models. In contemporary biomedical
research facilities, almost without exception, laboratory mice and
rats are examined for optimum health status prior to acceptance
as research models. Therefore, common ailments (i.e., respiratory
tract infections, diarrhea, and overt skin disease) are often grounds
to refuse admittance into an existing animal research colony. If
laboratory rodents develop these ailments after acceptance into an
experimental study, they most likely will be treated by extrapola-
tion of applicable clinical regimens employed in small animal and
exotic pet veterinary practice. Unfortunately, for more complex cases
that need critical care management, many treatment approaches to
veterinary emergencies cannot be applied directly to the laboratory
rodent.
For critical care of laboratory rodents, there is a scarcity of sources
that provide comprehensive, feasible, and response-oriented informa-
tion about clinical interventions specific to spontaneous and induced
models of disease. This textbook is the first of its kind devoted to
the challenges of critical care management for laboratory mice and
rats. This text was designed to serve as a specialized resource for all
persons and professionals who utilize rodent models for biomedical
research. The information herein emphasizes the varied approaches
to laboratory rodent patient care, health assessments, characteris-
tics of specific disease models, monitoring and scoring of disease
parameters, and humane interventions. Although preservation of
animal health and welfare is of primary consideration, achievement
of proposed research objectives should also be prioritized.
xiii
xiv preface
Within the field of laboratory animal practice, guidance and

recommendations for clinical treatments are rarely uniform across
the entire profession. The majority of the dosages and treatments in
this book are derived from published abstracts presented at national
American Association for Laboratory Animal Science (AALAS)
conferences (dating from the year 2000 to 2012), journal articles, and
personal consultation and experience. Many treatment modalities are
based on anecdotal information, pilot work, and empirical e vidence.
In the spirit of incorporating continued refinements into laboratory
animal practice, these treatment guidelines may be m odified further
for additional clinical support and improved patient-based care for
research animals.
This textbook is organized as five chapters: “General Approaches for
Critical Care” (Chapter 1), “Critical Care Management for Laboratory
Mice” (Chapter 2), “Critical Care Management for Laboratory Rats”
(Chapter 3), “Special Considerations for Critical Care Management
in Laboratory Rodents” (Chapter 4), and “Resources and Additional
Information” (Chapter 5). In addition, three appendices include a
glossary, medical supply list, and an abridged rodent formulary. As
endorsed by the Guide for the Care and Use of Laboratory Animals
(National Research Council, 2011), the expectation is for those clini-
cal recommendations herein to be applied with apt professional
judgment, including review of evidence- and performance-based out-
comes, while advancing animal welfare and science.
F. Claire Hankenson, DVM, MS, DACLAM

acknowledgments
I would like to thank so many of my colleagues in comparative
medicine, all of whom challenge the status quo to identify refine-
ments in laboratory animal practice on a daily basis. The following
individuals participated in discussions and information and

photograph exchanges during the course of this project: Drs. Valerie
Bergdall, Iris Bolton, Gillian Braden, Angela Brice, Ralph Bunte,
Andrew Burich, Adam Caro, Anthony Carty, Scout Chou, Laura
Conour, Dawn Dinger, Ramon Duran-Struuck, Nancy Figler, Trish
Foley, Margaret Fordham, Joanna Fried, Diane Gaertner, Travis
Hagedorn, Lisa Halliday, Troy Hallman, Lori Hill, Michael Huerkamp,
Marc Hulin, Samer Jaber, Bambi Jasmin, JanLee Jensen, Kari
Koszdin, Paul Makidon, Jim Marx, Tom Meier, Angela Mexas, Emily
Miedel, Guy Mulder, Stu Leland, John Long, Judy Nielson, Jane Olin,
Kate Pritchett-Corning, Jennifer Pullium, Jamie Rhodes, Karen
Rosenthal, George Sanders, Kim Saunders, J. Mat Schech, Pat
Sharp, Abigail Smith, Jennifer Smith, Peter Smith, Laike Stewart,
Doug Taylor, Christin Veeder, Susan Volk, Ashley Wathen, Wendy
O. Williams, Jolaine Wilson, and Norm Wiltshire.
My sincere appreciation extends to the staff at the national
American Association for Laboratory Animal Science (AALAS) office,
particularly John Farrar for his ready assistance with providing

materials presented throughout the text. Kimberly Bowe assisted
with gathering relevant resource materials, Karena Thek contributed
background material for rodent interventions, and Carrie Childs
Maute provided advice and comic relief, for which I am indebted.
And finally, this work would not have been completed without
Dr. Mark Suckow and John Sulzycki and their collective insights,
encouragement, flexibility, and patience.
xv
about the author
F. Claire Hankenson, DVM, MS, is the senior associate director in
University Laboratory Animal Resources, University of Pennsylvania,
Philadelphia, and is an associate professor of laboratory a nimal
medicine in the Department of Pathobiology at the School of

Veterinary Medicine. Dr. Hankenson obtained her veterinary degree
from Purdue University. Following veterinary school, she completed
her laboratory animal medicine residency and graduate work (MS,
microbiology) at the University of Washington, Seattle. She became
a Diplomate of the American College of Laboratory Animal Medicine
(ACLAM) in 2002. Following several years on faculty at the University
of Michigan in the Unit for Laboratory Animal Medicine, she tran-
sitioned to the University of Pennsylvania. Dr. Hankenson’s current
position combines administrative service, clinical effort, teaching
duties, and collaborative research. Her own research studies involve
investigations of refinements in the care and use of laboratory
rodents, particularly blood sampling, tail biopsy evaluations, and
humane endpoints. Dr. Hankenson has been active on committees
in the American Association for Laboratory Animal Science (AALAS)
since 2002, has served on the board of directors for ACLAM, and is
an ad hoc consultant to AAALAC, International.
xvii
1
general approaches
for critical care
overview
The characterization of animal models is becoming increasingly
sophisticated. Animal species that reliably mimic human disease
provide critical insights so that causative mechanisms can be
understood and lead to the development of novel drugs, diagnostic
procedures, and therapies. In the ardent hope to advance the clini-
cal practice of contemporary rodent medicine, in housing facilities
largely free of infectious diseases, individualized patient care is often
prioritized over traditional herd (colony) health diagnostic approaches.
As a subsequent benefit, improvements in individual animal health
will augment overall colony health measures. Within these colonies,
breeding success, production of offspring, and prolonged good health
are invaluable. The continued commitment to financing biomedical
research and maintaining vast colonies of laboratory rodent models
is due to the sheer variety and prevalence of those with unique and
irreplaceable genetic backgrounds.
Genetic engineering of laboratory animals is fueled by the desire
to unravel the mystery of disease based on contributions by single
and multiple genes, molecules and events associated with physiology,
development, and function. This dynamic scientific area holds promise
for development of new mouse and rat strains that rapidly contribute
to and further define applications of in vivo models for research
programs (Croy et al., 2001). Biomedical researchers should be aware
that both inherent and induced mutations may result in unexpected
phenotypes and disease syndromes; thus, genotype and secondary
1
2 critical care management for laboratory mice and rats
influences on the functionality of the animal’s immune system must

always be considered, along with housing, husbandry, experimental
treatments, age, gender, and body condition, when formulating
clinical treatment plans. Increased susceptibility of l aboratory rodents
to environmental variances, nosocomial bacteria, and infectious
diseases will ultimately require heightened v eterinary care. Oversight
by veterinary specialists, in collaboration with scientific investigators,
should ensure that all procedures for disease prevention, diagnosis,
and therapy are those currently accepted in veterinary and l aboratory
animal practice (National Research Council, 2011).
Assessment of clinical abnormalities in rodents may be difficult
and complex due to the stoic nature of these species. Exposures to
stressors (environmental or otherwise) should be kept to a mini-
mum; rodents have an innate response to escape from a perceived
“predator,” which can result in harm to themselves and a release of
systemic catecholamines that can predispose these species to respi-
ratory and cardiac arrest and severe pulmonary, cardiac, and renal
hypertension (Morrisey, 2003). Therefore, if the ultimate objective is
to maintain the highest standard of animal welfare, then outcomes
and treatments that maintain physiologic stability are ideal.
In rodents that are predisposed highly to stress, the r apidity of the
assessment and subsequent patient stabilization are e ssential prior to
complete evaluation for definitive diagnoses of c linical abnormalities
(Hawkins and Graham, 2007). Supporting a patient (“supportive
care”) on initial presentation can be life saving, particularly with the
prompt provision of oxygen, fluid therapy, and warmth (Doneley, 2005).
Murine intensive care units (M-ICUs), akin to an ICU in v eterinary
clinical practice, are extremely rare to n onexistent in the b iomedical
research environment; therefore, treatment plans for c ritical case
management must realistically factor in the u navailability of 24-h
healthcare monitoring capabilities.
Undesirable infectious pathogens and parasites that enter facilities
and affect rodent colony health should always be included in lists of
diagnostic differentials, but details on treatment and eradication of
these specific agents are left primarily to other reference manuals
(Danneman et al., 2012, Hrapkiewicz and Medina, 2007, Jacoby
et al., 2002, Kohn and Clifford, 2002, Sharp and Villano, 2012). Of
course, no reference text can substitute for specialized instruction
and advanced training in animal physiology, handling, and medi-
cine. Routine consultation with veterinary care professionals is
strongly encouraged, particularly for any challenging or unusual
clinical cases in laboratory animal species.
general approaches for critical care 3
obtaining a clinical history and medical records

Although the individual signs can vary between species and patients,
in general, health status assessments are comprised of a combi-
nation of: appearance, performance, productivity, and exhibition
of appropriate and species-specific behaviors (Clark et al., 1997a,
1997b, 1997c, 1997d). Obtaining a thorough clinical “history” of
the animal will be essential to integrate and analyze the relevant
information necessary for critical care diagnoses and interventions,
similar to what would be expected for human and pet medicine
(Aldrich, 2005).
Individual medical records can be created to provide information
about the condition being reported, potential research interactions
with proposed treatments, action expected from the investigational
team and timeframe for expected response, treatment options, and
detailed instructions on treatment administration (Couto et al.,
2003). Specifics about the laboratory rodent patient database ideally
should include the following:
• Genetic background, strain, or stock

• Gender
• Date of birth (age)
• Body weight (BW; grams)
• Food and water provisions (type, amount)
• Caging and bedding type
• Breeding status (nulliparous, multiparous)
• Microbiological status (specific pathogen free vs. known
Helicobacter positive, etc.)
• Enrichment provisions for exhibition of species-specific
behaviors (nontoxic, disposable cotton padding, plastic tubing)
• Housing arrangement (single housed, paired, multiple cage
mates) or if new cage mates have recently been introduced
• Housing parameters of the room: temperature, humidity,
illumination, noise, vibration
• Whether experimentally naïve or if experimental treatments
were provided
• Any adverse outcomes anticipated for the anticipated phenotype
• Recent surgical or nonsurgical procedures
• Administration of anesthesia or analgesia (type and time

when delivered)
• Transportation or exposure to new housing areas
• Timeline of clinical abnormality (development, progression)
• Corrective measures (treatment plan) taken as a result of
variation from normal health or behavior
• Doses, routes, and frequency of administration of any addi-
tional drugs or medications
• Resolution of the clinical abnormality (either corrected,
healed, or potentially removed from study)
Assessments of the critical patient must be comprehensive, orga-

nized, focused, and efficient (Figures 1.1 and 1.2) to identify as many
of the patient’s problems in the order of their importance for survival
(Aldrich, 2005).
Case Presentation
Date: Facility:
Species: Sex: Strain: Vendor:

Naive: Animal on Protocol:
Protocol Title:
Brief Clinical History:
Problem including differential diagnoses:
Diagnostic work-up: (X-ray; CBC, Chemistries, Urinalysis, Skin Scraping, Fur Pluck, Anal
tape test, Serology, Viral Culture, Culture and Sensitivity, etc.)
Progress and/or changes in treatment based on diagnostic results and/or poor progress:
Prognosis:
Necropsy/Histopathology:
Fig. 1.1 Clinical history (medical record) template for laboratory ani-
mal case management. CBC = complete blood count. (Modified from
the University of Pennsylvania, ULAR.)
Sick Animal Report

Date: / / Reported by:
Facility: Room: Cage Card # _Species: Sex:
PI Protocol # Number of affected animals /
Name of lab contact Phone #
Problem:
FIGHTING( Separate animals) SKIN LESION(not fighting) Ventral Dorsal
PROLAPSE (rectum, penis, or vagina) Place on Alpha-Dri
TUMOR ON BACK greater than penny size or ulcerated.
TUMOR other than on back any size, OR multiple tumors
OTHER, please describe
EMERGENCY*** Call clinical staff immediately
DEHYDRATED/HUNCHED (place 5 moist food pellets on cage floor)
DYSTOCIA TRAUMA/BLEEDING
MORIBUND PAINFUL BREATHING PROBLEM
CIRCLE AFFECTED AREA ON DIAGRAM TO THE RIGHT:
Mark affected area with an “X”
***REPORT EMERGENCIES TO THE CLINICAL STAFF IMMEDIATELY***

1. WEEKDAYS: Veterinary Technicians @ phone number XXX-XXX (M-F 7:30 am to 4:30 pm)
2. WEEKEND: Veterinary Technicians ALL Facilities @ phone number XXX-XXX (Sat-Sun 7:30 am to 4: 30 pm)
3. AFTER HOURS: Call Emergency Service to reach Vet On-Call @ phone number XXX-XXXX
Fig. 1.2 Sick animal report template for initial assessment performed
cage-side. PI = primary investigator. (Modified from the University of
Pennsylvania, ULAR.)
body condition scoring

Methods to develop body condition scores (BCSs) have been
described in a wide variety of domesticated, herd, and laboratory
animal species. The BCS serves as a semiquantitative method of
assessing body fat and muscle (see relevant sections on this topic
in Chapters 2 and 3). Generally, the BCS is independent of BW and
frame size yet can provide a consistent approach to health assess-
ments from appropriately trained personnel. Overall appearance
and body condition score is typically ranked on a scale from 1 (ema-
ciated) to 5 (excessive body fat), with an expectation that an animal
categorized as BCS 3 is of appropriate size and species-specific con-
formation. Extremes in BCS may correlate with, or be predictive of,
certain disease conditions. Animals with the lowest BCSs on the
scale (BCS 1–2) typically are frequently monitored for any poten-
tial humane interventions necessary to improve welfare; however,
it is not uncommon that the lowest BCS will result in veterinary
recommendations for removal from experimental procedures due to
a nimal welfare concerns.
relocation for physical examinations

Initial assessments of laboratory rodents will typically be performed
within the original housing rooms. This assessment is usually with
the housing cage removed from the rack and subsequently opened
within the room’s biosafety cabinet, laminar flow hood, or benchtop
area. It is recommended that veterinary treatments be brought into
the housing room, if at all possible, for application and administration.
These types of veterinary tools can include various topical ointments
(may be premixed or aliquoted into single-use vials), cotton-tip appli-
cators, bandage materials, and sterile fluids, needles, and syringes.
If animals need more intensive care or require anesthesia for
administration of the treatments, animals should be gently moved
to a housing cage without cage mates or into a disposable trans-
port container to transfer them to a dedicated procedure area or
laboratory environment. Cages should be gently manipulated, with
the preference for hand carriage, to avoid excessive jostling (e.g., as
would happen with placement on a wheeled cart) or the possibility of
dropping a cage en route to the destination.
Access to food and fluid will be necessary if the animals are away
from these supplies for longer than 1–2 h; in a transport situation, a
few softened feed pellets and nutritional gel supplements may be the
best option for rapid administration and placement onto the holding
cage floor of the ill animal.
monitoring critically ill rodents

Critical care management occurs when a clinical condition (see
Table 1.1), whether spontaneous, idiopathic, or experimentally induced
in laboratory rodents, may develop toward severe impairment, pain,
distress, further injury, and potential for death.
Some of these situations may be more appropriately referred to as
emergencies but will still require assessment, consultation between
veterinary staff and investigators, a formulated treatment plan, and
medical intervention (see Table 1.2). A representative list of examples
of laboratory rodent emergencies necessitating immediate attention
should include
• Blood or blood-stained discharges from any orifice

• Dehydration
Table 1.1: Diagnostic Differentials Based on Clinical Conditions

Observed in Laboratory Rodents
Clinical Condition Differentials for Consideration
Abdominal distention Fat, ascites, splenomegaly, hepatomegaly, neoplastic mass,
urinary tract obstruction, enlarged bladder, genital gland
abscessation
♀ only: uterine enlargement/pyometra/mucometra,
pregnancy
Abdominal pain Gastroenteritis, urinary tract obstruction, peritonitis,
abscess
Diarrhea Gastroenteritis, parasites, gastrointestinal neoplasia,
nonsteroidal anti-inflammatory drug (NSAID)
administration
Nasal/ocular discharge Conjunctivitis, corneal ulcer, glaucoma, respiratory tract
disease, distress
Pallor Shock, anemia (blood loss, potential gastrointestinal loss,
leukemia, other neoplasia, marrow suppression,
coagulopathy, chronic disease)
Ptyalism (salivation) Malocclusion, oral wound, drug reaction, foreign body
obstruction
Rectal prolapse Gastroenteritis, parasites, Helicobacter, Citrobacter,
secondary to parturition or tenesmus
Seizures Hypoglycemia, toxicity, trauma, central nervous system
mass, neoplasia, subdural bleeding, metabolic
disturbance, experimental induction, expected
phenotype
Stranguria Urolithiasis, cystitis, fight wound contracture and
obstruction, neoplasia, sex gland abscessation, bladder
atony, paresis/paralysis
Weakness (posterior) Systemic disorders (hypoglycemia, anemia, hypoxia),
myelopathy and muscle wasting (trauma, disk disease,
neoplasia, infectious)
Weight loss Gastroenteritis, neoplasia, cardiac disease, dental disease
Source: Modified from Ivey, EI, and Morrisey, JK. 1999. Vet Clin North Am Exot Pract
2:471–494.
• Dystocia
• Hypothermia (cold to the touch)
• Limb weakness or paralysis
• Moribund state
• Postsurgical complications, like dehiscence of incisions
• Prolapses of eyes/urogenital organs/tissues
• Rapid weight loss, emaciation and dehydration over 24–48 h
• Respiratory distress
• Seizures that are unrelenting
Table 1.2: Rodent Health A bnormalities, with Emergencies

Highlighted for Situations that Necessitate Immediate Clinical
Intervention and Consultation with Veterinary Staff
Behavior/Activity Tumors Respiration
Hyperactivity Tumor greater than 2 cm Gaspinga
Lethargic on back Rapid breathinga
Head tilt Tumor/growth anywhere Labored breathinga
Circling movements else on body
Tremors/twitching Multiple tumors Oral/Nasal
Uncoordination Ulcerated tumora Staining
Seizures Salivation
Continuous convulsionsa Limbs/Joints Nasal discharge
Swollen limb Incisor broken
Appearance/Condition Lameness Malocclusion
Infection Self-mutilationa Swollen muzzle
Discolored legs, feet, ears, Missing limba Frothy discharge
and tails Loss use of limb/paralysisa Discolored gumsa
Distended abdomen
Rough hair coat Eyes Feces/Urine
Hunched posture Cloudy eye(s)a Diarrhea
Tail missing Moist discharge Bloody feces
Tail swollen Dry, crusty discharge Feces absent
Weight loss Red-stained eye(s) Discolored urine
Moribund (near death)a Sunken eye(s) Urine absent
Profuse bleedinga Partially closed lids/
Cold to toucha squinting Urogenital and Anal
Eye not visible Region
Skin and Hair Coat Eyelid(s) red/swollen Prolapsed rectum
Skin swelling Scratching/rubbing at eyea Prolapsed penis
Scabs/fight wounds Prolapsed eyea Vaginal discharge
Skin flaking Discharge from penis
Skin discoloration Ears Prolapsed vagina/uterusa
Ulcerative lesion Scabbed ear Dystociaa
Trauma Swollen ear
Stained hair coat Discolored ear
Dehydration/skin tentinga Discharge from ear
Torn ear
Source: Courtesy of the University of Pennsylvania, ULAR.
a Indicates emergency.
• Self-mutilation
• Tumor burden that is ulcerated or interferes with mobility
In a critical care situation, the initial goal is to keep the rodent

patient conscious with appropriate thermal, fluid, and nutritional
support. The key approach is to stabilize the critical patient and not
to determine the immediate diagnosis (Mader, 2002). It is entirely
possible to inadvertently and irreparably harm a rodent patient

by trying to perform too many procedures in succession, and the
struggle of the patient during these events (e.g., venipuncture, cys-
tocentesis, or radiography) may be distressful and potentially fatal
(Paul-Murphy, 1996).
Clinical evaluation and assessment of critical care parameters
should be performed in any sick rodent; particularly, the tenets
of a irway/breathing/circulation (the ABCs of critical care) can be
checked by observing respiratory rate and perfusion of tissues
(e.g., color of ear tips, gums, rectal mucosa) in rodents (Flegal and
Kuhlman, 2004). To better relax and temporarily restrain a sta-
bilized rodent patient for overall physical assessments, one may
consider placing the animal under anesthesia (preferably isoflurane
or a similar agent delivered by nose cone connected to a vaporizer);
keep in mind that the influence of anesthesia will potentially bias the
objective clinical data points to be collected.
Relevant questions, similar to those asked in veterinary clini-
cal practice (Aldrich, 2005), should be reviewed on initial physical
examination of the critical laboratory rodent patient, such as the
following:
• Is the patient conscious?

• Is the patient trying to breathe?
• If not, ventilate if possible.
• Assess mucous membrane color, which should be pink/red
and not purple/blue (indicating a lack of oxygen).
• If the patient is trying to breathe, is air moving in and out of
the lungs?
• Observe thoracic wall movement.
• Auscultate breath sounds.
• Is the heart beating effectively?
• While the heart can be palpated for activity by gently
holding the thorax between thumb and forefinger, manual
assessment of beats per minute is extremely difficult.
• Auscultation can be performed using a pediatric stethoscope.

• Is there any disabling condition (trauma, neurologic injury)
that may affect outcome?
• Is the patient in shock?
• Assess
−− Decreased mental awareness.
−− Unpigmented mucous membrane color.
−− Capillary refill time (assess using rectal mucosa in
mice and rats): equivalent to the time it takes for blood
to return to the capillary bed after one compresses
t issue with a fingertip (Aldrich, 2005).
−− Relative heart rate (as pulses may be impossible to
detect in small rodents).
−− Colder extremities versus warmer central temperature
differences (Lichtenberger, 2007).
−− Consider potential for disseminated intravascular
coagulation, a complex systemic disorder involving the
generation of intravascular fibrin and the consump-
tion of procoagulants and platelets; thrombin ulti-
mately potentiates the coagulation cascade and leads
to small- and large-vessel thrombosis, with resultant
organ ischemia and organ failure.
• Is the animal dehydrated?
• Gently lift and pinch the skin over the back just behind the
scapula and let it return to its resting position; s
lowness
of return is correlated to various degrees of dehydration,
expressed as percentages of BW as follows:
−− marked speed of skin return: hydrated
−− moderate speed of skin return: 5–7% dehydrated
−− mild speed of skin return: up to 12% dehydrated
• Is the animal actively bleeding?
• Is the animal in pain?
• Consider various behaviors that can indicate pain, includ-
ing vocalization when handled, attempting to bite or
escape, an abnormal posture or ungroomed appearance,
and lack of movement within the cage or interacting with
cage mates.
pain recognition and assessments

Guidance on assessments of types of procedures in laboratory rodents
that result in painful outcomes has been reviewed (Kohn et al., 2007).
Those procedures described (see Table 1.3) are commonly requested
in rodent IACUC (Institutional Animal Care and Use Committee)
protocols and should be considered on a continuum of potential pain
and distress. Be aware that certain disease models that may also
induce pain, like those that may induce inflammation or neoplasia,
are described throughout the remainder of the text for assessments
and interventions. The list is not to be interpreted as exhaustive, and
the categorizations are only guides to assist in individual cases using
professional judgment and collaboration between veterinary staff
and research teams. Most, if not all, of the procedures listed are to
be conducted with the animal under sedation or general anesthesia.
Animals in critical condition should be assessed for indicators
of pain, particularly when they are actively involved in proposed
research projects. Assessing pain and subsequent management of
Table 1.3: Categorization of Painful Procedures Performed in the

Course of Research with Laboratory Rodents
Mildly to Moderately Moderately to Severely
Minimal to Mildly Painful Painful Painful
Catheter implantation Minor laparotomy Major laparotomy/organ
incisions incision
Tail clipping Thyroidectomy Thoracotomy
Ear notching Orchidectomy Heterotopic organ
transplantation
Superficial tumor Caesarean section Vertebral procedures
implantation
Retro-orbital sinus blood Embryo transfer Burn procedures
collection
Superficial Hypophysectomy Trauma models
lymphadenectomy
Ocular procedures Thymectomy
Multiple intradermal
antigen injections
Intracerebral electrode
implantation
Vasectomy
Vascular access port
implantation
Source: Kohn, DF, Martin, TE, Foley, PL, Morris, TH, Swindle, MM, Vogler, GA, and
Wixson, SK. 2007. J Am Assoc Lab Anim Sci 46:97–108.
pain may be challenging; in particular, one must be aware that each

animal will respond differently to painful stimuli (Klaphake, 2006).
Due to the considerable variability in pain responses, it is important
that pain assessment be performed by clinicians and skilled per-
sonnel with a comprehensive knowledge of the normal b ehavior and
appearance of the species and particular animal patient of concern
(Miller and Richardson, 2011). Successive observations by a single
experienced observer are likely to provide the best insight into the
resolution of pain and clinical improvement of the critical patient.
Unfortunately, to compound the challenge of the individual conduct-
ing the clinical observations, the sheer presence of the observer,
coupled with the application of certain treatments, may also have an
impact on animal behavior and muddy clinical interpretations.
Acute pain, from a known cause like an injury or surgical proce-
dure, should be treated with analgesia. Chronic pain may be associ-
ated with subtle physiological changes that may be more challenging
to effectively manage in the absence of identification of an inciting
cause. Pain scales (Table 1.4) have been developed to address the
unique needs of many different species, based on species-specific
behavior, and should be developed for individual animal models
with input from the veterinarian and IACUC and in accordance
with standards of the National Research Council (NRC) guidance
(Carbone, 2012, NRC, 2011, Stasiak et al., 2003).
supportive care for surgical procedures

Surgical modeling in laboratory mice and rats may include intra-
cerebral cannula implants, heart and lung transplants, coronary
artery ligations, stroke induction, device placements, radiotelemeter
insertion, hepatectomies, and manipulations of other major organs.
Refinements in surgery methods in rodents include the use of less-
invasive (nonmidline) incisional approaches (Chappell et al., 2011)
and laparoscopic procedures.
Prior to initiation of surgical procedures, it is imperative to e
valuate
whether the patient is at an appropriate plane of adequate anesthesia
and to monitor a variety of parameters to best assess the patient’s
physiologic status (Morrisey, 2003). These parameters can include
the following:
• Depth of anesthesia: Assess by direct visualization, gauging

the degree of muscle relaxation, reflex response, and response
Table 1.4: Template A ssessment for Pain E valuation in Rodents

Criteria Score Definition
Record BW and monitor 0 BW maintained or ↑, normal food
food/water intake and consumption and urine/fecal output.
urine/fecal output Baseline BW is scored as 0.
1 BW change is minor (loss of < 5% from
baseline BW).
2 Animal food consumption ↓ and water
consumption variable (loss of 10–15% from
baseline BW). Altered urine/fecal output.
3 Little to no food/water intake (loss of > 20%
from baseline BW).
Appearance 0 Animal “normal.” Hair coat smooth, lies flat,
with sheen. Eyes clear, bright, open, no
discharge. Posture and movements/
ambulation are appropriate for a healthy
animal.
1 Lack of grooming apparent. No other marked
changes.
2 Hair coat roughened; overall hunched
appearance. Eyes and nose may have
discharges or porphyrin (red) staining.
3 Hair coat very roughened. Ungroomed.
Abnormal posture. Eyes pale, sunken;
closure of lids.
Measurable clinical signs: 0 RR (regular, even) and BT are within
evaluate baseline physiologic norms. Limbs and feet warm.
respiratory rate (RR); Mucous membranes normal (gums and
record body temperature anus pink); extremities normal (ears and
(BT) after all cage feet pink).
manipulations 1–2 BT may be changed by ±1–2°C; RR ↑ by up
to 30% (rapid, shallow breaths; more
abdominal effort).
3 BT changes greater than ±2°C; RR ↑ by 50%
or RR markedly ↓ with little visible effort.
Unprovoked behavior: 0 Normal behaviors (exploring cage, grooming,
determine by cage-side feeding); BAR.
evaluation only 1–2 Abnormal behavior; less BAR; inactive when
activity expected (at feeding times, at night).
Guarding potentially painful area (limbs,
abdomen). Twitching, lameness.
3 Unsolicited vocalizations, self-mutilation,
grinding teeth, chattering, salivating,
restless, or immobile. Unresponsive, quiet.
Behavior responses to 0 Behavioral responses normal for expected
external stimuli (conduct conditions, like being restrained for
this assessment last) physical assessment.
(Continued)
Table 1.4: (Continued) Template A ssessment for Pain E valuation in

Rodents
Criteria Score Definition
Behavior responses to 1 Shows minor depression or minor
external stimuli exaggeration of responses.
2 Shows moderate signs of abnormal
responses; may be behavior changes (more
aggressive or more docile).
3 Animal may overreact to external stimuli,
may have weak responses, or be
nonresponsive.
Source: Modified from Kirsch, JH, Klaus, JA, Blizzard, KK, Hurn, PD, and Murphy, SJ.
2002. Contemp Top Lab Anim Sci 41:9–14.
Scoring: If a subscore of 3 is recorded more than once, then all subscores of 3 are to be
given an extra point (3 becomes 4). Any subscore of 3 is potentially serious, and
one should always attempt to determine the underlying cause or explanation
(e.g., recently dosed with opioid, recovering from anesthesia). Consult veterinary
staff if any subscores of 3 are recorded.
BAR = Bright, alert, responsive.
to stimulation (toe pinch); note that the palpebral response is

lost early in a surgical plane of anesthesia, so stimuli that are
more painful may be required to ensure that no response is
elicited, particularly prior to initiating a surgical incision or
procedure.
• Measurement of oxygen saturation of hemoglobin (or pulse
oximetry): Equipment is available commercially for rodents
but may not be on site during an emergency situation. If there
are apparent oxygenation or ventilation concerns, corrections
will need to be instituted (i.e., intubation, repositioning the
animal to maximize lung field expansion during respiration,
or tracheostomy and assisted respiration).
• Cardiovascular performance: Specialized electrocardiographic
equipment is available commercially (but may not be on site
during an emergency) for rodents to measure electrical activ-
ity of the heart, including atrial depolarization, ventricular
depolarization, and repolarization.
• Blood pressure: Typically, indirect blood pressure measure-
ments are obtained in smaller lab animal species; the pres-
sure cuff is placed snugly around a limb or tail over an
artery, and the monitor inflates the cuff past the point that it
occludes blood flow. In general, mean blood pressure values
over 60 mmHg or systolic blood pressures over 100 mmHg
indicate adequate tissue perfusion.
(A)
(B)
Fig. 1.3 (A) An anesthetized adult mouse in preparation for a pro-

cedure to be conducted. The funnel-shaped face mask is attached
to the non-rebreathing apparatus. A downdraft table is used to
capture waste anesthetic gases, and a thermogenic pack is used
for supplemental heat. (Reprinted with permission from Macmillan
Publishers Ltd. Yardeni, T, Eckhaus, M, Morris, HD, Huizing, M,
and Hoogstraten-Miller, S. 2011. Lab Anim (NY) 40:155–160.) (B) The
housing cage and anesthetic induction chamber can also be placed
in a biosafety cabinet to conduct brief procedures that require seda-
tion of the animal.
Pre-, peri-, and postoperative supportive care should include per-

formance of procedures in a designated area (Figure 1.3) and provi-
sion of sterile supplies and equipment, heat through a source similar
to those described (see relevant section on this topic), sterile warmed
fluids, palatable calorie supplements, and analgesics. It is critical
to ensure that any surgical draping allows for adequate view of ani-
mals to optimize patient monitoring. Animals should be monitored
until they are fully conscious from anesthesia and exhibiting clinically
normal behaviors.
Following surgery, it is imperative to manage potential pain and

monitor anesthetic recovery, which is typically the responsibility of
the research staff approved to perform the procedures. Postoperative
pain in rats and mice has been shown to reduce food and water
consumption; therefore, amelioration/prevention of anorexia is espe-
cially important (Flecknell, 2001). Studies have assessed the impact
of socially housing rodents versus individually housing a nimals
postoperatively; it has been shown that rats subjected to spinal cord
injury have a 20% less chance of survival when housed individually,
whereas socially housed female mice need less time to fully recover
from telemetric implant surgery (Van Loo et al., 2007). The Guide
for the Care and Use of Laboratory Animals (NRC, 2011) emphasizes
the importance of social housing of social species; therefore, every
effort should be made to socially house rodents after they have recov-
ered from anesthesia or experimental procedures. Keep in mind that
male mice may need to be housed singly; many strains are known
to display behaviors suggestive of anxiety and overt aggression if
exposed to unfamiliar male mice.
In the immediate hours and days postoperatively, trained person-
nel should frequently observe and closely monitor rodents during the
recovery period, particularly to assess changes in appetite, dehy-
dration, lethargy, or abnormal healing of surgical sites (Hoff et al.,
2006). Normal circadian rhythms, as assessed in telemeterized
rodents, may also take several days to return to preprocedural levels
(Weinandy et al., 2005).
Once an animal has regained consciousness, the residual effects of
many anesthetics may persist for up to 48 h, resulting in decreased
food and water intake and prolonged ataxia (Flecknell, 1987). It is
useful to record BW before and after anesthetic procedures as weight
losses are typically seen during the anesthetic and recovery periods
in rodents. BW may take several days to return to preanesthetic levels,
even if food and water consumption appears within normal limits
(Hayton et al., 1999, Lawson et al., 2001). It is often recommended
to provide food and water immediately after recovery, including solid
pellets on the cage floor and nutritional gel supplements, to prompt
appetite following anesthesia. Typically, BW will decrease from base-
line levels following anesthetic events accompanied by surgery, with
a return to baseline of presurgical weights occurring within 2 weeks
after the procedure (Van Loo et al., 2007). Scoring of body condition
(as described previously and in forthcoming chapters) is a helpful tool
to monitor the pre-, peri-, and postanesthesia appearance and health
of rodents (Easterly et al., 2001, Ullman-Cullere and Foltz, 1999).
Analgesics may be necessary at scheduled intervals for several

days, even up to a week or more, after the procedure. Analgesic d osing
based on rodent BW tends to be relatively high due to their fast metab-
olism and relatively small size. Dose rates of opioids given orally are
particularly elevated due to the considerable first-pass m etabolism
by the liver for these drugs. Dosing will vary at times, often based on
strain; therefore, patients should be observed prior to and following
administration of analgesics (Miller and Richardson, 2011). Further
discussion of these topics is provided in other chapters.
supplemental heat provision

Rodents have relatively high metabolic rates compared to many larger
domesticated species. Due to the high surface area to relatively low
body mass ratio in rodents, heat loss and the development of hypother-
mia (colder core temperatures) are to be avoided. It is extremely impor-
tant to provide thermal support at the start of anesthetic induction
and continuing throughout full recovery, from anesthesia to regained
consciousness (Gardner et al., 1995, Wixson et al., 1987). Body heat
is lost rapidly, and this loss is accelerated when fur is removed or
clipped and liquid disinfectants (particularly a lcohol-based ones)
are applied; in fact, animals undergoing prolonged procedures may
exhibit up to 8% loss of core body heat during just 90 min of general
anesthesia or prolonged sedation in the absence of appropriate sup-
portive care. Placement of laboratory rodents directly on surface bench
tops or metal examination tables is not recommended as these can act
as a heat sink and can further chill an already-hypothermic rodent. Use
of clean disposable diapers or sterile towels is preferred prior to plac-
ing the rodent patient on the examination surface.
To provide additional and supplemental warmth, warm-water
recirculating blankets have been highly effective (Caro et al., 2012),
and microwaveable or automated thermogenic gel packs, reflective
foils, surgical drapes, and heating lamps can be used. Open-style
incubators (ThermoCare®, Incline Village, NY) and Bair Hugger ®
blankets (Arizant Healthcare, Eden Prairie, MN) are more sophis-
ticated heating methods that can be part of an intensive care/post-
operative area in laboratory a nimal settings. For more impromptu
methods of warming, histology slide tray warmers can be adapted in
the laboratory setting; as well, the use of disposable ThermaCare®
activated hand warmers (which can later be placed in the recovery
cage itself) and protective latex/nitrile gloves filled with warm water
can be placed near animals to provide thermal support. Care should

be taken to ensure that animals do not become too warm as hyper-
thermia can occur easily once heat sources are provided due to the
vasodilatory effects of most a nesthetics (Morrisey, 2003).
For monitoring of temperatures in rodents, there are limited
options available that will provide accurate assessments. Microchip
transponder systems that remotely read subcutaneous temperatures
have been tested (Hankenson et al., 2013) and were adequately simi-
lar in readings to traditional rectal thermometry (Chen and White,
2006, Kort et al., 1998) and intra-abdominal telemetry (Vlach et al.,
2000). Noncontact infrared industrial thermometers and temporal
artery thermometry have been tested on rodents but may not be con-
sistently representative of core body temperatures achieved by other
methods; human pediatric ear thermometers may be of use in larger
rats, but are too large for the ear canal of most laboratory mice.
The targeted temperature to which a rodent should be warmed is
95–100°F. Keeping the body temperature as constant as possible from
onset to end of anesthesia has been linked to reduction in recovery
times by 20–30 min (Luboyeski, 2008).
drug therapy
Administration of therapeutic treatments (drugs) is an integral
aspect of critical care medicine. Laboratory animal patients need
to be evaluated carefully prior to treatments for individual factors
(whether experimentally induced or spontaneous) that can affect
the distribution and metabolism of therapeutic drugs (Hackett and
Lehman, 2005). It is important to be aware of the types of drugs
administered, particularly with respect to the potential interactions
they may have with other provided treatments. It will be e ssential to
have a plan for the appropriate dose, frequency, route, and unique
patient factors that might influence drug dosages (Hackett and
Lehman, 2005). Often, for rodent patients in particular, injections are
preferred to best deliver experimental and pharmaceutical agents.
Understanding the different injection techniques, with regard to rate
of delivery as well as volume limits, is important when choosing the
appropriate injection method.
Routes of Drug and Fluid Administration

The routes of drug and fluid administration are shown in Figure 1.4.
IM IV SC ID
Fig. 1.4 Differing routes of administration of substances, including

intramuscular (IM), intravenous (IV), subcutaneous (SC), and intra-
dermal (ID) routes. (Reprinted with permission from AALAS. Turner,
PV, Brabb, T, Pekow, C, and Vasbinder, MA. 2011. J Am Assoc Lab
Anim Sci 50:600–613.)
• Subcutaneous (SC). This is one of the most common sites for

drug and fluid administration in laboratory rodents. The
excess skin over the back of the rodent between the shoul-
der blades (scapulae) accommodates boluses of injectate; vol-
ume overload is less likely with subcutaneous injections than
other administration routes.
• Intraperitoneal (IP). Injections of fluid bolus into the peritoneal
cavity are routinely performed in laboratory rodents. The nee-
dles should be directed toward the lower left or right quadrant,
with a retraction of the plunger to ensure that gastrointesti-
nal structures and the bladder have not been inadvertently
penetrated. Fluids should be warmed when administered to
avoid induction of hypothermia following bolus. Advantages
of intraperitoneal injection include rapid absorption of large
volumes of fluid.
• Oral (PO). Hypertonic solutions and solutions of high caloric
value are often delivered by syringe placed into the diastema
(space between the incisors and premolars/molars) for the
animal to ingest. Oral gavage can be performed, but this may
unnecessarily stress the critical patient. Small volumes are
essential, with 0.05 ml often the limit per dose of what can be
administered to the reluctant and critical patient (Klaphake,
2006).
• Intravenous (IV). Tail vessels are typically the best route in
rodents for intravenous access; however, the time needed
to successfully place and secure (to prevent chewing by the

patient) an intravenous catheter should be balanced against
the need for immediate fluid delivery. Typically, in veteri-
nary practice, the intravenous access is the most critical for
emergencies; however, this is routinely difficult to access in
clinically healthy mice and rats and more challenging in the
ill rodent, which may already be volume contracted.
• Intraosseus (IO). Intraosseus injection of fluid permits rapid
vascular access to bone marrow but should only be used in
very small animals.
Note: Unless the patient is critical, very small, severely dehydrated,
and hypothermic and has no accessible vessels, an intraosseus cath-
eter should be the last choice for fluid administration (Mader, 2002).
Intraosseus catheters are painful, are difficult to keep patent in
small animals, can cause peripheral damage during placement, and
can serve as a nidus for development of osteomyelitis. Common injec-
tion sites include the tibial tuberosity, femur trochanteric fossa, wing
of the ileum, and the greater t rochanter of the humerus.
• Intramuscular (IM). Intramuscular injections of bolus fluids

are not recommended due to the limitations of muscle mass.
For rats, injection of very small volumes into the m
usculature
of the tongue can be rapidly absorbed due to preferential
circulation to the head and brain during shock. On the

contrary, certain emergency drugs, like epinephrine, may

require intramuscular administration. Anterior hind leg
muscles should be avoided to prevent the risk of inadvertently
harming the sciatic nerve (Klaphake, 2006).
Alternative drug routes for mice and rats include intratracheal

instillations (diluted doses of drugs like epinephrine, atropine, lido-
caine, and dexamethasone delivered by catheter into the endotracheal
tube or instilled directly via tracheostomy), gingival injections, and
as a modified suppository per rectum (Hackett and Lehman, 2005).
Intracardiac injections are appropriate only in cardiac arrest situations
when no other method of drug administration has been successful.
Training videos are available online to review these approaches
(http://w w w.jove.com/video/2771/manual-restraint-common-
compound-administration-routes-mice; Machholz et al., 2012).
Properties of the agent injected must also be considered to ensure it
does not cause irritation if given via the selected technique. Certain
injection sites may require restraint of animals or general anesthesia
of the animal. All rodent patients under critical care should be moni-
tored closely for adverse drug effects. Further discussion of these
topics and the reference formulary for this text are provided in the
final chapters and appendices.
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2
critical care management
for laboratory mice
introduction
The continued demand for laboratory mice as biomedical models of
disease necessitates the refinement of diagnostics and treatments
for this species. Further, many health conditions and unique strain-
specific behaviors in mice can be monitored and managed for improved
animal and overall colony health (Bothe et al., 2005). Development
of mouse strains and maintenance of experimental models require
significant investments of research funds and intellectual capital
put toward specific medical model discovery and progress. Therefore,
emphasis will be placed on means to promote longevity of individual
animals in lieu of postmortem diagnostics, if at all possible. General
information about working with laboratory mice is best reviewed in
the companion text, The Laboratory Mouse (Danneman et al., 2012).
Further background information on strains, stocks, and genotypes
can also be obtained by visiting the originating vendor source web-
sites; additional resources are highlighted in Chapter 5.
overall assessments
When assessing a laboratory mouse, it is essential to obtain as much
information as possible about the animal and its use in research to
gain the greatest portfolio of information prior to finalizing differential
25
diagnoses. The complement of information (the clinical and

experimental “history”) should include the background strain of the
mouse; the origin (vendor and pathogen status); any genetic manip-
ulations (which may manifest as phenotypic abnormalities); gender;
age; and specifics about experimental manipulations. When consid-
ering impacts on animal health in the research setting, knowledge
of physical factors like phenotype, body weight (BW), body condition
score (BCS), and potential illness related to the model are paramount.
Environmental factors may also be a confounding factor for
research studies and a great influence on health, specifically factors
in the macroenvironment (e.g., housing room) and those in the
microenvironment (e.g., the housing cage). Macroenvironmental

influences include light cycle and intensity, noise or vibration expo-
sure, and fluctuations in room temperature and humidity. In the
microenvironment, the type and volume of diet and water sources,
frequency of cage changing and types of bedding substrates, and fluc-
tuations in humidity, temperature, and air changes within the cage
are to be considered with respect to maintenance of animal health. It
has been shown, particularly in static housing cages, that the tem-
perature and humidity can be significantly elevated for p rolonged
time periods following autoclaving and heat sterilization practices
(Ward et al., 2009). Therefore, precaution should be taken to ensure
a nimals are placed in cages with bedding approximating the cage
ambient temperature to avoid exposures to heat and hyperthermia,
which can be rapidly fatal.
general medical approaches to physical

examination and health assessments
When presented with a critically ill mouse, to minimize stressors on
the animal, it may be best to prioritize actions and not try to tackle
everything clinically at once. Breaking up the diagnostic and treat-
ment steps into smaller stages has been shown to subject animals to
less stress and lead to improved survivability in exotic animal spe-
cies; in other words, a tentatively diagnosed live patient is preferable
to a confirmed diagnosis at necropsy (Doneley, 2005).
Acquiring a clinical history of the rodent patient is not unlike
that done for any patient in clinical veterinary practice (see details
in Chapter 1), and template medical record sheets and sick animal
reports are provided (Figures 1.1 and 1.2) for mice noted to be in less-
than-optimal health condition.
critical care management for laboratory mice 27
Table 2.1: Miscellaneous Parameters for the Laboratory Mouse

Parameter Value
Lifespan 2–3 years
Age of sexual maturity 6 weeks
Gestation 19–21 days
Adult body weight 28–40 ga
Blood volume 76–80 ml/kg = 2.3–2.4 ml total for a 30-g mouse
Food intake 12–18 g/100 g BW/day
Water intake 15 ml/100 g BW/day
Packed cell volume (PCV) 38.5–45.1%
Glucose 106–278 mg/dlb
Body temperature (rectal) 36.5–38.0°C (97.5–100.4°F)
Respiratory rate 80–230 breaths/min
Heart rate 500–700+ beats/min
Source: Adapted from Banks, RE, Sharp, JM, Doss, SD, and Vanderford, DA. 2010.
Mice, pp. 73–80. In Exotic Small Mammal Care and Husbandry. Wiley-Blackwell,
Ames, IA; Danneman, PJ, Suckow, MA, and Brayton, CF. 2012. The Laboratory
Mouse, 2nd edition. CRC Press, Boca Raton, FL; and Suckow, MA, Danneman,
P, and Brayton, C. 2001. The Laboratory Mouse. CRC Press, Boca Raton, FL.
a Weights will vary depending on diet, age, stock or strain, gender.
b Values are dependent on collection method and may be influenced by anesthesia.
Typical values for biologic parameters in mice are presented in

Table 2.1. The size of the typical adult laboratory mouse ranges from
about 28 to 40 g, with less distinction by weight between gender than
other species; this small size makes it difficult to precisely quantify
body temperature, heart rate, and respiration rate without the use of
telemetric implants or other specialized equipment.
Physical Examination
Familiarity with the appearance of a routine clinically healthy
mouse is key to ensure recognition of one that develops abnormal
clinical signs. Visual examination of the animal is the most criti-
cal step in assessing the overall physical condition of the labora-
tory mouse. Observation of the animal in its home cage environment
is critical prior to performing a physical examination; this permits
overt lesions, behavioral abnormalities, and general activity to be
assessed rapidly. Animals in poor health will likely benefit from
placement in warming incubators or containers where warmth and
oxygen (flow rate 1–2 L/min) may be administered automatically
(Klaphake, 2006).
Prior to manual restraint and handling of laboratory mice, disposable
nitrile/latex gloves should be donned. Gentle single-handed restraint
Fig. 2.1 Retrieval of mice from a cage can be conducted using a

grip at the tail base to lift the animal or gently cupping the whole
a nimal (not shown) and placing it on the cage lid. The animal can
then be held by the scruff and turned on its back to better perform
palpation of the abdomen for abnormalities or held upright for

injections or gavage. (Images courtesy of University of Michigan,
ULAM.)
of the mouse (Figure 2.1) will allow for the ability to closely observe
skin and hair coat conditions, any ocular discharge or abnormalities,
tooth overgrowth, abnormal masses, or unusual presentations in the
anogenital region. Keep in mind that animals in critical c
ondition may
need to be sedated to perform these assessments and mitigate stress
levels. Gentle palpation of the abdomen, using a pincer technique
with the thumb and forefinger, should help to confirm pregnancy in
females that may present with dystocia and to identify abnormalities
like growths, enlarged lymph nodes (lymphadenopathy), or bladder
distention. A nonpregnant abdomen is generally “soft and doughy”
to the touch, and one has difficulty defining structures as partic-
ular organs; for example, something firm in the distal colon that
is not consistent with fecal pellets may require further diagnostics
(Klaphake, 2006).
Physiological aspects, like body weight, activity, and b ehavior

ssessments, are useful to measure and monitor serially. Hair coat
a
quality should be reviewed regarding location areas of a lopecia
(baldness), open or closed wounds, or poor grooming. In a ddition,
respiratory status (difficult or labored breathing with a more frequent/
diminished rate than expected) should be evaluated. Relative
perfusion status, ascertained by the color of mucous membranes,
reflects the transport of fluid and oxygen in blood to meet meta-
bolic needs. For a mouse, perfusion can be most easily ascertained
by rectal mucosal inspection; as well, the color of nonpigmented
ears and tails may assist with this assessment. Collectively, these
physiologic measures provide a crude interpretation of the “ABCs”
(airway/breathing/circulation) of critical care medicine. Finally, the
particular experimental use of the mouse, as described and approved
in an Institutional Animal Care and Use Committee proposal, must
be considered, and any adverse effects of the experimental proce-
dures should be documented.
Body Condition Scoring

Assessing general body condition, as a means for assessing health
status in any animal species, is an excellent tool to apply toward
mice. The use of a BCS (body condition score) scale (generally on a
range from 1 [wasted; emaciated] to 5 [obese]) is greatly enhanced by
the adherence to definitions of numerical health diagrams that rep-
resent each score on the scale (Ullman-Cullere and Foltz, 1999). This
tool is exceptionally valuable for any laboratory animal group with
variable experience in working with mice, as it provides a uniform
health assessment tool (Figure 2.2).
Overall percentages of weight loss should be monitored, yet may or
may not be pertinent, depending on the disease model and whether
the animals are expected to or may spontaneously develop tumors
(Paster et al., 2009) or ascites. Typically, weight loss of more than
20–25% from preexperimental baseline may warrant critical care
measures and potentially euthanasia, depending on institutional
policies.
Clinical Assessments of Ill Health and Pain in Mice

Mice are prey species; as such, they are conditioned to suppress
overt painful and ill behaviors, particularly when being handled by
personnel. Clinical assessments of ill health and pain in mice have
BC 1
Mouse is emaciated.
• Skeletal structure extremely prominent;
little or no flesh cover.
• Vertebrae distinctly segmented.
BC 2
Mouse is underconditioned.
• Segmentation of vertebral column evident.
• Dorsal pelvic bones are readily palpable.
BC 3
Mouse is well-conditioned.
• Vertebrae and dorsal pelvis not prominent;
palpable with slight pressure.
BC 4
Mouse is overconditioned.
• Spine is continuous column.
• Vertebrae palpable only with firm pressure.
BC 5
Mouse is obese.
• Mouse is smooth and bulky.
• Bone structure disappears under flesh and
subcutaneous fat.
A “+” or a “–” can be added to the body condition score
if additional increments are necessary (i.e. ...2+, 2, 2....)
Fig. 2.2 Schematic for scoring mouse body condition. (Reprinted

with permission from AALAS. Ullman-Cullere, MH, and Foltz, CJ.
1999. Lab Anim Sci 49:319–323.)
been described (Kohn et al., 2007, Miller and Richardson, 2011) and
include the following:
• Vocalization, particularly when handled or a painful area is

palpated
• Reduced grooming or piloerection, leading to a “ruffled fur”
appearance
• Reduced level of spontaneous and exploratory (sniffing,
rearing) activity to the point that mice may not be moving: a
“moribund condition”
• Hunched posture, with potential “guarding” of abdomen and
reduced mobility
• Squint-eyed appearance (either unilateral or bilateral)

• Increased aggressiveness on handling; may bite without
warning
• Distanced from cage mates
• Reduced BCS, likely secondary to reduced nutritional intake
or experimental model resulting in muscle wasting and
weight loss
• Self-mutilation (excessive licking, biting, scratching) of the
painful area
• Discharge originating from the eyes, nose, ears, or perineal
region
• Abnormal postures, ataxia, circling, and raised tail position
• Palpation of unexpected masses
Monitoring Frequency
A detailed and descriptive plan for scheduled monitoring of research
animals both before and after an experimental procedure, including
the provision of therapeutic treatments and supportive care, should
be included in the IACUC protocol submission. Investigators should
be aware that, as the potential for pain/distress in animals rises,
there should be an increasing intensity of monitoring and frequency
of observations performed.
Objective Scoring Systems

Professional and clinical judgments are inherent to the evalua-
tion of an animal’s well-being and are essential to the ultimate
decision for administration of treatments or planned removal from
studies for euthanasia, based on welfare rationale. To standardize
approaches for care and treatment, objective data-based approaches
to predicting demise and imminent death, when developed for spe-
cific experimental models, should facilitate the implementation
of timely euthanasia before the onset of clinically overt signs of
moribund state (Toth and Gardiner, 2000). Individualized scoring
systems are a common means by which humane interventions/
endpoints can be defined and implemented. Examples of template
scoring systems (Figure 2.3; Table 2.2), by which the health of lab-
oratory rodents can be judged, are provided (Adamson et al., 2010,
Scoring Characteristics
Active curiosity • Moving quickly around cage

(score = 1) • Frequent standing at sides of cage and close to
cage mates
• Active investigation of surroundings
Mildly decreased activity • Reduced movement around cage

(score = 2) • Little to no investigation of surroundings
• Seeks shelter/prefers corners of cage
• Will move around cage when stimulated by gentle
handling
Severely decreased activity • No movement around cage
(score = 3) • May be moribund or only move slightly when
stimulated by gentle handling
• Typically isolated from cage mates
Fig. 2.3 Template scoring system for overall rodent activity in

the home cage; a score of 3 over the course of 24 h would indicate
potential removal from a study and likely euthanasia.
Hankenson et al., 2013). To eliminate the inherent subjectivity

in clinical assessments, it is advised that a consensus on over-
all health and activity scoring be established through hands-on
t raining of observers by the principal investigator or veterinary

staff.
A novel method of potential pain assessment in laboratory mice
is through coding of facial expressions of pain, also referred to
as the Mouse Grimace Scale (MGS) (Langford et al., 2010). The
technique was adapted from the use of facial expressions in
human infants; in mice, orbital tightening, nose bulges, cheek
bulges, and changes in ear and whisker position are the five
“action units” scored on a 0–2 scale for their prominence in still
photographs taken from digital video of mice in either a baseline
or pain c ondition due to an irritant (Figure 2.4). The MGS displays
high accuracy and r eliability, can quantify pain of duration up to
about 1 day, and is sensitive to detecting weak analgesic effects.
Dedicated observation of cohorts of laboratory mice resulted in the
identification of these subtle distinctions, like the bulging altera-
tion from the t ypical smooth appearance of the nose and cheek in
control a nimals. Use of the grimace scale has been applied to mice
receiving analgesia to determine efficacy of pain relief (Matsumiya
et al., 2012).
Table 2.2: Visual A ssessment Scoring System that A lso Can Be Used

to Score A nimals Postprocedurally
Coordination Overall
Score Hair Coat Eyes and Posture Condition
0 Normal, well groomed, Open, Normal Normal
smooth, sleek alert
1 Not well groomed Squinted Walks Roughened
awkwardly or appearance but
slightly otherwise
hunched, activity and
otherwise behaviors within
active, mobile normal limits
2 Rough hair coat, Squinted Hunched, Slightly
unkempt surgical to closed abdominal depressed, poor
site stretching appearance,
observed, behaviors not
reduced activity within normal
limits due to
agitation
3 Very rough hair coat, Closed Hunched, Depressed,
hair loss, unkempt stumbles when increasingly poor
surgical site moving, appearance,
inactive abnormal
behavior
4 Hunched, not Unresponsive
moving
Source: Modified from Adamson, TW, Kendall, LV, Goss, S, Grayson, K, Touma,
C, Palme, R, Chen, JQ, and Borowsky, AD. 2010. J Am Assoc Lab Anim Sci
49:610–616.
Characteristics of the animal (hair coat, eyes, coordination, and overall condition) are
scored independently and then summed and averaged to obtain a final pain
index score.
veterinary care measures

Administration of Fluids
Evidence of dehydration has been documented as an outcome of
potential pain and distress, related to underlying clinical conditions,
experimental interventions, or husbandry parameters. Dehydration
may be assessed by performing a skin tent or gentle pinch of scruff
over the scapulae of the mouse and assessing the time that passes
for the skin to return to normal placement. A prolonged return
time indicates a degree of dehydration that should be ameliorated.
Any administered fluids should be from a sterile source to avoid
introduction of infectious disease agents. In the critical rodent patient,
Not Present Moderate Obvious

“0” “1” “2”
Fig. 2.4 Representative photographs of the Mouse Grimace Scale

for a mouse at baseline (facial grimacing not present, 0); a mouse
with moderate facial grimacing (1); and a mouse with obvious facial
grimacing (2). (Reprinted with permission from AALAS. Matsumiya,
LC, Sorge, RE, Sotocinal, SG, Tabaka, JM, Wieskopf, JS, Zaloum, A,
King, OD, and Mogil, JS. 2012. J Am Assoc Lab Anim Sci 51:42–49.)
the subcutaneous (SC) route of administration may be most readily

accessed; a balanced electrolyte solution (potentially with 2.5% dex-
trose) at an initial rate of 40–50 mg/kg twice daily is recommended
and can be administered through a butterfly needle and attached line
to allow less restraint of the animal (Klaphake, 2006). Prophylactic
sterile fluid administration, 0.9% NaCl or polystarch given subcuta-
neously at a dose of 1.0 ml, can significantly improve survival rates
in mouse models of cancer receiving carcinogen treatments (Smith
et al., 1999). Additional fluid support is beneficial in terms of raising
arterial blood pressure, although there may be a negative impact on
changes in organ water content and increased potential for anemia
(Zuurbier et al., 2002).
Water and fluid replacement sources are gaining in p opularity,
expanding from products initially developed as sustainable fluid
sources for the duration of rodent shipping and transport. The
provision of these water replacements, in disposable single-use

containers, is typically done on the cage floor for rapid access by
those animals in ill health. These supplementary fluid sources, when
combined with food, can maintain the health of rodents for at least
7 days in the absence of routine water sources (Luo et al., 2003).
Additional critical care considerations for nutritional support, fluid
administration, and available products are provided in Chapter 4.
Blood Sampling
Blood collection, or venipuncture, is a common procedure
performed in animal research for experimental, routine, or c ritical
care r easons (Danneman et al., 2012, Hoff, 2000, Hrapkiewicz and
Medina, 2007, Suckow et al., 2001). Sampling allows for testing
of serum chemistry parameters, as well as complete blood counts
(CBCs). Sampling sites in mice (Table 2.3) include the retro-orbital
sinus (typically with animals under anesthesia), facial vein (with
animals conscious), medial and lateral saphenous veins, and
tail vessels (Horne et al., 2003). Retro-orbital bleeds are readily
performed using m icrohematocrit c apillary tubes formulated for col-
lection of microvolumes of blood with a ppropriate anticoagulants.
It has been shown that a drop of 0.05% ophthalmic proparacaine
hydrochloride solution, directly onto the eye to be bled significantly
reduces the incidence of responsiveness to retro-orbital blood col-
lection (Taylor et al., 2000).
Mandibular bleeds (also referred to as using the facial vein, super-
ficial temporal vein, submandibular, or cheek bleed) can success-
fully be performed with puncture of the vascular bed using a 20- to
22-gauge needle or small disposable lancet. These are recommended
in mice weighing more than 20 g (Figure 2.5).
Table 2.3: Recommended Sampling Sites and Related Information for

Blood Collection in Mice
Approximate Range
Anatomical Site Anesthesia? of Volume Collected Comments
Lateral tail vein Not required Up to 1% of BW
Lateral saphenous Not required Up to 1% of BW
vein
Tail clip Not required Up to 1% of BW ~1 mm of distal end of
tail should be clipped
Retro-orbital Required Up to 1% of BW Should alternate eyes
vasculature for repeated sampling;
topical anesthetic
drops prior to bleeds;
ophthalmic ointment
should be applied
following bleed
Submandibular Not required Up to 1% of BW Limited to adult mice;
vessels single sampling
recommended
Cardiac Required 500+ µl Terminal procedure
only
Source: Modified from University of Pennsylvania, ULAR.
(A) (B)
Fig. 2.5 The mandibular bleeding method is used to collect samples

typically from unanesthetized mice. Mice are manually restrained by
a one-handed grip, and a disposable lancet or needle tip is then used
to prick the facial vessels (A). Drops of blood can be collected directly
into a microhematocrit tube for processing (B).
Anatomic differences in the skull between mouse strains may

make it difficult to consistently find the vessels for access. Care must
be taken with any method to ensure that structures (globe of the eye,
ear canal, muscles, nerves, and bones) surrounding the sampling site
are not injured; this may be better accomplished by inserting the flat
edge of the lancet parallel to the masticatory muscle (Forbes et al.,
2010). Typically, bleeds from the facial vein are recommended only
for one-time, nonserial collections (Coman et al., 2010, Tomlinson
et al., 2004). Nude mice have been documented as more susceptible
to hematoma formation and tissue damage following sampling from
the submandibular location (Nugent Britt et al., 2011). It is impera-
tive that manual pressure be applied to achieve hemostasis following
blood collection. Controversies surrounding facial sampling revolve
around whether anesthesia should be used for this method. As well,
facial sampling has the potential for adverse outcomes, including
rapid clotting of blood, which prevents accurate serology results; for-
mation of hematomas; blood exiting through a pierced ear canal;
punctures into the oral cavity; inconsistency of volume acquisition;
and high potential for pain and distress, leading to stupor and ataxia
if conducted improperly. As well, there is a rare chance for significant
and life-threatening hemorrhage to occur (Forbes et al., 2010).
For critical animals, it may be useful to access the saphenous vein
on the hind limb or employ the tail-clip method (Figure 2.6), by which
the very end of the tail is removed to allow for a drop of blood to be
Fig. 2.6 Modified tail-clip technique. Demonstration of the minimal

restraint used during the modified tail-clip blood collection procedure
and the ability to pipette blood directly from the tail. (Reprinted with
permission from AALAS. Abatan, OI, Welch, KB, and Nemzek, JA.
2008. J Am Assoc Lab Anim Sci 47:8–15.)
collected (Abatan et al., 2008). For further description, in the modi-

fied tail-clip technique, the mouse is placed on a wired surface to
allow for gripping by toenails.
During the tail-clip collection procedure, the animal handler
moves with the animal and only restricts animal movement if the
mouse attempts to escape from the working surface. Only the d istal
1 to 2 mm of the tail need to be clipped, and a capillary pipette (flushed
with an anticoagulant like EDTA [ethylenediaminetetraacetic acid]
or heparin) is then used to collect 20-μl samples from the exposed
tail tip. Immediately after collection, styptic powder (Kwik-stop®) can
be applied to the tail tip for hemostasis. For repeated sampling, the
surface of the clip site can be disrupted without the need to remove
additional tail tissue (Abatan et al., 2008).
Total blood volume (TBV) in a mouse has been defined as about
6–7% of BW (Hrapkiewicz and Medina, 2007, Raabe et al., 2011).
Because blood regeneration rates may vary among mouse strains, a
safe guideline for maximum volume of blood to remove from other-
wise-healthy animals is 1% of BW (Klaphake, 2006, Paul-Murphy,
1996). For example, 0.30 ml (or 300 µl) can be withdrawn at a single
sampling from a 30-g mouse; importantly, this volume may be more
than needed and should be diminished to the minimum required
for obtaining test results in the critical patient. Following sampling
of 1% BW volume, replacement fluid therapy (0.5–1.0 ml SC or IP
[intraperitoneally] of sterile isotonic fluid) should be provided.
If blood is to be collected serially (e.g., over a period of days), then

sampling volumes should not exceed about 20% of BW within a
1-week period to avoid weight loss and anemia. Anemia generally is
defined as a red blood cell (RBC) count, hemoglobin concentration, or
hematocrit (HCT) value lower than 2 standard deviations (SDs) below
the mean of a normal population (low end of normal range for HCT
is ~35%); hemoglobin has been highlighted as the most direct and
sensitive measure for detecting anemia at 2 SD below the mean of
baseline hemoglobin values (Raabe et al., 2011). Permissible frequen-
cies have been further delineated for 10- to 14-week-old clinically
healthy C57BL/6 mice for collection of weekly blood samples for up
to 6 weeks: about 15% of TBV weekly from males is acceptable, and
up to about 25% of TBV weekly from females is acceptable, without
adverse effects (Raabe et al., 2011).
When dealing with a sick mouse, it will be critical to decide the
most informative or essential tests to run with limited volumes of
blood. Blood smears can be made with fresh whole blood and the
remainder of the sample collected in anticoagulant for select serum
chemistry and further testing (Wiedmeyer et al., 2007).
Body Temperature Monitoring

Body temperature monitoring in mice, while a useful aspect of the
diagnostic profile, is most often logistically challenging to perform.
Often, simple handling of sick mice will provide some indication of
whether they are excessively cool or warm to the touch. Mechanisms
of obtaining temperatures in rodents can include contact methods
(rectal, surface probes) and indirect methods (telemetry devices,
implanted microchips, infrared laser).
Rectal probes require gentle placement and positioning d uring
procedures in sedated mice, but are efficacious for monitoring
temperature (McCann and Mitchel, 1994). Telemeterized animals,

or those with an implantable device typically surgically inserted
into the peritoneal cavity, are most easily monitored for variations
outside the normal body temperature interval of about 95°F to

100.5°F; however, microchip transponders (Bio Medic Data Systems,
Seaford, DE), surface temperature probes, and infrared noncontact
thermometers may also be suitable for taking temperatures in cer-
tain mouse strains and for assessment of disease model progression
(Byrum et al., 2011, Hankenson et al., 2013, Miller and Haimovich,
B., 2011, Newsom et al., 2004). Placement of animals on circulating
water b
lankets (Figure 2.7) and microwaveable padding (Figure 2.8)
Fig. 2.7 Anesthetized mouse on warm-water recirculating blanket

(T/Pump® Classic TP650, Gaymar ®) placed on manufacturer’s setting
of “medium” heat. Animal’s eyes were treated with topical ophthal-
mic tears to maintain moisture. (Image courtesy of the University of
Fig. 2.8 Reusable heating pad (SnuggleSafe® Microwave Heatpad,

West Sussex, UK) used for thermal support of anesthetized mice with-
out a cover (left) and with manufacturer’s cover (right). (Reprinted
with permission from AALAS. Taylor, DK. 2007. J Am Assoc Lab Anim
Sci 46:37–41.)
has been successful for maintaining body temperatures with rodents

under anesthesia (Caro et al., 2012, Charles et al., 2005, Taylor, 2007).
Body temperatures may vary depending on the biomedical model.
For example, in mouse models of septic shock (Nemzek et al., 2004)
and infectious herpesviral disease (Hankenson et al., 2013), non-
transient drops in body temperatures are a criterion for determining
humane endpoints prior to spontaneous death.
Endotracheal Intubation
Mouse endotracheal (ET) intubation (Figure 2.9) has been refined to
enhance the ability to mechanically ventilate and provide inhalant
anesthesia to this species (Hamacher et al., 2008, Rivera et al., 2005,
Spoelstra et al., 2007, Tonsfeldt et al., 2007).
Direct visualization of the arytenoid cartilages can be p erformed
with a handheld light source combined with an otoscope. Mice
anesthetized with inhalant anesthesia (e.g., isoflurane) can be
placed supine on a tilt board or an incisor bar (Burns et al., 2005)
or on any sanitizable surface (plexiglass) that puts the animal at
an incline (Rivera et al., 2005). Mice are restrained with a rubber
band placed beneath the incisors and around the support board.
Transillumination is accomplished using a fiber-optic light beam or
by aiming a horizontal microscope light at the midtracheal level. The
patient’s tongue can be held aside with a pair of forceps shielded
with polyethylene (PE) tubing or held flat against the lower jaw with
the bent end of a small weighing spatula. The backlit larynx is then
visualized, and a 20-gauge, 1.25-inch Teflon intravenous catheter, or
species-specific ET tube, is inserted into the trachea. Verification of
accurate placement is noted by presence of condensation on the tube
A B C
Fig. 2.9 Intubation of the mouse with (A) retraction of the tongue

and insertion of the intubation system (arrow points to light at the
end of the fiber), (B) verification of tube placement with a modified
disposable plastic transfer pipette (arrow points to the male Luer
connector), and (C) securing the endotracheal tube to the muzzle
of the mouse. (Reprinted with permission from AALAS. Rivera, B,
Miller, S, Brown, E, and Price, R. 2005. Contemp Top Lab Anim Sci
44:52–55.)
and gentle inflation of the lungs with a small disposable pipette. Once
intubated, mice can rapidly be moved from the inclined support board
and positioned so that the ET tube is attached to the inhalant anes-
thesia flow for the duration of the procedure of interest. Note that in
the critically ill mouse, intubation may be extremely challenging and
should be attempted only as a last resort to gain airway access if tra-
cheostomy cannot be performed (see relevant section in Chapter 4).
Injections and Oral Administration

Injections can be performed routinely using multiple routes (Table 2.4)
for the mouse, including subcutaneous, intradermal (ID), intraperi-
toneal, intratracheal (IT), and intravenous (IV) routes, as described
in Chapter 1. As well, retro-orbital injections in mice have been
described as an alternative to tail-vein injections, to deliver volumes
up to 150 µl in adult animals and up to 10 µl in neonates (Yardeni
et al., 2011).
Administration of drugs and fluids can also be done by oral
gavage and voluntary ingestion. Creative approaches to disguis-
ing medications in palatable substances have been successful,
like p rovision of analgesics in Nutella® chocolate spread (Goldkuhl
et al., 2008, Jacobsen et al., 2011, Kalliokoski et al., 2011, Strom
et al., 2012), although mice may need to acclimate to the novel sub-
stance prior to drug incorporation. Honey (in 100-µl doses adminis-
tered by syringe) has been used to deliver antiparasitic medications
Table 2.4: Recommendations for Injection Dose Limits Based on

Weight of Laboratory Mice
Injection Limits (ml/kg)
Route PO SC IP IMa IV (Bolusb) IV (Slow)
Dose (ml/kg) 10 10 20 0.05 5 25
Weight (kg)
0.020 0.20 mL 0.20 mL 0.40 mL 0.001 mL 0.10 mL 0.50 mL
0.025 0.25 0.25 0.50 0.001 0.13 0.63
0.030 0.30 0.30 0.60 0.001 0.15 0.75
0.035 0.35 0.35 0.70 0.002 0.18 0.88
0.040 0.40 0.40 0.80 0.002 0.20 1.00
0.045 0.45 0.45 0.90 0.002 0.23 1.13
0.050 0.50 0.50 1.00 0.002 0.25 1.25
a Technique is discouraged in mice due to small muscle mass.
b A bolus is a larger dose given over a shorter period of time.

daily for up to several weeks (Kuster et al., 2012). Therapeutics

delivered in peanut butter pills have been described; in brief, pea-
nut butter is heated to 37°C, and drug doses are then mixed in
for approximately 15 min. Pilling molds are used to isolate single-
dose a pplications, which can be frozen at -80°C until needed. Mice
readily consume the pellets, ensuring delivery of the complete dose
(Cope et al., 2005).
Urine Sampling
Urinalyses in mice are challenging due to volume limitations;
however, the propensity for mice to urinate on handling assists in
the collection of free-catch samples (Chew and Chua, 2003). Urine
droplets can be collected into plastic well plates and then aliquoted
by pipette for appropriate assessment and assays (Table 2.5) (Kurien
et al., 2004, Kurien and Scofield, 1999). Facilitating urination in the
mouse can be done by applying equal pressure in a gentle massaging
manner at both sides of the lower back near the tail, with the thumb
on one side and the fore and middle fingers on the other side, rubbing
up and down; this application of pressure to the caudal back area
of the mouse facilitates expression of a maximum volume (>50 µl) of
urine for collection (Chew and Chua, 2003). Alternatively, gentle, firm
pressure on the bladder with the thumb and index finger can stimu-
late urination, which can be collected on clean plastic wrap, pipetted
into sterile microfuge tubes, and stored immediately at −20°C until
use (Maier et al., 2007).
Critically ill mice should be stabilized prior to attempting uri-
nary catheterization if urine collection by other methods has been
unsuccessful. Urinary catheterization should only be performed on
Table 2.5: Parameters for Urine in the

Laboratory Mouse
Parameter Value
Color Clear or slightly yellow
Volume 0.5–2.4 mg/24 h
Specific gravity 1.030
pH 5.0
Glucose 0.5–3.0 mg/24 h
Protein 0.6–2.6 mg/24 h
Source: Adapted from Danneman, PJ, Suckow, MA,
and Brayton, CF. 2012. The Laboratory Mouse,
2nd edition. CRC Press, Boca Raton, FL.
anesthetized mice. Aseptic technique (see Chapter 4, “Perioperative

Care Considerations”) and atraumatic approach should be used
during placement of a urinary catheter. Prior to insertion of the
catheter, the external urinary orifice should be gently cleansed
using a disinfecting (e.g., chlorhexidine) solution. The individual
performing the catheterization is advised to don sterile surgical
gloves, use a sterile catheter, and apply a small amount of sterile
water-soluble lubricant on the external urinary orifice. Additional
sterile lubricant should be applied in a thin layer to cover the sur-
face of the urinary catheter for ease of insertion into the urinary
orifice (St. Claire et al., 1999). The diameter of the urinary catheter
should be the minimum that can be inserted into the bladder and
still prevent urinary leakage around the catheter. The distance
from the external urinary orifice to the neck of the bladder should
be estimated prior to catheter insertion.
The anatomy of the female mouse is unique in that the uri-
nary orifice is external and just anterior to the vaginal opening.
Catheters for adult female mice can be made by using number 10
(1.8-French) Intramedic ® PE tubing. A guidewire can be threaded
through the PE tubing to increase the rigidity of the catheter.
Care should be taken that the tip of the guidewire does not extend
past the end of the catheter. Guidewires can be made of stainless
steel surgical wire (Ethicon, Somerville, NJ) and are coated with
a water-soluble lubricant to ease placement and removal from the
PE tubing. The approximate distance from the external urinary
orifice to the neck of the bladder for a 20-g female mouse is 10 mm
(St. Claire et al., 1999).
Once urine is collected, standard veterinary refractometers require
approximately 60 µl of mouse urine to generate a reliable reading
of the urine specific gravity value (Forbes-McBean and Brayton,
2012). Elevated plasma creatinine levels have routinely been used
as a marker of reduced kidney function in animal studies; however,
historically these have been difficult to measure for mice. Because
blood urea nitrogen (BUN) concentration increases as kidney func-
tion declines, plasma BUN is a decent alternative to creatinine as a
high-throughput screen for evaluating kidney function in mice. In
addition to BUN as a marker for kidney function, the ratio of urinary
albumin concentration to creatinine concentration is commonly used
as an indicator of kidney damage in animal studies. It has been dem-
onstrated that chromagens in mouse plasma do not interfere with
autoanalyzer methodologies for quantifying BUN c oncentrations
(Grindle et al., 2006).
abnormal, critical, and emergent conditions

Categories of laboratory rodent health concerns are discussed in
alphabetical order to facilitate locating topical information. Under
each topic, the “cause and impact” has been provided, and “poten-
tial treatments” offer suggestions about procedures, therapeutic
treatments, or husbandry and environmental alterations. Every
attempt has been made to provide citations from the literature for
evidence-based medical outcomes. For those health concerns that
list drug therapy options, please refer to the rodent formulary pro-
vided in Appendix C for additional details on dosages and route of
delivery.
Abdominal Swelling
• Cause and impact: Animals may present acutely with an
enlarged or swollen abdominal (peritoneal) cavity. This may
be caused by ascites fluid accumulation (see further section
if presumed research related), organomegaly, pregnancy (in
females), hemoabdomen, enlarged bladder, subcutaneous
edema, or neoplasia, among other differentials (Figure 2.10).
The impact on the mouse can be severe due to pressure placed
on the thoracic cavity, secondary respiratory difficulty due to
restricted ability of the lungs to expand, and anemia due to
blood loss into the abdomen.
A B C
Fig. 2.10 Abdominal swelling reported in a mouse (A). The animal

was subsequently found to have subcutaneous edema (B) and 12 ml
of serosanguinous fluid free in the abdomen (C). (Images courtesy of
the University of Pennsylvania, ULAR.)
• Potential treatments: Determining the differentials will be

largely based on physical examination and palpation to iden-
tify masses within the abdominal cavity; however, it may be
that abdominal contents cannot be appreciated due to the
volume of dilation. Fine-needle aspirates conducted over the
caudal ventral midline can determine if the extracted fluid is
urine. Aspiration off the midline to the right, using a ventral
approach, may assist with removal of free abdominal fluid,
which can be evaluated for cellular and proteinaceous com-
ponents (transudate, modified transudate, or exudate) by
microscopic review of a fluid drop placed on a glass slide.
Experimental information and assessment of gender of cage
mates will assist with ruling out the potential for a pregnant
mouse; as well, nipples may be prominent in pregnant mice.
If the abdomen appears swollen due to a large neoplastic
growth, and depending on the importance of the mouse to
the research outcomes and colony, exploratory surgery may
be performed to extract any growths from the peritoneal cav-
ity. These mice will require close monitoring postoperatively,
particularly due to the potential for heightened preoperative
stress and weakened physiologic condition.
Abscessation
• Cause and impact: Coagulase-positive Staphylococcus
aureus is commonly found on the skin of animals and has
been reported as a primary cause of facial abscesses in
mice (Figures 2.11 and 2.12) (Lawson, 2010). An oral route
of infection has been suggested, and it has been verified that
introduction of bacteria occurs through piercing of the oral
mucosa by pelage or vibrissae (hairs) following grooming or
barbering activities. Hair then can become entrapped in the
periodontal spaces as a side effect of these activities. Severe
localized periodontal bone loss in the oral cavity, secondary
to hair ingestion and abscessation, has been confirmed by
micro-computed tomography (micro-CT).
Ultimately, abscess formation can occur anywhere on the
body at a site where the skin integrity has been altered and
bacterial contamination introduced (Figure 2.13).
• Potential treatments: Abscesses in mice, depending on loca-
tion and size, can be treated similarly to those in other species.
A B
Fig. 2.11 Mouse that presented with a facial mass (A), determined

to be an abscess. (Image courtesy of the University of Pennsylvania,
ULAR.) (B) Facial abscess (ventral view) with associated draining
tract. (Reprinted with permission from AALAS. Lawson, GW. 2010.
Comp Med 60:200–204.)
A B C
Fig. 2.12 Adult female Swiss Webster mouse housed in a conventional

facility was reported with a submandibular swelling (A). Physical
examination concluded that the animal was in thin body condition,
hunched, with a draining tract noted on left ventral aspect of m
andible
(B). Degree of abscessation precluded ability to flush and drain the
area (C), and the animal was humanely euthanized. Culture results
confirmed infection with S. aureus. (Images courtesy of University of
Fine-needle aspiration can be performed to d etermine if the

extracted material is purulent; culture and antibiotic sensi-
tivity testing can be performed to best determine a ntibiotic
treatment. Under anesthesia and using aseptic conditions,
abscesses can be lanced, drained, flushed, and most often
left to heal by second intention. Topical antibiotic ointment
(Neosporin®; polymyxin B sulfate + neomycin sulfate + bac-
itracin zinc) is most commonly utilized at the site daily or
every other day (EOD) for 3 to 5 days; systemic antibiotic
Fig. 2.13 Abscess draining tract in an adult female FRG (with

humanized mouse liver) mouse that presented with difficulty walk-
ing. This strain is extremely immunodeficient, and the entire colony
was maintained on a ntibiotic (enrofloxacin-treated water) to amelio-
rate further secondary bacterial infections. All nutritional, fluid, and
caging materials were a utoclaved into the room to further diminish
the possibility of secondary infectious disease. (Image courtesy of
University of Pennsylvania, ULAR.)
administration may be selected for additional coverage d

uring
the healing process. Analgesics should be applied to manage
pain secondary to the eradication of the abscess, dependent
on the level of tissue manipulation.
Cage Flooding with Subsequent Hypothermia

• Cause and impact: Water bottle or automatic watering
system malfunctions that result in cage flooding can be a

source of significant morbidity and mortality in mice used
in biomedical research. Following an investigation into the
cause for leaking water valves, the findings identified a com-
bination of bedding materials and rodent fur lodged in failed
water valves (Ogeka, 2009). Even relatively minor floods from
watering sources into the cage can lead to significant hypo-
thermia and potentially death if left uncorrected.
• Potential treatments: Prompt warming is an important
first aid procedure for wet mice, with the target temperature
for rewarming equivalent to 90–100°F. Since these mice are
typically conscious, despite hypothermia, external warmers
are advised. These can include heat lamps (250 W), micro-
waveable gel packs (wrapped and placed within the cage
in a location that prevents direct contact with animals),

recirculating warm-water blankets, and reusable chemically
activated heating pads (placed under the cage). Rewarming
first aid stations can be assembled and remain permanently
in housing areas to facilitate prompt treatment of any ani-
mals affected by cage flooding (Figure 2.14).
Heat lamps can be beneficial for rewarming purposes but
can lead to overly hot temperatures (120°F) in less than 15
min when placed at a distance of 4 inches from cage lids.
Therefore, placement of hypothermic mice, in a dry cage with
bedding, at a distance of 6 inches from a heat lamp for 35 min
will achieve a warming temperature of 100°F, which will then
remain elevated for 60 min (Hedrick et al., 2009).
A standard rewarming practice of the University of
Colorado-Denver is to scruff a damp mouse and manually
submerge the animal in warm water. Water should be warm
to the touch but not excessively hot. The animal is kept in
the warm bath for 1-2 minutes. These mice are then dried
throughly and placed in a dry and clean cage set-up.
Microwaveable pads heated to 90°F can be placed within
the dry cage with mice for 10–20 min and will keep the cage
A B
Fig. 2.14 An example of how a flooded mouse cage appears following a

malfunction of the watering system (A); this r ewarming first aid s
tation
(B) was developed for damp or hypothermic rodents recovering from
a cage flood event, crafted from a commercial histology slide warmer
(front) with a customized polycarbonate extender (rear) to support the
full length of rodent cage and allow for warm and cool thermal zones.
Note that the station can also be used for postoperative recovery pur-
poses. (Images courtesy of Emory University; M. J. Huerkamp.)
warmed for 1 h; however, there is additional preparatory time

associated with microwaving for this device (including having
ready access to the microwave and ensuring the pad has not
been overheated or contains “hot spots” from uneven heat-
ing). Reusable chemical pads can be activated manually, and
they heat within 30 s. These chemical pads can be placed
within a new and dry cage of mice for 20 min and will keep
the cage warmed to 90–97°F for greater than 1 h (Shomer and
Berenblit, 2008).
Cages exposed to heat sources should never be left unat-
tended for prolonged periods as internal cage t emperatures can
reach limits detrimental to the animals (Hedrick et al., 2009).
Cannibalization
• Cause and impact: In rodent breeding colonies, there are
often legitimate concerns about the potential for mutilation
and cannibalization of neonates, resulting in the loss of
valuable research animals. The causes for cannibalism are
thought to be linked to stressors on the mother, including
handling or disrupting neonates too soon after delivery or
environmental influences, like construction noise and vibra-
tions. Cannibalism may also be strain related in more aggres-
sive mouse strains or may be conducted by male mice (which
have fathered the offspring).
• Potential treatments: Husbandry practices could involve a
decreased change cycle of cages of breeding mice with new
litters, such that after parturition these cages should be left
undisturbed (i.e., not changed to clean bedding) for at least
2 days postpartum. Breeding rooms can have altered light–
dark cycles for maximizing production, caging m aterials may
be tinted or colored, and enrichment materials (e.g., paper
enrichment shacks and plastic tubing) can be placed in the
environment to provide a degree of shielding for the dam.
Providing nesting material is essential, and females will
deliver pups into nests, which typically provide a softer and
warmer surface (than corncob bedding) for altricial neonates
during nursing and development.
Modification of poor maternal behaviors, particularly in
strains known to readily cannibalize, can be attempted.
Maternal administration of perphenazine on the day before or
morning of parturition to dams has been reported to decrease

the incidence of cannibalism in colonies of interferon-γ and
interleukin (IL) 4, IL-10, and IL-12 knockout mice of the DBA/1
and C57BL/6 background strains. Perphenazine (2–4 mg/kg)
can be supplied in water bottles. Results have shown that
medicated dams weaned 76.4% of their pups, compared with
untreated dams that weaned only 59.4% of their pups. Timing
of the administration of perphenazine did not appear to have
a significant impact on efficacy (Carter et al., 2002).
Nutritional supplementation with specialty treats to “dis-
tract” the dam from engaging in cannibalism have been
used with success and are described further in Chapter 4,
“Nutritional Therapy Considerations.”
Conjunctivitis
• Cause and impacts: The appearance of reddened, crusty, and
swollen conjunctiva in mice may be due to a number of causes
and should be treated as a painful condition that should be
monitored for improvement (Figure 2.15). Historically, in nude
mice (Bazille et al., 2001), conjunctivitis has been linked to
the contamination of the conjunctiva with cotton fibers from
nesting material that results in chronic irritation. In hairless
mice (SKH1) that have been reported with bilateral conjuncti-
vitis and blepharitis, the disorder is related to body and facial
hair shedding during the first weeks of the neonates’ lives
(Rosenbaum, 2010).
Fig. 2.15 Representative mice with presentation of severe conjuncti-

vitis. Spontaneous development of clinical signs (left); experimental
development secondary to ocular exposure to herpesviral infection
(right).
• Potential treatments: Husbandry practices should be

altered so that cages are changed with increased frequency
during the shedding of the neonates’ hairs. Known affected
strains may be moved from static housing to ventilated caging
as the incidence of conjunctivitis is increased in static hous-
ing conditions, likely due to the minimal airflow preventing
removal of the shed hairs from the facial and ocular area
(Rosenbaum, 2010).
Routine prophylactic cleaning of the conjunctiva with
sterile swabs and saline to remove fibers and hair is also

effective. Removal of material embedded in the eyelids can be
done with a swab primed with topical triple antibiotic (e.g.,
Neosporin) or a related antibiotic ophthalmic ointment (Swan
et al., 2010).
Cross Fostering of Neonates/Mouse Pups

• Cause and impact: In the course of breeding mice for
research studies, instances may arise when there is a loss
of a nursing mother, either at the time of delivery or prior to
weaning of the neonates or pups. The litter will not survive
unless there is immediate intervention by provision of a foster
mother or supplements provided by personnel.
For experimental and biosecurity reasons, intentional
cross fostering of pups can be used to eliminate potentially
harmful murine pathogens, including murine norovirus
(MNV), mouse hepatitis virus (MHV), and Helicobacter from
contaminated lines of mice.
• Potential treatments: Placement of orphaned pups, regard-
less of whether they are age matched or not age matched to
pups of the foster mother, has been successful for survival,
with pups up to 12 days of age. Pups to be fostered should
be gently rubbed with bedding (for transference of odors
from the foster mother) or intermixed with the litter of pups
belonging to the recipient dam to best facilitate orphaned pup
acceptance (Hickman and Swan, 2011).
To eradicate murine pathogens in newborn pups, litters
should be less than 24 h old and from cages in which bedding
material was changed within 24 h of planned cross fostering.
Note: Syphacia obvelata has not been eliminated successfully
through this cross-fostering technique (Artwohl et al., 2008).
Cross-fostered mice should be tested at a minimum of

4 to 12 weeks of age to ensure specific pathogen-free status
(Artwohl et al., 2008). In particular, for elimination of MNV,
it has been shown that cross fostering of neonatal mice from
MNV-infected to naïve dams is successful when pups are 1 to
3 days of age (Buxbaum et al., 2011, Compton, 2008). For elim-
ination of Helicobacter spp., mice should be fostered within
24 h of birth (Singletary et al., 2003, Truett et al., 2000).
Milk substitutes (replacers) for artificial rearing of pups
have been utilized for many years and have been based on rat
milk substitutes (RMSs; derived from cow’s milk) and canine
pup milk replacers (Auestad et al., 1989, Hoshiba, 2004).
More recently, milk substitutes have been formulated follow-
ing analysis of the components of actual milk collected from
ICR, BALB/c, and FVB/N mouse strains (Yajima et al., 2006).
Mouse formula includes protein (purified bovine casein and
whey), edible oils (up to five types, based on RMS), and vita-
mins similar to the RMS.
Administration of the milk replacer has been performed
using gastrostomy catheters (placed surgically; Figure 2.16)
and hand feeding with a surrogate nipple and nursing bottle
(Beierle et al., 2004, Hoshiba, 2004).
Dystocia
• Cause and impact: Dystocia results from the inability of the
uterus to respond to fetal signals appropriately and leads to
a delay in onset or completion of pup delivery (Narver, 2012);
it is one of the most common problems in rodent breeding
colonies. It should not be assumed that laboratory mice deliv-
ering during daylight hours are in dystocia; delivery of pups
is genetically based and may occur outside the night cycle
(Murray et al., 2010, Narver, 2012). Once identified, dystocia
is an emergency requiring intervention to preserve the life of
pups as well as the dam. Dams with dystocia may be noted
to have bloody vaginal discharge or may have pups actively
lodged in the vaginal opening.
Cross fostering (see relevant section on this topic) of surviv-
ing pups to another nursing female mouse is often required
for valuable pups of mothers that decompensate during par-
turition and require euthanasia.
B C
D E
DP
Fig. 2.16 The percutaneous insertion of gastrostomy tubes into

mouse pups. (A) The mouse pup is in the ventral view, with the
stomach and spleen visible. The milk-filled stomach of the pup is
easily seen through the transparent skin of the anterior abdominal
wall. (B) After adequate anesthesia, a small stitch in a U manner is
placed through the abdominal wall, incorporating the anterior wall
of the stomach, thus pulling the stomach tight to the abdominal
wall. (C) A needle is used to create a small stab wound through the
skin and into the anterior gastric wall. (D) A thin wire is passed
through the stab incision into the stomach, and polyethylene t ubing
is passed over the wire into the stomach. (E) The purse-string suture
is tied, and the wire and tubing are tunneled under the skin of the
abdomen toward the thorax. (F) The wire and tube are tunneled
under the skin of the thorax, out the nape of the neck, and the wire
is removed. The final tunneled gastrostomy tube is depicted in the
ventral, dorsal, and lateral views. (Reprinted by permission from
Macmillan Publishers Limited. Beierle, EA, Chen, MK, Hartwich,
JE, Iyengar, M, Dai, W, Li, N, Demarco, V, and Neu, J. 2004. Pediatr
Res 56:250–255.)
• Potential treatments: Different measures may be taken to

promote delivery, including physical removal of pups lodged
in the birth canal, which can be done manually with lubri-
cation and gentle retraction using forceps. Provision of sup-
portive sterile fluids (1–3 ml SC of warmed 0.9% NaCl or
lactated Ringer’s solution [LRS]), calcium gluconate salt to
increase contraction strength (100 mg/kg IP given 10 to15
min before oxytocin), with subsequent oxytocin (0.1–1.0 unit
SC) may help stimulate delivery and potentially improve the
health of the mother (Narver, 2012). It should be clarified that
little scientific evidence exists demonstrating the benefits of
oxytocin, a strong uterotonic drug, to dystocic mice (Narver,
2012, Schowalter et al., 2011); however, anecdotal reports
indicated it is administered frequently to dams in dystocia.
Administration of oxytocin may have adverse and confound-
ing effects on research, especially for behavior studies, and
administration may be contraindicated (Narver, 2012).
Despite the known critical role of prostaglandins in reg-
ulating murine parturition, prostaglandin therapy (2.5 µg
prostaglandin F2α [PGF2α] SC) has not been shown to be
more effective than oxytocin for alleviating dystocia (Chan
and Washington, 2011). Analgesics, from drug classes other
than nonsteroidal anti-inflammatory drugs (NSAIDs), may
be administered in addition to other supportive care mea-
sures mentioned. There is concern that administration of
NSAIDs, like ketoprofen, will inhibit cyclooxygenase, which is
responsible for production of the prostaglandins essential to
successful parturition in mice.
Promotion of environmental enhancements, including nest-
ing material (cotton bedding or paper strips), minimizing dis-
turbance to the cage, and judicious supportive care with fluids,
caloric supplements, and palatable softened food, in addition
to heating devices, may sometimes be sufficient to achieve
vaginal delivery by animals in good condition (Narver, 2012).
Any mouse in a compromised state during pup delivery should
be frequently monitored, up to every 1 to 2 h, to determine if
parturition is progressing and to assess health of the mother
(Schowalter et al., 2011). Options for management approaches
for dystocia (Figure 2.17) are provided (Narver, 2012).
For extremely valuable pups, the decision can be made to
perform a cesarean section for salvage of pups remaining in
Mouse Dystocia Reported
At cage change
(Disturbed) UNDISTURBED
With type Genetically

Observe throughout cage: If or unknown manipulated
bright, alert, and responsive (BAR),
DO NOT DISTURB
Recheck at end of shift;
± recheck next day; Physical
± provide supplements exam Dystocia
Dystocia expected or
expected unknown
NOT OBSTRUCTED
Productive Continued problems: Birth canal dilated;
Pup in birth canal characterize any Contact
delivery See UNDISTURBED Contact investigator
vulval discharge investigator
re: euthanasia or
re: treatment
treatment/C section;
If BAR and NOT
GOOD CONDITION: Poor proceed as directed
OBSTRUCTED
BAR, scant bleeding or condition (see GOOD
blood-tinged discharge CONDITION) OR
Lubricate good body condition euthanasia to
and manually Contact vet if remove from
remove pup(s) Nursing care + C section treatment or breeding pool
Contact C section is elected
OR investigator
Medical treatment depending on re:
veterinary assessment euthanasia
Productive Continued problems; and research study: Consider Ensure that Begin or continue
delivery see NOT OBSTRUCTED Is there behavioral work? breeder’s dystocia propensity tracking dystocias per
Is it a neurochemical study? age and action plan breeding pair for line
Is it a cardiovascular study? and colony are documented Consider breeder age
Document treatment results inbreeding in protocol and colony inbreeding
Fig. 2.17 Decision tree for dystocia management of mice. (Reprinted with permission from AALAS. Narver, HL.
2012. Oxytocin in the treatment of dystocia in mice. J Am Assoc Lab Anim Sci 51:10–17.)
the uterus; however, anecdotal reports indicated that these

females do not return to normal breeding status if they sur-
vive the operative procedure, and that pups do not typically
survive. If pups are collected in utero, they should be kept
warm. It is imperative to stimulate activity by stroking pups
gently with sterile soft materials (e.g., bandaging supplies or
gauze) to stimulate responsiveness and breathing. Prior to
placing pups with a foster dam, ensure that the animals are
pink, taking breaths, and are responsive to stimuli.
Fight Wounds
• Cause and impact: Aggression in group-housed male labo-
ratory mice is a widely recognized occurrence that can range
from mild to severe as a clinical concern (Van Loo et al.,
2003). In brief, male mice prefer social housing to individual
housing; however, dominant males in a group-housed cage
will show aggression toward subordinates. Aggressive behav-
iors have been linked to genetic background, odor cues, and
the lack of an available “escape” from human handling or
other mice within the cage. Injuries (Figure 2.18) are often
targeted along the dorsal rump and tail area, as well as in the
anogenital region.
A B C
Fig. 2.18 Fight wounds affecting the dorsal aspect of various male

mice. Wounds may be at the surface of the skin without readily
apparent hair loss (A); wounding may appear like ulcerative d
ermatitis
(B); healing of the lesions may occur by second intention, with
sloughing of the haired scab over time (C). Ventral aspects of these
animals should be physically examined to d etermine any further
extent of injury. (Images courtesy of University of Pennsylvania,
ULAR.)
• Potential treatments: Animals must be physically handled

and thoroughly examined to palpate the extent of potential
wound injuries both dorsally and ventrally.
Cohoused males with fight wounds should be separated
for the betterment of animal welfare, with the dominant male
(most often the sole mouse in the cage with no apparent
wounds) moved to an individual cage when aggression reaches
unacceptable levels and group-housed males have incurred
wounding.
Individual housing is recommended for highly aggres-
sive strains such as Swiss/CD-1 and FVB; however, the
sheer impact of isolation from conspecifics may lead to
increased aggression, and the addition of enrichment
devices to the cage may be of benefit. Nesting material is the
enrichment device of choice for group-housed male mice.

For g roup-housed mice with the dominant animal removed,
transference of used nesting material from the old housing
cage to the new cage with the dominant male can further
reduce a ggression. Interestingly, it has been suggested that
housing male mice in groups of three diminishes a ggression
(
compared to groups of five or eight), indicating that the
dominance hierarchy is more stable in smaller groups
(Van Loo et al., 2000).
With topical treatments similar to those applied for ulcer-
ative dermatitis (UD; see relevant section on this topic), sys-
temic analgesics (meloxicam 5 mg/kg daily for 3 to 5 days,
administered SC or by mouth [PO]), and separation from
aggressive mice, injured males very often recover to good
health, with scabbing and eventual sloughing of the wounded
skin.
Note: Males with severe wounds may need to be euthanized,
particularly if the anogenital region is scarred to a degree
that the animal cannot urinate due to injuries surrounding
the urethral opening and obstruction of urine outflow.
Fractures/Orthopedic Problems
• Cause and impact: Traumatic injuries related to fighting,
improper handling and procedures, entrapment in caging
equipment, and nutritional deficiencies may lead to broken
bones (fractures) and related orthopedic concerns. Affected
mice may present with a limp or lameness or other type of

gait abnormality. A broken tail may appear kinked, with little
evidence of swelling or bruising.
• Potential treatments: Depending on site of fracture (tail,
limb), animals should be placed under frequent observa-
tion, assessed for ambulation, provided with pain medication
(at least once daily for 3 days after the presumed fracture
occurs), and possibly imaged to identify the site of the lesion
by dual-energy X-ray absorptiometry (DXA) scanning or
microCT (Figure 2.19).
Rodents typically can ambulate well despite tail and limb
fractures and may not show overt evidence of distress. In time,
fractures should heal on their own, depending on location and
severity of the breakage. Bandages can be placed to immo-
bilize fractures; however, restraint collars (see Chapter 4,
“Restraint Collar Considerations”) may need to be placed to
prevent bandage removal (Hawkins and Graham, 2007).
Fig. 2.19 Fracture models of murine tibial injury and repair by

pinning may be imaged using DXA scans (A). This imaging tech-
nology can assess tail health and bone density (B), but greater
d istinction of vertebrae is capable using microcomputed tomography
(C). (Images courtesy of University of Pennsylvania; K. Hankenson.)
If the animal is unable to ambulate or typical mobility is

compromised, prompt euthanasia should be considered for
animal welfare.
Hemorrhage
• Cause and impact: Hemorrhage (active bleeding) may be
secondary to a number of physiological abnormalities, includ-
ing trauma, thrombocytopenia, or experimental treatments.
Certain mouse models of hemophilia (see relevant section on
this topic) may exhibit this clinical symptom following routine
procedures, like blood sampling. As well, bleeding may be due
to lacerations, secondary to fighting, or because of improper
hemostasis following tail clipping for genotyping.
• Potential treatments: Evidence of blood on any animal or
in the housing cage should require immediate attention to
ascertain the source and potentially to provide hemostasis
to stop continued blood loss. Depending on the degree and
source of blood, one can apply direct pressure to the site or
styptic p owder. Silver nitrate sticks are not recommended as
they tend to be an irritant and leave a persistent chemical
“burn” on the skin following use. Cautery applied using cord-
less disposable high-temperature loop tips (e.g., MediChoice®)
work well for small lesions as long as the animals are under
anesthesia at the time of application. If assisted wound
closure is necessary, it is recommended to use stainless steel
staples or tissue glue as routine sutures may be chewed out,
and bandages may be poorly tolerated. Consider application
of an appropriate size restraint collar on mice to prohibit
oral access to suture sites (see Chapter 4, “Restraint Collar
Considerations”).
Moribund, Weak, or Paralyzed Condition

• Causes and impact: Hind limb weakness (paresis) and
paralysis in laboratory mice may be associated with trauma,
dysfunction, and weakness of the musculoskeletal and
nervous systems, as well as infections caused by undesirable
colony pathogens that may particularly affect immunodefi-
cient strains. These animals may present with abnormal gait,
ataxia, or dragging one or more limbs during ambulation.
Fig. 2.20 An adult female BALB/c mouse, 12 days postinoculation

with herpesvirus, demonstrated neurological deficits, including trem-
ors and an inability to grip and place feet normally. The animal was
treated with supportive fluids and supplemental gel diet; the experi-
mental protocol did not permit the administration of anti-inflammatory
or analgesic medications due to interference with research outcomes.
Additional causes of weakness and paralysis may include

inherited neurologic diseases, like myelin disorders or neu-
ronal degeneration. Clinical signs develop in young mice car-
rying certain recessive mutations (i.e., jp/Y, shi, mnd, wst).
Neoplasia and nonneoplastic diseases, such as osteoarthri-
tis, bone fractures, or peripheral neuropathies (Figure 2.20),
may also occur, particularly with increasing age of animals
(Ceccarelli and Rozengurt, 2002).
Models of spinal muscular atrophy (SMA), a neurodegener-
ative disease of human children, have been established, with
SMA mice dying by 2 weeks of age if untreated. Mortality
has been linked to the phenotype of muscle weakness, but
secondarily to malnourishment as the affected pups are out-
competed for access to nursing during the preweanling phase
(Narver, 2011).
Moribund animals are typically those that are alive but non-
responsive to gentle manipulation by personnel, and they tend
to be isolated from cage mates. This state may be expected
for certain experimental models, but typically is irreversible
despite concerted efforts to administer supportive care.
• Potential treatments: Institutions and laboratories need to
ensure screening of biological materials prior to injection into
colony animals to ensure cells and murine-derived injectables

are pathogen free for agents, including MHV, mouse encepha-
lomyelitis virus, and lactate dehydrogenase-elevating virus
(LDV).
Certain infectious disease models may induce a moribund
state from which animals can recover. It is critical to increase
the frequency of monitoring and determine humane endpoints
that eliminate prolonged suffering (see Chapter 4, “Humane
or ‘Clinical’ Endpoint Considerations” and “Experimental
Autoimmune Encephalomyelitis and Demyelinating Disease
Model Considerations”). Animals that are paralyzed will need
to have their bladders expressed two to three times daily, and
topical lanolin ointment (Lansinoh®) should be placed on the
ventral abdomen and hind limbs to prevent or minimize urine
scalding of the skin.
Affected animals can be provided with fluid adminis-
tered subcutaneously and offered nutritional support at the
level of the cage floor (see Chapter 4. “Nutritional Therapy
Considerations” and “Fluid Therapy Considerations”).
Softened bedding substrates can be provided for comfort,
and particularly weakened or moribund animals should be
provided with heat and potentially separated to avoid further
injury from cage mates.
Mouse models, like that for SMA, can also be provided with
hand feeding to support nutritional requirements; antibiotic
and analgesic medication may also be provided as needed
(Wagner et al., 2011). Provision of ad libitum nutritional gel
supplements on the cage floor has been shown to improve
caloric intake and promote survival in neurodegenerative
mouse models with persistent tremors (Black et al., 2011).
More often than not, moribund and paralyzed animals will
require euthanasia if there is no improvement or change in
activity status within 24 h of initial presentation.
Mortality (Sudden Death)

• Cause and impact: Sudden death in mice is a common
occurrence, often without any premonitory signs, or may be
secondary to conditions like acute toxicity, degenerative dis-
ease, seizures, subclinical infection, or vascular dysfunctions
that were unrecognized prior to death.
There may be strain-related conditions of which to be aware,

such as spontaneous death in apparently clinically healthy
FVB/n mice with no premonitory symptoms at around 4
months of age. These mice have been described to have wet fur
below the mandible and on the ventrum of their necks at the
time of death. Histology on these animals identified multifocal
areas of neuronal necrosis and loss in the cerebrum, which
has been linked to the “space cadet syndrome” described
for FVB/NCR mice. While both genders are affected, female
FVB are more predisposed to sudden death. The condition is
thought to result from neuronal necrosis in the brain due to
seizure activity. Incidence of this condition may be underdiag-
nosed and should be considered in evaluation of FVB/n wild-
type and transgenic phenotypes (Rosenbaum et al., 2007).
• Potential treatments: While there is no treatment for ani-
mals that succumb to sudden death, the opportunity should
not be overlooked to necropsy animals that have recently died
(within 4 h) and perform histopathology on fresh tissues to
attempt to determine the root cause of death.
Ocular Lesions
• Cause and impact: Ocular abnormalities (Figure 2.21) are
frequently identified in laboratory mice and may appear
w ithout any obvious etiology. Eyelids may be squinted closed
over the eye; the globe itself may have alterations (ulcers) or
opacities (cataracts); there may be discharge noted; or an
altered size and shape of the eyeball may lead to exophthal-
mos (forward projection of the globe out of the socket). Any
abnormal swelling or mass development in or around the eye
should be reported. Animals will often appear to be otherwise
behaviorally normal despite the ocular lesion.
It is important to be aware that many common labora-
tory mouse strains (e.g., C3H, FVB/N, SJL/J, SWR, and
some outbred Swiss mice) are blind due to genomic muta-
tions (Danneman et al., 2012). Mice with microphthalmia
often have abnormalities in a variety of ocular structures;
this condition is common in C57BL/6 and related mice, with
increased incidence in females compared to males.
Additional things to rule out should include strain-
related disease, glaucoma, congenital abnormalities, trauma
A B
C D
Fig. 2.21 Representative ocular lesions that should be reported as

clinical cases for monitoring of progression and management of pain:
(A) cloudy left eye, (B) bulging or proptosis of the left eye, (C) retro-
orbital neoplasia of the right eye, and (D) corneal pitting and ulcer-
ation of the left eye, likely secondary to failure to apply ophthalmic
ointment while under anesthesia. (Images courtesy of the University
of Pennsylvania, ULAR.)
(
perhaps secondary to a recent retro-orbital bleed), com-
pound administration, light sensitivity due to any expected
neurological disorder, retro-orbital abscessation, or neopla-
sia. Acute reversible corneal lesions have been documented
in mice, attributable to a side effect of xylazine for anesthesia
(Calderone et al., 1986).
• Potential treatments: Certain ocular lesions can be avoided
through the routine use of eye lubrication ointment (e.g.,
PuralubeTM or Rugby® Sterile Artificial Tears Ointment
Lubricant–Ophthalmic Ointment) for any mouse undergoing
anesthesia for any procedure. This avoids desiccation of the eye-
ball that has been otherwise shown to lead to corneal opacities.
Routine prophylactic cleaning of the conjunctiva with
sterile swabs and saline will assist with removal of debris.
Fluorescein stain can be applied to check for corneal ulcers
and abrasions. Prior to staining, proparacaine (0.5%; 1 to 2
drops per eye) may be applied topically directly to the globe
for anesthesia.
Application of topical ophthalmic ointment, with or without

added antibiotics, is warranted as a first-line approach to an
eye injury. Certain lesions will be painful, with notation of
animals scratching at the eye and face; these animals should
receive topical anesthetic drops (proparacaine 0.5%) and sys-
temic analgesics (meloxicam 5 mg/kg SC) for pain relief.
If an animal presents with proptosis, the lid margins
around the globe should be retracted gently following clean-
ing of the eye surface and provision of topical ointment for
lubrication. Gentle pressure should be applied to the intact
globe to reduce the prolapse, and topical wetting drops and
ophthalmic antibiotics can be provided for 7 to 10 days after
replacement. Topical steroids should be avoided, but NSAIDs
appropriate for ophthalmic application can be considered
(Hawkins and Graham, 2007).
Surgical enucleation can be performed under anesthesia
for critical injuries, including ulcerations (Cote et al., 2011).
Perineal Swelling
• Cause and impact: Perineal swelling presents as an enlarge-

ment of subcutaneous tissues on the ventral and distal
abdomen in both males and females and is typically benign
(Figure 2.22). Male mice in particular may be reported for
Fig. 2.22 Representative perineal swellings that should be reported

as clinical cases for monitoring of progression, potential urinary
obstruction, and management of pain: female mouse with an absces-
sation of the clitoral gland (left); male mouse with abscessation of
the preputial gland (right). (Images courtesy of the University of
erineal swelling, which is believed to have a genetic compo-

p
nent, and there may be translocation of abdominal organs
into the perineal space or cysts within the bulbourethral
glands (Hill et al., 2002). Preputial gland abscesses second-
ary to bacterial infection have long been recognized as a
clinical issue in laboratory mice, and often Staphylococcus or
Pseudomonas are cultured (Hong and Ediger, 1978). Bacterial
agents may enter through the urethra or potentially through
fight wounds in males. Male mice have also been found to have
cholesterol granulomas with hemorrhage, in conjunction with
bulbourethral dilations (Dardenne et al., 2011). Females have
been reported to have clitoral gland abscessation (Alworth
and Nagy, 2009) and swellings that were diagnosed subse-
quently as mammary adenocarcinomas, likely strain related
in origin (Naff et al., 2005).
It is important to train animal care and investigative
staff to recognize these, and other, reproductive anomalies
in breeding colony mice. Because of the swollen perineum,
mice with imperforate vaginas are often mistaken as males.
The presenting complaint may be failure of a breeding unit
to produce pups, when the issue is actually that a normal
female has been paired with a second female with an imper-
forate vagina.
• Potential treatments: Many of these perineal swellings tend
to be incidental findings in mice that are otherwise bright,
alert, responsive, and in good body condition. Fine-needle
aspirates can be performed on the swellings to ascertain
if they contain purulent material, as many are caused by
secondary bacterial infections. While research data can be
obtained from animals with perineal alterations, if a genetic
component is suspected in affected animals, and if these mice
are in breeding colonies, it may be of benefit to cull affected
mice to avoid perpetuation of the condition in offspring
(Rubino et al., 2004).
If the swelling is due to imperforate vagina in the female,
the condition can be treated with surgery, yet with a limited
chance that the female will return to reproductive function
following surgery (Ginty and Hoogstraten-Miller, 2008):
• The vaginal membrane can be transected to release the
retained mucoid debris, and the newly formed orifice
can be further enlarged using blunt scissors to ensure
an adequate opening and good drainage. Take precau-

tion when flushing the vagina and uterus as these thin,
dilated, and flaccid tissues may be damaged.
• Supportive care involves instillation of triple antibiotic
ointment with hydrocortisone into the vaginal orifice,
followed by 1 to 2 drops of a local anesthetic agent (e.g.,
bupivacaine) along the incision site for 3 days.
• Antibiotics (trimethoprim-sulfa SC) can be administered
twice daily in 1 ml 0.9% saline for up to 7 days.
Realistically, the percentage of mice that return to breeding
soundness postsurgery is low, potentially due to the presence
of other reproductive abnormalities or to permanent damage
caused by the extreme distension of the uterus. Therefore,
surgical repair may be primarily incorporated as a salvage
procedure so that valuable females can be used for other
experimental manipulations.
Poor Body Condition

• Cause and impact: Mice may present with thin, hunched,
and ruffled appearance (Figure 2.23) without much forewarn-
ing; this may be related to a wide range of factors involving
anatomy and physiology, strain phenotype, and experimental
manipulations. Mice may have teeth that have overgrown due
to malocclusion and thus be unable to prehend and ingest
food. As well, immunodeficient strains may have underlying
(subclinical) infectious disease that should be considered as a
differential and may result in isolation of these animals from
the colony. Animals may have ruffled fur and hair loss due
to inflammation, autoimmune responses, pruritis and subse-
quent scratching, or due to ill health, such that the mouse is
not attending to self-grooming habits.
• Potential treatments: The logical and prioritized causes
should be treated first, typically including administration of a
bolus of fluids subcutaneously, nutritional support, and heat
supplementation. Dental assessments should be conducted
to determine if there is a loss of alignment of incisors (maloc-
clusion); if so, these teeth can be trimmed with nail scissors
and should be monitored for future overgrowth. Softened food
and supplements can be provided on the cage floor following
trimming.
A B
C D
Fig. 2.23 Mice can present with overall poor body condition, identi-
fied by lack of normal activity, ruffled fur (A, B, and D); hunched or
abnormal postures (A–D); or thin appearance in the range of a 1–2
on the BCS scale (A, C, and D). Often, animals in poor condition are
enrolled in experiments intended to cause an adverse outcome, or the
condition may be related to inherent immunosuppression or underly-
ing spontaneous disease, like tumorigenesis. A thorough h istory and
experimental description will be essential to compile the necessary
database of information to formulate a treatment plan. (Images cour-
tesy of the University of Pennsylvania, ULAR.)
Blood samples can be collected to assess serum chem-

istry and complete blood counts, as well as provide serum
to conduct a pathogen screen (Wiedmeyer et al., 2007).
Ectoparasites should be ruled out through skin scrapes, and
fungal cultures should be collected to discern if this might be
contributing to hair loss.
If the animal is not eating a provided experimental source of
food or pellet or if the treatments are potentially toxic or unpal-
atable, this should be further discussed with the research
team to ensure the animal is meeting its caloric needs.
If animals were expected to succumb to experimental
disease, the treatment efforts should be aimed at assist-

ing with comforting the animal in an attempt to achieve the
experimental endpoint. BCS should be assessed daily and
BWs checked routinely to track any further losses. Animals

that become quiet, less alert, and unresponsive should be
considered for euthanasia prior to reaching a moribund state
and spontaneous death.
Rectal Prolapse
• Cause and impact: An eversion of the rectal mucosa beyond
the rectal opening (Figure 2.24) is not an uncommon finding
in laboratory mice and may range from mild to severe enough
to warrant euthanasia. This prolapse may be due to strain-
related phenotypes, the efforts of parturition, or intestinal
infection (e.g., Helicobacter spp.) or other conditions that cause
diarrhea or straining to defecate. The mucosa may remain
moist and the prolapse actually identified due to adherence of
bedding substrate to the rectum in an animal that otherwise
has a normal body condition and activity level.
• Potential treatments: Husbandry management would
indicate that the bedding substrates should be changed to
softened paper materials for animals with rectal prolapses.
Any adhered bedding substrate should be removed from the
mucosal tissue to determine the severity and state of the
Fig. 2.24 Rectal prolapse of moderate severity; the mucosa is dark

red with pinpoint areas of hemorrhage. This mouse would need fur-
ther assessments to ensure normal defecation can occur, and topical
ointment should be applied to keep the tissue moistened. Changing
the bedding to a soft substrate could prevent further irritation of the
tissue. (Image courtesy of the University of Michigan, ULAM.)
prolapse, and moisturizing treatment with triple antibiotic

ointment or lanolin can be applied to dry and desiccated
tissues.
If pathogen infection is suspected (e.g., pinworms,
Helicobacter, or Citrobacter), treatment with antiparasitics or
antibacterial therapy appropriate to the pathogen should be
instituted promptly.
Successful correction of rectal prolapse has been described
using a technique that permits the repositioning of the rectum
inside the anal cavity, thus preventing tissue cyanosis and
necrosis (Koch, 2010). Topical triple antibiotic ointment with
hydrocortisone is first applied to the perineal area to reduce
inflammation. The rectum is then pushed back inside the
body using a blunt probe, then a small amount of tissue glue
is applied on the borders of the anus to temporarily maintain
the rectum in place and prevent it from prolapsing when the
mouse is passing feces. Daily application of antibiotic ointment
topically at the site is recommended, along with daily admin-
istration of an injectable NSAID (e.g., carprofen or meloxicam
5 mg/kg SC) for 3 consecutive days if this analgesic choice
does not interfere with the research outcomes (Koch, 2010).
If rectal prolapses are prevalent in particular strains, scor-
ing systems for severity of rectal prolapse to determine points
for humane intervention may be beneficial (Figure 2.25).
Respiratory Distress
• Cause and impact: The respiratory system for mice has been
reviewed (Kling, 2011), and adverse clinical signs dependent
on some aspect of the respiratory system can include nasal
discharge, ocular discharge, sneezing, audible “chattering,”
dyspnea, open-mouth breathing, cyanosis, and head tilt.
As mice are obligate nasal breathers, the development of
respiratory distress in the face of infectious pulmonary disease
may be rapid. Infectious agents that may be i mplicated include
viruses (e.g., Sendai virus) and bacteria (e.g., Mycoplasma). If
the animals are deemed to be in stable condition (not overly
stressed), one may consider using imaging methods of radi-
ography to ascertain if there are consolidations, opacities, or
other abnormalities in the lung field that are contributing to
the condition.
Rectal Prolapse
SEVERE
MILD MODERATE
Animal: Quiet, alert,
Animal: Bright, alert, Animal: Bright, alert,
responsive
responsive responsive
Prolapse Size:
Prolapse Size: Prolapse Size:
Protrudes 1–2 mm Protrudes >7 mm from
Protrudes 3–7 mm from
from anal opening; anal opening;
anal opening;
Tissue status: Tissue status:
Tissue status:
Moist, pink Ulcerated, dry, dark
Moist, pink
red to black
Recommend
May apply triple euthanasia
Consult with
antibiotic EOD to principal
May apply triple
maintain moisture investigator
antibiotic ointment
of tissue; switch to determine
EOD to maintain
bedding to paper preferred
moisture of tissue; course of
product to reduce
monitor progression action for
irritation of tissue.
the
Is this a valuable,
treatment
irreplaceable or plan
pregnant animal ?
NO YES
Recommend euthansia Continue to monitor progression and

provide daily treatments
Fig. 2.25 Decision tree for management of rectal prolapse in rodents.

(Modified from University of Washington, DCM.)
• Potential treatments: Respiratory distress in any animal

warrants prompt administration of supplemental oxygen; it is
recommended to place rodents in a small induction chamber
or place the entire animal inside a large anesthetic face mask
to deliver oxygen rapidly. The chamber can be kept somewhat
cool relative to ambient temperatures as animals in distress
may have increased core temperatures (hyperthermia) due to
their respiratory efforts. Rodents should be observed closely
for improved breathing patterns and changes in mucous
membrane color toward a pink/red hue.
If an undesirable infectious pathogen is suspected as the

root cause, it is recommended to quarantine the affected ani-
mal and potentially perform euthanasia, tissue harvest, and
serology to diagnose the agent. These diagnostics may be
critical if there is concern of pathogen spread into additional
mouse colonies in the facility.
For certain animals, depending on the experimental plan,
it may be of use to provide light sedation to calm the respi-
ratory effort and diminish anxiety. It may be appropriate to
administer diazepam (1–3 mg/kg IP) or midazolam (0.5–1.0
mg/kg IP) as described (Oglesbee, 2011). If a bacterial cause
is confirmed, antibiotic sensitivity testing will direct selection
of appropriate systemic antibiotic treatments.
Seizures
• Cause and impact: Generalized seizures are caused by
paroxysmal cerebral dysrhythmias and are characterized by
loss of consciousness, muscle contraction (tonus), and jerking
(clonus) (Aldrich, 2005). Seizures may present in animals with
a sudden onset of shaking or chewing; with circling, momen-
tary paralysis or “freezing”; or with convulsions (Figure 2.26).
Episodes typically are brief and may arise spontaneously
after handling of rodents; they can be accompanied by auto-
nomic dysfunction (urination/defecation). In status e
pilepticus,
seizures occur in rapid succession without recovery between
them; this intense neuronal activity can cause metabolic
derangements with damage to neurons and brain swelling.
The exertional muscular activity during seizures can predis-
pose the rodent to hyperthermia, hypercapnia, hypoxemia,
and metabolic acidosis (Vernau and LeCouteur, 2009).
Models of epilepsy (Fisher, 1989) may be desired for certain
experimental protocols, and seizures may be linked to the
transgenic or knockout strain or may be due to development
of a deleterious mutation (Pesapane and Good, 2009).
Seizures in FVB mice have been described (Goelz et al.,
1998); observations of seizure activity were made of mice
while in their cages, when handled for tail tattooing and fur
clipping, as well as during facility fire alarms. The majority
of affected animals were female FVB/N. Clinical presenta-
tions included facial grimace, chewing automatism, ptyalism
Fig. 2.26 Sequence of events leading to full seizure in an affected

mouse: (A) After handling, the mouse is placed on the top of a
Shepherd Shack® (Shepherd Specialty Papers, Portage, MI) and starts
to experience paralysis. (B) Paralysis worsens, and the animal’s
mouth opens; the mouse twists and shakes. (C) The animal falls
from the Shepherd Shack and is convulsing with its mouth open. (D)
The animal is fully recovered after approximately 5 min. (Reprinted
by permission from Macmillan Publishers Limited. Pesapane, R, and
Good, DJ. 2009. Seizures in a colony of genetically obese mice. Lab
Anim (NY) 38:81–83.)
with matting of the fur around the neck and on forelimbs,

and clonic convulsions that, at times, progressed to tonic
convulsions and death. In some mice, only nonspecific signs
of disease were noted, such as lethargy, moribund state, and
matting of the fur (from hypersalivation); blood glucose values
remained within normal limits.
Animals may undergo multiple unobserved seizure
episodes, culminating in a terminal lethal convulsion; these
animals will likely be found dead yet appear in good body
condition without other evidence of an overt cause of death.
In keeping with reports of finding spontaneously dead FVB/N
mice, it has been documented that susceptibility to seizure
activity increases during the dark cycle, when the laboratory
rodents in a typical research facility would not be routinely
observed by personnel who could otherwise track and report
on seizure activity (Goelz et al., 1998).
Generalized spontaneous seizure disorders in mice have
been documented in dilute lethal (dl), quaking (qk), and
wobbler-lethal (wl) strains. Certain inbred strains, including
DBA/2J, SJL/J, and LP, may have generalized lethal seizures
when exposed to auditory stimuli (Fuller and Sjursen, 1967).
Audiogenic seizure susceptibility varies widely between
strains and appears to be influenced by genetic and environ-
mental factors (Goelz et al., 1998).
• Potential treatments: Neurologic examinations in rodents are
not typically easy to perform and may be based on observations
of return to clinically normal behaviors after a seizure, includ-
ing returns to expected eating and drinking patterns and
activity. It is important to note that animals may be neurologi-
cally abnormal for days after a seizure (Vernau and LeCouteur,
2009), which can have impact on the collection of accurate
experimental data and the overall model under investigation.
As well, once an animal has seizured, it may be indistinguish-
able from normal cage mates within a period of a few hours.
Questions to consider when trying to determine the history of
the seizures (particularly if noted in a colony of genetically sim-
ilar animals) include the following (Pesapane and Good, 2009):
• Is this condition linked to a gene of interest in this line of
mice?
• Is it correlated with gender or age at onset?
• Could there be an environmental or infectious cause?

• Is this condition heritable?
Emergence of seizures in a colony that otherwise has not
previously been reported should instigate a review of breeding
record keeping to determine the frequency of occurrence and
to determine if this trait originated with a single breeding pair
linked to a particular cohort of offspring maintained within
the colony.
Drug treatments are not typically described in mice,
although diazepam is a first-line agent of treatment in other
species (Vernau and LeCouteur, 2009). If seizures are unre-
sponsive to benzodiazepines, one can attempt treatment with
phenobarbital (4 mg/kg intramuscularly [IM], give twice
at 20 min apart; continue this 12 h later at 2 mg/kg PO).
Dextrose may also be administered if hypoglycemia is present
postseizure (Hawkins et al., 2007).
Trauma
• Cause and impact: As mentioned in other sections, traumatic
injuries in routinely housed laboratory mice may be caused by
improper handling or intracage fighting, improper administra-
tion of experimental agents (e.g., tumor cells injected extravas-
cularly), or entanglement in caging equipment (Figure 2.27).
Fig. 2.27 Trauma to the tail caused by inappropriate tail vein injec-
tions of tumor cells (left) and by damage from cage mates follow-
ing injury from cage materials (right). These animals should receive
pain management and potentially surgical tail amputation to remove
the affected necrotic tissues. (Images courtesy of University of
Fig. 2.28 Trauma to the ears related to metal ear tag placement.

A nude mouse ear became entrapped in cage materials, resulting
in avulsion and necrosis of the tissue, which required amputation
of the ear (left); foreign body reaction to the metal tag may result in
self-injury and scarring (right). It is recommended that the ear tags
be removed if an allergic response is suspected. (Images courtesy of
Trauma may have an impact on any part of the body,

including the face, limbs, paws, or ears, often due to metal-
lic identification ear tags becoming stuck in wire bar feed-
ers, j-feeders, and wire bar lids. These tag entrapments are
precipitated by the routine exploratory and climbing activities
in which mice engage as part of their species-specific behav-
ior (Figure 2.28).
Animals may also self-injure, related to their attempt to ame-
liorate painful sensations, and cause severe self-trauma to eyes,
to existing dermatitis, or to limbs that may have become injured
secondary to environmental or experimental influences.
• Potential treatments: Animals should be rescued from any
compromising entrapment in the cage as soon as it is noted
and then further evaluated for injury and potential fractures
to legs and tail. Any active hemorrhage should be immediately
assessed for source of the bleeding, and manual h emostasis
should be provided.
Ears may be avulsed and necrotic following entrapment;
removal of pinnal tissue can be done with pen-like cautery
tools to provide a clean dissection of the affected tissue away
from healthy tissue. Animals should be under systemic anes-
thesia prior to the cautery recommended for amputation of
ear tissue.
Animals that potentially drop a distance when being han-

dled by personnel should be recaptured and examined for
injury; in most contemporary housing facilities, it is typically
not acceptable to place an animal that has fallen onto the
floor or out of the flow hood or biosafety cabinet back into
the original cage due to biosecurity concerns. Instead, these
animals should be singly housed (quarantined) for observa-
tion and isolated due to their potential contamination from
exposure to macroenvironmental elements.
Ulcerative Dermatitis
• Cause and impact: In the research animal community,
there are diverse anecdotes and hypotheses about the etiol-
ogy of ulcerative dermatitis (UD). In contemporary facilities,
dermatitis is noted in mice from multiple background strains,
not just those with a C57BL/6 background. Development
of severe lesions can occur rapidly; areas of ulceration can
expand, most often due to self-mutilation, in the course of
less than 24 h. The more commonly noted sites of ulcerative
dermatitis are between the scapulae and on both dorsal and
ventral aspects of the neck area; however, facial dermatitis
is not uncommon (Figure 2.29). A chronic nidus of inflam-
mation, such as that occurring from oral mucosa pierced
by shed hairs during grooming, has a significant link to the
presence of ulcerative dermatitis (Duarte-Vogel and Lawson,
2011, Lawson, 2010).
Ulcerative dermatitis has been reviewed comprehensively
(Hampton et al., 2012, Kastenmayer et al., 2006), and the
most important underlying factors have been documented to
best synergize treatment modalities. The skin disorder has
been attributed to some combination of infectious, genetic,
behavioral, nutritional, immunological, endocrinological,
environmental, and neurological factors. Neurological “skin-
picking” disorders have been implicated as well. (Dufour
et al., 2010).
Histological assessments of UD lesions from C57BL/6 mice
have indicated that the earliest detectable lesions involve fol-
licular dystrophy, with degradation of the inner root sheath
and defects in the hair fiber cuticle that may puncture the fol-
licles and lead to inflammatory reactions (Taylor et al., 2005).
Fig. 2.29 Typical appearance of ulcerative dermatitis in mice.

(Left) Mouse with facial dermatitis around the eyes, with scarring
and retraction of the mandible due to lesions under the chin; nasal
abrasions are likely secondary to excessive grooming activities.

(Reprinted with permission from AALAS. Lawson, GW. 2010. Comp
Med 60:200–204.) (Right) Mouse with severe ulcerative lesions and
hair loss over the scapulae, affecting at least 10% of the body area.
(Images courtesy of University of Michigan, ULAM.)
Progressive necrosing dermatitis of the pinna has been

described in CD-1 outbred mice; initial lesions may be subtle
and thus missed. However, peripheral necrosis of the pinna
may develop with serous discharge, and the pruritus may
cause the mice to self-injure and excoriate the area (Slattum
et al., 1998).
• Potential treatments: Importantly, one should rule out
other infestations, such as mites, that may lead to pruritis
and ultimately dermatitis (Smith et al., 2003). Reports of
attempted treatment (e.g., topically applied corticosteroids,
antibiotics, or antiseptics) for pinnal necrosing dermatitis
may not eliminate lesions but may help to control progression
(Slattum et al., 1998).
It is critical in treatment of UD to understand that no
application will likely eradicate all signs in all animals;

however, even partial resolution or inhibition of further devel-
opment can be an improvement to animal welfare and avoid
the outcome of euthanasia prior to the animal reaching its
anticipated experimental endpoint.
A comprehensive list of UD treatment types and options
is provided next (in alphabetical order, depending on
administration route, and with further specific instructions

from available resources):
• Topical application (daily or EOD depending on severity of
lesions):
−− Antibiotics: Selection should be based on culture and
antibiotic sensitivity results of skin lesions; Neosporin
or similar products are most commonly used as a front-
line approach.
−− Calamine liquid suspension with zinc oxide (cala-
mine lotion).
−− Caladryl® (camphor/pramoxine/zinc):
−− Main advantage of Caladryl is mitigating the scratch-
ing to allow lesions to dry and heal; improvement has
been seen in up to 65% of cases (Crowley et al., 2008).
−− Clean affected skin gently with chlorhexidine solu-
tion with sterile gauze.
−− Apply Caladryl once daily in thin layer for 5 to 7 days
using clean cotton swabs or clean gloved finger.
−− Healing ensues sequentially after cessation of
scratching, with resolution of erythema and drying
of lesions.
−− Treatment should continue after lesions have healed
grossly and should continue or be reinstituted if
scratching is noted.
−− Treatment may be less successful in mice with mul-
tiple UD lesions or advanced full-skin-thickness
lesions.
−− Chlorhexidine (0.2% solution):
−− Dilute 1 ml 2% chlorhexidine gluconate in a test
tube containing 9 ml distilled water (1:10 dilution)
and shake thoroughly.
−− Using sterile cotton swab or gauze pad, apply a
generous amount to affected areas once daily for
7 to 14 days, followed by application of topical tri-
ple antibiotic ointment for severe cases (Lumpkins
et al., 2006, Delgado et al., 2011).
−− Mice were noted to stop scratching within 3 days of
this treatment, and most treated mice were lesion
free for up to 7 months.
−− Cyclosporin (0.2%) in a 2% lidocaine gel:

−− Apply mixture topically and supplement with 50 µg/
ml gentamicin twice daily (Feldman et al., 2006).
−− Derm Caps®: A proprietary combination of omega-3
and omega-6 fatty acids (contains safflower oil, a source
of linoleic acid; borage seed oil, a source of gamma
linolenic acid; fish oil, a source of e
icosapentaenoic and
docosahexaenoic acid; and vitamin E).
−− Diabetic foot ulcer medications: May be beneficial
due to inclusion of platelet-derived growth factors
−− EMLA® (Eutectic Mixture of Local Anesthetics): A mix-
ture of 2.5% prilocaine and 2.5% lidocaine cream that
provides topical anesthesia with prolonged contact
time (>15 min).
−− Gentian violet (GV): An anti-infective ointment in an
alcohol-based solution.
−− Green clay (montmorillonite):
−− Clay is mixed with water to reach a thickened
consistency and should be allowed to stand for 1 h
prior to use.
−− Shave around the affected skin area (with mice
under anesthesia) and then clean shaved area with
dilute chlorhexidine and allow to air dry.
−− Apply clay poultice to a thickness of 5 mm and
reapply every 3 to 4 days, as long as animal does
not remove clay (Figure 2.30).
−− Improvement in UD was noted that ultimately
resolved in 123 of 150 similarly treated mice (Martel
and Careau, 2011).
−− Lansinoh: Lanolin ointment used for keeping skin
moist. It serves as a nontoxic emollient and is success-
ful for treatment of dry skin for pups and nude mice
(Taylor et al., 2006).
−− Natural unpasteurized honey (Mathews and
Binnington, 2002).
−− New Skin®: Liquid bandage substance.
−− Panalog®: Contains antibacterial, anti-inflammatory,
and antifungal components.
−− Silvadene®: A 1% silver sulfadiazine ointment.
A B
C D
Fig. 2.30 Green clay therapy for mice with UD: (A) untreated mouse;
(B) treatment of UD with topical green clay; (C) healing by day 4
postapplication; and (D) appearance of the mouse 3 months later.
(Reprinted with permission from AALAS. Martel, N, and Careau, C.
2011. Tech Talk 16:2–3.)
−− Sterile saline (hypertonic): To remove debris and

bacteria.
−− Place 100 ml distilled water in screw-cap bottle.
−− Add 4.4 g of sodium chloride and shake well; loosely
screw cap on bottle and steam sterilize bottle on
liquid cycle in an autoclave.
−− Apply saline with sterile cotton ball; apply to crusts
and ulcerative areas twice daily for 2 to 3 days.
−− Flush remainder of lesion using a sterile syringe or
pipette; as healing progresses from granulation to
epithelial tissue, treatment can be reduced to EOD
(Mangete et al., 1992).
−− Steroids: To combat inflammation.
• Systemic: May include antibiotics, anti-inflammatories,
antihistamines, and analgesics, along with the following:
−− Vitamin E
−− When added to food (dosed at 1,600–3,400 IU/kg),
has been shown to improve lesions greater than
Day 1 Day 15 Day 50
Water
1
mg/kg
Fig. 2.31 Progression of healing UD lesions; animals on the bottom

row were treated with systemic maropitant citrate (dosed at 1 mg/kg
IP) compared to controls in the top row. (Reprinted with p
ermission
from AALAS. Williams-Fritze, MJ, Carlson Scholz, JA, Zeiss, C,
Deng, Y, Wilson, SR, Franklin, R, and Smith, PC. 2011. J Am Assoc
Lab Anim Sci 50:221–226.)
50% at the low-end dose (Barnard et al., 2006,

Lawson et al., 2005); Derm Caps® can be the source
of v itamin E, with 0.1 ml/day dripped onto a food
pellet.
−− Maropitant citrate: A neurokinin 1 (NK1) receptor
antagonist.
−− For lesion improvement (Figure 2.31), dose once
daily at 1 mg/kg IP for 5 to 10 days (Williams-Fritze
et al., 2011).
• Other miscellaneous adjunctive therapies:
−− Toenail clipping (especially rear feet):
−− It is well documented that reduction in self-
mutilation from scratching can diminish progres-
sion of dermatitis (Disselhorst et al., 2010, Martel
and Careau, 2011, Mufford and Richardson, 2009).
−− To avoid scruffing the mice over the ulcerated areas
for restraint purposes, a 50-ml conical tube can be
used as a restraint device that p rovides access to
the rear feet (please ensure holes are drilled in the
end of the tube for air exchange).
Fig. 2.32 Performance of hind limb toenail clipping to diminish fur-

ther self-injury in mice with ulcerative dermatitis. (Reprinted with
permission from AALAS. Martel, N, and Careau, C. 2011. Tech Talk
16:2–3.)
−− Small scissors, potentially ocular scissors, are rec-

ommended to avoid removing more tissue than the
toenails (Figure 2.32).
−− Alternatively, nail trims may be performed on rodents
during anesthetic recovery from other procedures.
−− Removal of ear tags: This can be done with wire cut-
ters or using microscissors inserted into the closed tag,
followed by opening of the scissor blades to pop open
the tags for removal.
−− Sterilization of food and bedding: To diminish
potential hypersensitive elements or contaminants
(infectious pathogens).
−− Environmental manipulations: Could involve the
provision of cage enrichments to distract from scratch-
ing and a change to hypoallergenic bedding materials.
Scoring systems for UD (Figure 2.33) are helpful, consid-
ering that many of these lesions persist for weeks to months
without noticeable improvement, despite a v
ariety of treatment
attempts (Hampton et al., 2012). Given the variable success
rate for eradication of UD, it is recommended that humane
endpoints be established if the lesions develop to involve the
eyelids, if the lesions are deep enough such that muscle lay-
ers are exposed, if greater than 20% of the body is affected,
A Scratching Number (S number)* Score

None 0
<5 1
5–10 2
> 10 3
* Number of scratches in a 2 minute period.
B Character of Lesion (COL) Score

No lesion present 0
Excoriations only or one, small
1
punctuate crust (≤ 2 mm)
Multiple, small punctuate crusts or 2
coalescing crust (>2 mm)
Erosion or ulceration 3
C Length of Lesion** Score
0 cm 0
< 1 cm 1
1 cm–2 cm 2
> 2 cm 3
**Length of lesion is determined by measuring the longest

diameter of the largest lesion identified. This measurement
should involve the lesion only and not cross over clinically
normal skin.
D Regions Affected Score
None 0
Region 2 or 3 1
Region 2 and 3 2
Region 1 +/– other affected regions 3
E Calculated Severity Score
[(A + B + C + D) ÷ 12] × 100
Fig. 2.33 Ulcerative dermatitis scoring system. Simplified descrip-

tion for (A) scratching number, (B) character of lesion, (C) length
of lesion, and (D) regions affected. Scores from A, B, C, and D are
used in the (E) UD scoring system formula to generate the calculated
severity score (Reprinted with permission from AALAS. Hampton,
AL, Hish, GA, Aslam, MN, Rothman, ED, Bergin, IL, Patterson, KA,
Naik, M, Paruchuri, T, Varani, J, and Rush, HG. 2012. J Am Assoc
if ambulation is affected (particularly through wound con-

traction around a limb), or the treatments have continued
for longer than 4 weeks without improvement (see Chapter 4,
“Humane or ‘Clinical’ Endpoint Considerations”).
Vaginal or Uterine Prolapse

• Cause and impact: Subsequent to parturition, female mice
may exhibit vaginal prolapse, or even uterine prolapse
(Figure 2.34), with eversion of tissues outside the external
orifice. Typically, although these mice are ill, often exhibiting
pyloerection, hunched posture, and respiratory dysfunction,
they often are still able to actively nurse their pups. It may be
unclear on initial report which tissue has prolapsed; thus, it will
be important to involve a veterinary assessment to distinguish
a severe rectal prolapse from a vaginal/uterine prolapse, which
otherwise may appear to be the same to untrained personnel.
• Potential treatments: If the female is in poor health and
the prolapse is severe, euthanasia may be warranted, and
any pups should be cross fostered to an appropriate dam (see
relevant section on this topic).
Fig. 2.34 Prolapsed uterus secondary to delivery of a litter; corn-

cob bedding is adhered to the prolapsed tissue. (Image courtesy of
When the animal is relatively bright, alert, and behav-

ing normally, one can consider correction of the prolapse
performed as follows (Castro et al., 2010):
• Reduction of prolapse is done under anesthesia; isoflurane
is most preferred for its rapid induction and wide safety
margin. Analgesia is provided with carprofen (5.0 mg/kg
SC) and fluid replacement (1.0 ml 0.9% NaCL SC).
• Exposed mucosal surfaces should be gently rinsed with
saline and cleared of any adhered materials (bedding, etc.)
using forceps.
• Topical triple antibiotic ointment should then be applied
to the clean tissues, and a lubricated 0.75-inch, 20-gauge
polytetrafluoroethylene intravenous catheter (without nee-
dle) is placed into the lumen of the uterus and vagina.
• Utilizing the catheter, the exposed tissue is manually
manipulated to invert it into the proper anatomic position.
• Purse-string sutures, with 4–0 polyglycolic acid suture,
are then placed around the vaginal orifice prior to removal
of the catheter to prevent inadvertent recurrence of tissue
eversion.
• Postprocedurally, mice can be administered a 3-day
course of carprofen (5.0 mg/kg SC) and oral antibiotics.
research-related medical issues

Additional topics concerning laboratory rodent health that warrant
further information herein due to their prevalence in c ontemporary
research environments are listed next in alphabetical order to
facilitate their location. Under each topic, “background” informa-
tion is provided, and “potential treatments” offer suggestions about
procedures, therapeutic treatments, and further considerations.
Ascites Production
• Background: Tumor-producing and ascites-producing cell
lines are injected into mice as a method to produce antibod-
ies as a research reagent; this in vivo technique typically is
used when no other in vitro method can be implemented. Any
animal cell lines should be tested and demonstrated free of
murine pathogens and other transmissible agents that poten-

tially could contaminate animal colonies, infect humans, or
introduce unwanted experimental variables. If animals are
not monitored appropriately, ascites production can be life
threatening due to tumor growth, metastatic spread, infiltra-
tive growth, and ultimately respiratory distress.
• Potential treatments: The essential treatment for this model
is to ensure that mice are monitored for weight changes and
development of abdominal swelling due to ascites. Ascites
production most commonly occurs within 7 to 14 days after
the cells are injected, and the following are suggestions for
management of these animals:
• On the day of and prior to cell inoculation, the mouse
should be weighed; this weight is recorded as the “initial
weight.”
• Mice that have been inoculated must be weighed at regu-
lar intervals; these intervals should be as described in the
IACUC protocol and based on the expected rate of abdomi-
nal fluid accumulation.
• Daily clinical observations should be made for assess-
ments related to changes in posture; activity; food and
water intake; respiratory patterns (labored, depressed,
or accelerated); body condition (e.g., rough hair coat, pale
ears or eyes); and severe abdominal distention.
• Ascites fluid is to be collected before BW becomes 20%
greater than the initial weight or abdominal distention
leads to significant health problems.
• Fluid should be harvested following antiseptic preparation
of the site using 18- to 22-gauge hypodermic needles. Each
time the abdomen is tapped, a fresh disposable needle and
syringe are to be used.
• Warm saline or LRS (2–3 ml SC) may be given at the time
the animal is tapped to avoid hypovolemic shock if large
volumes of ascitic fluid are removed.
• Mice should be observed for at least 30 min following
a tap.
• The limit to the number of allowable abdominal taps
should be established in the IACUC protocol; the recom-
mended maximum is three taps, with the third tap per-
formed after euthanasia.
• If the abdominal tap does not relieve abdominal disten-

tion, the abdomen of the mouse should be gently palpated
to determine if distention is due to solid tumor growth.
Clinical signs of hypovolemic shock include hunched pos-
ture, roughened hair coat, anorexia, dehydration, weight loss,
loss of body condition, inactivity, difficulty in ambulation, pal-
lor of the ears and eyes, tachypnea, and dyspnea. Persistence of
these signs after tapping of the abdomen warrants immediate
notification of the veterinary staff or consideration of euthana-
sia of the animal. Mice should be euthanized promptly if asci-
tes fluid becomes blood tinged or turbid or if mice show signs
of poor condition, such as huddling, ruffled coat, or inability
to reach food and water (IACUC-UPENN, 2010).
Experimental Autoimmune
Encephalomyelitis Mouse Models
• Background: As a model for multiple sclerosis (MS), the
study of experimental allergic encephalitis (EAE) in mice is
commonly undertaken. This model may also be referred to
as experimental allergic encephalomyelitis, which has the
same acronym, EAE. MS is believed to be an autoimmune
disease mediated by autoreactive T cells with specificity for
myelin antigens. Animals are expected to become weak and
may develop an acute, chronic, or relapsing-remitting disease
course. In general, the disease progresses with ascending
paralysis, dysfunction in normal ability to eat due to lesions
in cranial nerves, and lingual paralysis. Animals may lose
BW quickly and will likely develop poor overall condition due
to inability to masticate food or access provided feedstuffs.
• Potential treatments: As disease progresses to excessive
weakness, paralysis, and weight loss, softened food can be
delivered via oral gavage, and 0.9% NaCl SC can be injected
for supportive care. Additional softened food should be placed
on the cage floor, water bottles should have elongated sipper
tubes to facilitate access by an animal that cannot stand,
and urinary bladders should be manually expressed at least
twice daily by personnel (Miller and Ito., 2011).
Scoring mechanisms for disease progression may be use-
ful for determining the humane endpoints of the experi-
ment; more information is available in Chapter 4, “Humane
or ‘Clinical’ Endpoint Considerations” and “Experimental

Autoimmune Encephalomyelitis and Demyelinating Disease
Model Considerations.”
Hemophilic Mouse Models

• Background: Mouse models for hemophilia demonstrate pro-
longed blood clotting times of at least 30 min (the clotting
time for a healthy C57BL/6 mouse is less than 5 min). This
delay in clotting causes hemorrhage when mice are wounded,
potentially due to lesions associated with fighting, experimen-
tal injection sites, or spontaneous bleeds. Mice may present
with blood in the cage; weakness and lethargy; pallor to the
paws, mouth, or gums; and a bloated or swollen appearance,
attributed to hemoabdomen.
• Potential treatments: Hemorrhage should be prevented by
keeping compatible animals in the same cage (with the intent
to eliminate aggression and fighting), maintaining pressure
over a site that has the potential to bleed for a prolonged time
period after a procedure, and handling and monitoring ani-
mals carefully to avoid trauma. Bleeding sites may be cleaned
and cauterized, with administration of subcutaneous fluids,
if necessary (Jones et al., 2005). Silver nitrate sticks are not
advisable for use in mice due to adverse skin irritation; stud-
ies have shown success in achieving hemostasis in factor VIII
knockout mice using ferric sulfate in a gel formulation (sup-
plied in a 1-ml syringe with a small, flat-tipped applicator).
This gel formulation has been shown to be superior to the use
of fiber cellulose pads, aluminum chloride (both powder and
liquid formulations), and ferric subsulfate (both powder and
liquid formulations) (Turner et al., 2011).
In a study of hemophilic PL/J mice (factor IX deficient),
D-dimer, a highly specific and reliable test for the diagnosis
of thrombotic conditions such as disseminated intravascular
coagulation, can be applied for hemophilic mice (Trammell
et al., 2006).
Obese Mouse Models

• Background: For mice that are genetically engineered or
spontaneously develop overweight conditions, consider-
ations should be made for a variety of aspects of their care.
More than 50 different rodent models of obesity are avail-

able for use in biomedical research projects (Good, 2005).
Some obese mice may grow to 60–70 g BW and therefore,
as per housing density standards of the National Research
Council (NRC) guide (NRC, 2011), should be housed with
fewer cage mates. In particular, for obese rodents, one must
take into account the type of housing, the housing condi-
tions (group or singly housed), cage shelf level, the room
temperature, and environmental enrichment as all of these
issues can have an impact on behaviors, food intake, and
BW gain (Good, 2005).
Diabetes, which can accompany the obese phenotype, typi-
cally results in increased water consumption and in increased
urine production. As well, fluctuations in room temperatures
can have an impact on weight gain, fat deposition, and core
body temperatures for obese rodents.
Choices of anesthetic agents used for obese rodents should
take into account the agent’s fat solubility as overweight
a nimals and certain knockout mice will display obesity and
reduced sensitivity to certain anesthetic agents (e.g., pento-
barbital and tribromoethanol). Performing surgery in these
compromised mouse models may result in adverse effects like
hypoglycemia, difficulty with dehiscence of incision sites, and
anesthesia effects, including delayed recoveries and poten-
tially fatal outcomes due to depressed respiration linked to
slower metabolism of drugs.
• Potential treatments: Providing supportive care and spe-
cialized environments will be best for these obese animals.
For animals with evidence of diabetes, more frequent cage
changes and provision of more absorbent bedding sub-
strates should be used in the mouse cages to compensate for
increased urine production.
Environmental temperatures and humidity should be
closely monitored and potentially lowered for those animals
with obesity.
For those obese mice undergoing surgery, attention to the
minimal time for fasting, both presurgical and postsurgical, is
key (see Chapter 4, “Fasting Considerations”). To prevent dehis-
cence of surgical sites, it is recommended to avoid surgical skin
clips for this model and instead close incisions with continuous
suture patterns, using a minimal suture size. Maintaining the
anesthetic dose of isoflurane to no more than 1.0%, with an

oxygen flow rate of 0.4 L/min, can facilitate recovery issues
and maintain animals at a reasonable depth of anesthesia. As
well, supplemental heat should be provided as outlined previ-
ously (see relevant sections on this topic). Application of these
refinements has been shown to contribute to a survival rate of
about 94% for gastrointestinal procedures performed in obese
and diabetic mice (Baran and Johnson, 2012).
Opportunistic Infections in
Immunodeficient Mouse Models
• Background: As a side effect of genetic engineering of mouse

strains, many of these animals have mild-to-severe impairment
of aspects of the functional immune system. This can lead to
overwhelming bacterial infections in these animals, even when
housed in barrier-level conditions and when handled by per-
sonnel donning appropriate personal protective equipment.
Klebsiella oxytoca is a common opportunistic agent that has
been isolated from urogenital tract infections and abscesses
of mice, as well as being identified as the etiology for otitis,
keratoconjunctivitis, meningitis, lymphadenitis, and pneu-
monia (Bleich et al., 2008).
At times, the source of pathogens may come from experimental
agents administered to immunodeficient mice. For example,
animals that appear to rapidly lose body condition may be
described as “wasting.” Wasting disease, with septic arthri-
tis, has been associated with the injection of a Mycoplasma-
contaminated biological into severely immunodeficient mice
(Dodd et al., 2003).
• Potential treatments: In the modern facility, it is difficult
to maintain a consistently sterile and aseptic housing envi-
ronment, and it requires a great deal of oversight, labor, and
equipment. Autoclaving of all types of equipment and supplies
(caging, watering, and food) that come into contact with the
immunodeficient mice is of great benefit. In addition, biologi-
cal agents (to include differing cell types) should be screened,
prior to injection, for any opportunists or contaminants.
Common husbandry practices for immunodeficient trans-
genic mouse colonies (and for postirradiated animals that
have an altered immune system) include the provision of a
source of water acidified to less than 3.0 pH as an approach

to prevent bacterial contaminations of the water. Laboratory
mice provided with acidified water may consume less fluid
and have slower growth rates and may weigh less than age-
matched controls on untreated water (Craig et al., 1996).
Sterile autoclaved water may provide a reasonable alterna-
tive for a fluid source that does not necessarily expose mice to
bacteria other than normal flora (Styer et al., 2004). Husbandry
management would indicate that the acidified water should
be routinely monitored to ensure the pH is not so acidic that it
becomes unpalatable to mice. As well, nutritional supplemen-
tation in the form of sterile solidified gels may be provided (see
Chapter 4, “Nutritional Therapy Considerations”).
Radiation Exposure
• Background: Xenotransplantation, using immunodeficient

mouse models (e.g., NOD/SCID or SCID strains), is a key
experimental technique for the study of stem cell biology.
The immune system of animals is eliminated using radiation
(total body irradiation, TBI) and then the blood cell types may
be reconstituted with transplantation. TBI of 3 to 3.5 Gy is
used to minimize competition from endogenous bone m arrow
cells and ensure maximum engraftment of donor hemopoi-
etic cells. In the interim between radiation exposures and
acceptance of cell transplants, mice are extraordinarily vul-
nerable to acquiring infections and developing related disease
syndromes.
Irradiation doses greater than 10 Gy have been reported to
cause dental abnormalities in C57BL/6 mice; as well, during
a series of xenogeneic transplantation experiments, develop-
ment of brittle incisor teeth (Figure 2.35) was noted in NOD/
SCID mice at approximately 5 to 7 weeks after nonmyeloab-
lative TBI at 3 Gy (Larsen et al., 2006). This abnormality
was associated with rapid weight loss in the mice due to the
inability to prehend rodent chow.
Graft-versus-host disease has been described in immu-
nodeficient strains used for study of human tumor biology
and adoptive immunotherapy. Animals may present with
weight loss, scruffy hair coat, and hunched posture, with a
poor prognosis (Figure 2.36).
Fig. 2.35 Damage to incisors after nonmyeloablative total body irra-

diation may complicate NOD/SCID models of hemopoietic stem cell
transplantation. Teeth should be trimmed, and animals should have
softened food and nutritional support provided on the cage floor.
(Reprinted with permission from AALAS. Larsen, SR, Kingham, JA,
Hayward, MD, and Rasko, JE. 2006. Comp Med 56:209–214.)
Fig. 2.36 Following irradiation, mice may present in poor body

condition and be hunched and lethargic. This animal had been
gamma-irradiated 3 weeks previously and was provided with daily
supportive care, including nutritional supplements, fluids, and anti-
biotics. (Image courtesy of University of Pennsylvania, ULAR.)
• Potential treatments: Multiple aspects of husbandry care

(e.g., bioexclusion practices, health monitoring, water qual-
ity, use of antibiotics) are to be considered when housing
mice that are undergoing irradiation or bone marrow trans-
plantation. Common husbandry practices for postirradiated
animals include the provision of a source of water acidi-
fied to less than 3.0 pH as an approach to prevent bacte-
rial contamination. It cannot be overemphasized that the
first 7 to 10 days after transplantation are the most crucial;
close monitoring of the recipient mice by the laboratory and

husbandry staff is highly recommended to identify any pos-
sible health problems during this phase and beyond (Duran-
Struuck and Dysko, 2009).
Following irradiation, treatments often include administra-
tion of oral antibiotics and provision of softened feed (which
may be provided with high fat/calories for additional energy).
Feed and water consumption can be evaluated to gauge how
to use these vehicles for antibiotic delivery.
In one study, consumption of water from sipper tubes for
irradiated C57BL/6 mice (dosed at 67 cGy/min) was tracked;
it was found that consumption of acidified water dropped by
30% within 1 day of exposure and continued to decrease to
about 1.5 ml/day (from 4 ml/day) at 3 weeks. Overall, when
wetted chow was available, the intake of fluid directly from
sipper tubes was decreased by half. Consumption of acidified
water containing ciprofloxacin after irradiation was similar
whether grape flavoring or sugar was added for increased pal-
atability (Plett et al., 2008).
Fluid replacement with LRS (1–2 ml SC) immediately follow-
ing irradiation and twice daily for up to 7 days postirradiation
may be instrumental in minimizing loss of animals. LRS can
be combined with antibiotics (i.e., enrofloxacin at 85 mg/kg
SC twice daily) prior to injection. In contrast, trimethoprim-
sulfamethoxazole in the drinking water has not been shown
to be of overall benefit to mouse health following irradiation
(Ramirez et al., 2005).
Early recognition of any subsequent dental abnormalities
following TBI is beneficial. As well, trimming of teeth to allow
mandibular and maxillary incisor occlusion, coupled with
provision of softened chow and supportive care, can improve
the dental health of irradiated animals to maintain them
through experimental phases.
Humane endpoints should be established and may include
cage-side scoring of body postures, appearance, and activity
levels. It has been shown in C57BL/6 mice receiving an LD50
(median lethal dose) dose of 845 cGy that those a nimals achiev-
ing cage-side scores indicative of declining condition had cor-
responding mortality rates of 78–100%. The effort should be
made to preemptively e uthanize a nimals prior to spontane-
ous death (Nunamaker et al., 2012).
Following euthanasia of mice with graft-versus-host

disease, histologic assessments may reveal underly-
ing dermatitis, colitis and hepatitis, nephritis, arthritis,
meningoencephalitis, and vasculitis (Duran-Struuck et al.,
2010).
Streptozotocin Induction for Diabetic Models

• Background: Streptozotocin (STZ)-induced diabetes in
mice is often used to model diabetes mellitus and its com-
plications as well as other pathologies. In studies of diabe-
tes p
rogression and effects of newly developed treatments,
experimental results may be difficult to interpret because
blood glucose levels (BGLs) of untreated diabetic ICR mice
tend to decline substantially during typical experimental
time spans of 8 to 11 h. To address this problem, several
experimental conditions have been examined that might
affect BGL stability, including STZ dose, initial mouse
weight, fasting regimens and light–dark cycle within the
room. Interestingly, it has been shown that diabetes severity
is dependent on initial mouse weight, and that weight loss
after diabetes induction is less severe in heavier mice (Dekel
et al., 2009).
• Potential treatments: BGLs can be stabilized in diabetic
mice, particularly for those animals that are not undergo-
ing treatment with insulin, by regulating the amount of food
offered to mice during the experiment (Dekel et al., 2009).
Tumor Burden in Mouse Models

• Background: Rodent tumor models are extremely prevalent
in laboratory animal facilities. Tumor cell suspensions are
often implanted subcutaneously over the flank and scapular
areas to evaluate growth and immune responses to the trans-
planted cells; also, tumors can be transplanted following
growth in another animal (Workman et al., 2010). Tumors
can be induced by chemical carcinogens, by radiation, by
surgical anastomoses, and using viral and bacterial agents.
Spontaneous tumors can develop in certain strains, and
tumor burden should be managed based on how the animal’s
overall health and body condition fares (Figure 2.37). Note that
many rodents may show only subtle signs of clinical disease
A B C
Fig. 2.37 Varying degrees of tumor burden in laboratory mice:

(A) focal subcutaneous tumor induced in a nude mouse; (B) large
ulcerated tumor following induction in a nude mouse; (C) mass that
has encompassed the right forelimb to the extent that ambulation
is impaired (this animal would be recommended for euthanasia for
humane reasons). (Images courtesy of University of Pennsylvania,
ULAR.)
until late in tumor development; often, tumor incidence will

develop with increased age and other c omorbidities of aging.
Comprehensive listings of syngeneic, xenogeneic, and autoch-
thonous tumor models are available for review (Workman
et al., 2010).
Ear tag neoplasms have been documented in certain
strains of mice, including transgenic animals with a FVB/N
background. Related neoplasms have been identified as squa-
mous cell carcinomas (in 9% of mice > 300 days old) (Baron
et al., 2005) or fibrosarcomas (Everitt et al., 2002) in locations
closely associated with the presence of metal (nickel-copper
alloy) ear tags.
Lymphoma may present as a combination of clinical signs,
including abdominal or subcutaneous masses, anemia,
hunched posture, and poor body condition with ruffled fur.
When tumors are not present in subcutaneous locations,
scoring systems and schematics for identifying tumor bur-
dens should be established (Paster et al., 2009).
Intracardiac injection of human tumor cells into anesthe-
tized nude mice is an established model of bone metastasis.
However, intracardiac injection of some human tumor cell
lines causes acute neurologic signs and high mortality, mak-
ing some potentially relevant tumor cell lines unusable for
investigation.
• Potential treatments: In genetically modified mice, par-

ticular care measures are necessary to ensure detection
of unexpected sites of tumor development (Workman et al.,
2010). Individual institutional guidance should be followed
with respect to size of allowable tumors and increased moni-
toring of animal health (see Chapter 4, “Tumor Development
and Monitoring Considerations”). BW may not vary signifi-
cantly from baseline as tumor masses develop; therefore,
incorporating BCS assessments may be beneficial for moni-
toring any declines in health related to model development.
Other scoring criteria for intra-abdominal tumors could
include level of activity after unprovoked and provoked
stimulation, fur integrity, posture, breathing effort, abdomi-
nal and musculoskeletal palpation, and measurement of cir-
cumference of the abdomen and torso (Schenk et al., 2012).
Diagnostics may include biopsy of the mass for histologi-
cal analysis of cells and tissues, using fine-needle aspirates
(FNAs) and impression smears, and potentially ultrasonog-
raphy for masses in the reproductive tract, abdomen, and
mammary glands (Hochleithner and Hochleithner, 2004).
If tumors have ulcerated, meaning that the tumor has
outgrown its available blood supply and is now showing tis-
sue necrosis, this should be closely monitored and treated
with topical antibiotics and potentially analgesics. Certain
studies may warrant the tumor to become ulcerated to test
experimental treatments. However, most models that lead
to ulcerated tumor formation are typically close to experi-
mental endpoints; therefore, animals should not be kept
alive longer than is absolutely necessary to avoid welfare
concerns.
Studies have shown that intracardiac injection of tumor
cells can induce a hypercoagulable state leading to plate-
let consumption and thromboemboli formation, and that
pretreatment with intravenous injection of low-molecular-
weight heparin (LMWH; enoxaparin) blocks this state.
In addition, intravenous injection of enoxaparin before
intracardiac injection with two different small-cell lung
carcinoma lines, H1975 and H2126, dramatically decreased
mouse mortality while still generating bone metastases.
Therefore, reduction of mortality by pretreatment with
LMWH increased the types of cells that can be studied in
A B C
Fig. 2.38 Removal of auricular tumor secondary to ear tag placement

(A). Amputation of the pinna, using cordless cautery loop tips, was
performed to remove the mass (B). The mouse received triple antibi-
otic ointment applied topically twice daily for 7 days and meloxicam
(5 mg/kg SC) for 3 days postsurgery. The amputation site had healed
within 10 days (C). (Images courtesy of University of Pennsylvania,
ULAR.)
this metastasis model and decreased the number of a

nimals
used (Stocking et al., 2009).
Surgical excision provides a potential cure and is gener-
ally more effective than chemotherapy or radiation ther-
apy for those animals with experimentally induced tumors
(Figure 2.38); tumor removal results in more cures than all
other modalities combined (Mehler and Bennett, 2004).
Mammary neoplasia in mice is almost always malignant;
therefore, aggressive resection is recommended if it is not an
intended aspect of the experimental model. Unfortunately,
recurrence and metastasis of this tumor type is common
after surgical resection (Mehler and Bennett, 2004).
For mouse models of melanoma, alternative endpoints
have been proposed since the animals may have heavy tumor
loads in the absence of other clinical abnormalities (Narver,
2013).
Chemotherapeutics may be administered for treatment,
depending on tumor type. Chemotherapy-related fatigue
may be present and persistent for up to 4 weeks after treat-
ment, linked to toxicity of the agents administered (Ray et al.,
2008). Confirmation of the tumor type should be made by
histopathology of tumor tissue samples harvested at surgery

or at the humane or experimental endpoint.
Urogenital Disease in Mouse Models

• Background: Chronic estrogen exposure has been a ssociated
with decreased muscular contraction of the urinary bladder
and subsequent urinary retention. This can lead to suscep-
tibility of mice to ascending infections of the urinary tract,
potential for bacterial colonization of the bladder ( cystitis),
and urine scalding around the perineal area.
Uroliths are typically found more often in male mice, in
association with mouse urologic syndrome (MUS). Intact male
mice housed on wire flooring have significantly higher rates
of MUS than those animals housed directly on bedding; clini-
cal signs include urine staining around the prepuce, edema,
perineal ulcerative dermatitis and alopecia, and bladder
distention (Everitt et al., 1988).
Spontaneous struvite urolithiasis has been described
in estrogen-treated ovariectomized female nude mice with
cystitis induced by Staphylococcus intermedius. Concurrent
factors of a moist ulcerative dermatitis with associated bac-
terial infection, as well as predisposition to urine retention
and vesicouretal reflux, may result in ascending cystitis and
subsequent uroliths. Female nude mice, of normal body con-
dition, have been documented to present with superficial and
ulcerative skin lesions around the perineal area and cranial
to the tail base.
• Potential treatments: Cases of urine scalding in mice may
be treated topically with Neosporin ointment once daily or
application of Silvadene (similar to treatment for burns). Note
that nitrofurazone ointment is a known carcinogen; therefore,
it is not recommended for use in rodents (Langlois, 2004).
Administration of systemic antibiotics can alleviate cystitis
and potentially alleviate urolith burden (Gibbs et al., 2007).
A bedding change in mouse cages from corncob substrate to
sterilized paper bedding has been linked to a 60% decrease in
incidence of severe urogenital disease; this simple husbandry
practice change can subsequently lead to an increased
survival rate for study animals to reach their experimental
endpoints (Simmons et al., 2002).
euthanasia
Euthanasia is the process of inducing painless death in animals. To
the greatest extent possible, animals being euthanized should not
experience pain, fear, or other significant stress prior to their death.
Carbon dioxide (CO2) exposure or narcosis is a frequently used eutha-
nasia method in the laboratory for small animals due to its rapid
onset of action, safety, low cost, and ready availability. Exposure
times for carbon dioxide differ dramatically depending on the age
of the mouse to be euthanized; mice older than 21 days of age typi-
cally require 5 min of exposure (Pritchett et al., 2005). Investigations
into the potential advantages of premedicating or anesthetizing mice
prior to CO2 exposure have led to the conclusions that these ancillary
approaches do not diminish the behavioral effects of exposure to a
low flow rate (defined as displacement of 20% of the cage volume per
minute) of CO2 (Valentine et al., 2012).
Cervical dislocation as a rapid means of physically causing
death has been shown to potentially have unacceptably high rates
of failure (up to 21%) for mouse euthanasia (Carbone et al., 2012).
Injectable and inhalant methods may be preferable to physical means
unless individuals have received specific hands-on t raining. Further
discussion is provided in Chapter 4, “Euthanasia Considerations,”
and in the AVMA Guidelines for the Euthanasia of Animals (American
Veterinary Medical Association [AVMA], 2013).
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3
for laboratory rats
introduction
The laboratory rat continues to be broadly studied as a model s pecies
for investigating disease pathophysiology. Rats are second only to
laboratory mice in the number used for biomedical research; for-
tunately, due to their similarities, many treatment applications
described for mouse models can be extrapolated for administration
to rats. Three main advantages to using laboratory rats for experi-
mental purposes for studies are their comparatively larger size,
coupled with their 3-week gestation period and production of large
numbers of offspring. In addition, there are known scientific areas in
which the laboratory rat is more similar to humans than the mouse,
including the vascular system, the complexity of the rat brain for
study of cerebral disorders, and the enzymatic ability of the rat liver
to metabolize drugs. The continuing increase in rat genetics data
and the rat genome have led to centralization of this information;
these resources are highlighted further in Chapter 5. General infor-
mation on working with laboratory rats is best reviewed in the com-
panion text The Laboratory Rat (Sharp and Villano, 2012). Further
background information on strains, stocks, and genotypes can also
be obtained by visiting the originating v endor source websites, and
additional resources are highlighted in Chapter 5.
113
overall assessments
When assessing laboratory rats, as described for laboratory mice,
it will be essential to compile a thorough database of information
on health status, research project enrollment, and any potential
procedures or treatments already administered. Additional routine
aspects of any critical care “history” (see details in Chapter 1) should
include the background strain, gender, and age to gain the great-
est portfolio of information prior to finalizing differential diagnoses.
Further, any changes to the animal’s environmental and housing
parameters should be reviewed for contribution to the clinical signs.
These can include macroenvironmental influences of lighting, noise
and vibration, and temperature and humidity of the room; as well,
the microenvironment of the cage (diet, water source, housed singly
or with other rats, bedding substrate) is to be considered with respect
to maintenance of animal health.
general medical approaches to physical

examination and health assessments
If rats present in a critically poor state, as determined by the listing
of abnormal health conditions in Chapter 1 (see Table 1.2), it will be
essential to minimize stress and prioritize clinical interventions into
smaller diagnostic and treatment steps. A medical record informa-
tion template and sick animal reporting sheet are provided and can
be used for any rats noted to be in less-than-optimal health (see
Chapter 1, Figures 1.1 and 1.2). Typical values for biologic param-
eters in rats are presented in Table 3.1. The size of the average adult
laboratory rat can range widely depending on gender, with females
typically lighter weight than males (overall ranging from 250 to
greater than 500 g). Despite the increased body size when compared
to laboratory mice, there still are strong limitations on the ability to
precisely quantify temperature, heart rate, and respiration rate with-
out the use of telemetry or other special equipment in rats.
Physical Examination
Knowledge of the appearance of a laboratory rat in clinically normal
health will be key to ensure recognition of abnormal clinical signs
should they appear. Visual examination of the animal is the first
critical care management for laboratory rats 115
Table 3.1: Miscellaneous Parameters for the Laboratory R at

Parameter Value
Lifespan 2.5–4.0 years
Puberty 50 ± 10 days
Gestation 21–23 days
Male body weight 450–520 ga
Female body weight 250–400 ga
Blood volume 57–70 ml/kg = 17.1–21 ml total/300-g rat
Food intake 5–6 g/100 g BW/day
Water intake (ml/100 g BW/day) 10–12 ml/100 g BW/day
Packed cell volume (PCV) 35–57%
Glucose 80–300 (mg/dl)b
Body temperature (rectal) 35.9–37.5°C (96.9–99.5°F)
Respiratory rate 70–150 breaths per minute
Heart rate 250–600+ beats per minute
Source: Adapted from Banks, RE, Sharp, JM, Doss, SD, and Vanderford, DA.
2010. Rats, pp. 81–92. In Exotic Small Mammal Care and Husbandry.
Wiley-Blackwell, Ames, IA; and Sharp, PE, and LaRegina, MC. 1998. The
Laboratory Rat. CRC Press, Boca Raton, FL.
a Weights will vary depending on diet, age, stock, or strain.
b Enzyme values are dependent on collection method and may be influenced by
anesthesia.
step in assessing the overall physical condition of the laboratory rat

and should be done with the rat in its home cage or housing setup
prior to manual examination. Rats in critically poor health may ben-
efit from access to supplemental heat and increased oxygen (flow
rate 1–2 L/min) exposure (Klaphake, 2006).
As with standard handling practices for laboratory animals, to
prevent transmission of potential human pathogens and unwanted
exposure to animal allergens, fresh disposable gloves should be
donned prior to manual restraint and handling of laboratory rats.
Handling of rats throughout their time in the research environment
will assist with their acclimation to this interaction with personnel.
Data on a handling approach called “tickling” suggest that stress
associated with handling and intraperitoneal (IP) injections is mini-
mized using this technique (Cloutier et al., 2010). Playful handling
includes gentle manipulation and petting of the rats for a few min-
utes, both before and after a procedure, for the patient to develop
further acclimation to its human handlers.
Rats are difficult to lift by the scruff and will often vocalize and
struggle against this type of restraint. Instead, a firm grip on the
tail base of the rat will facilitate lifting the animal out of the cage
(Figure 3.1), followed by placement of the animal on a stable surface
A B
C D
Fig. 3.1 Retrieval of rat from cage can be conducted using a firm

grip at the tail base (A) to lift the animal and then placing it into the
crook of the staff member’s arm (B). Alternatively, the rat is placed on
a firm surface or benchtop and can be calmed by covering the eyes
with a towel, which also serves to protect the handler from a nimal
bites (C). The two-handed method of restraint allows for a partner
to administer a physical examination or treat with therapeutics
(D). (Images courtesy of University of Michigan, ULAM.)
(e.g., on a laboratory bench, flow hood surface, or in the crook of

the handler’s arm held adjacent to the torso). Gentle two-handed
restraint is preferred to best assess overall condition.
Handling allows for the ability to closely observe skin and hair
coat conditions, any ocular discharge or abnormalities, tooth over-
growth, abnormal masses, or unusual presentations in the anogeni-
tal region. Physiological aspects, like body weight (BW), activity, and
behavior assessments, are useful to measure and monitor serially.
Heart and lung sounds should be auscultated and can be performed
using a pediatric stethoscope. Hair coat quality should be reviewed
regarding location of areas of alopecia (baldness), open or closed
wounds, or poor grooming. In addition, respiratory status (difficult
or labored breathing with a more frequent/diminished rate than
expected) should be evaluated. Relative perfusion status, ascertained
by the color of mucous membranes (and potentially by color check of
ear and tail tissue), reflects the transport of fluid and oxygen in blood
to meet metabolic needs. Collectively, these physiologic measures
provide a crude assessment of the “ABCs” (airway/breathing/circu-

lation) of critical care medicine.
Rats in a critical state may need to be sedated to perform p hysical
assessments while minimizing stress responses. Gentle palpation of
the abdomen, using a pincer technique with the thumb and forefinger,
should help to confirm pregnancy in females and further identify
abnormalities like abdominal masses and growths, m ammary tissue
enlargement, or bladder distention. Finally, the particular experimen-
tal use of the rat, as described and approved in the approved p
roposal
to the IACUC (Institutional Animal Care and Use Committee), must be
considered, and any adverse effects of the experimental procedures
should be documented.
Body Condition Scoring

Assessing general body condition, as described in mice, remains an
excellent semiquantitative tool to apply toward rats for assessing
health status. The use of a body condition score (BCS) scale (gener-
ally on a range from 1, for wasted or emaciated, to 5, for obese) is
greatly enhanced by the incorporation of available cartoon diagrams
that represent each score on the scale (Hickman and Swan, 2010).
The uniformity of the diagrams can be exceptionally valuable for
assessments done by a laboratory animal group with variable levels
of experience in working with rats (Figure 3.2).
Overall percentages of weight loss should be tracked in rats yet
may or may not indicate a loss of health condition, depending on
the disease model and whether the animals are expected to develop
spontaneous or experimental tumors or other syndromes. Typically,
weight loss of more than 20–25% from preexperimental baseline may
warrant critical care measures and potentially euthanasia, depend-
ing on institutional policies.
Clinical Assessments of Ill Health and Pain in Rats

Rats, though predators of some animals, are considered as prey spe-
cies in the biomedical research environment. As such, similar to
mice, they are conditioned to suppress overt painful behaviors, par-
ticularly when being handled. The following are clinical assessments
of ill health and pain in rats (Kirsch et al., 2002, Kohn et al., 2007,
Miller and Richardson, 2011, Roughan and Flecknell, 2004):
• Vocalization, particularly when handled or a painful area is

palpated
BC 1
Rat is emaciated
• Segmentation of vertebral column prominent
if not visible.
• Little or no flesh cover over dorsal pelvis. Pins
prominent if not visible.
• Segmentation of caudal vertebrae prominent.
BC 2
Rat is under conditioned
• Segmentation of vertebral column prominent.
• Thin flesh cover over dorsal pelvis, little
subcutaneous fat. Pins easily palpable.
• Thin flesh cover over caudal vertebrae,
segmentation palpable with slight pressure.
BC 3
Rat is well-conditioned
• Segmentation of vertebral column easily
palpable.
• Moderate subcutaneous fat store over pelvis.
Pins easily palpable with slight pressure.
• Moderate fat store around tail base, caudal
vertebrae may be palpable but not segmented.
BC 4
Rat is overconditioned
• Segmentation of vertebral column palpable
with slight pressure.
• Thick subcutaneous fat store over dorsal
pelvis. Pins of pelvis palpable with firm
pressure.
• Thick fat store over tail base, caudal vertebrae
not palpable.
BC 5
Rat is obese
• Segmentation of vertebral column palpable
with firm pressure; may be a continuous
column.
• Thick subcutaneous fat store over dorsal
pelvis. Pins of pelvis not palpable with firm
pressure.
• Thick fat store over tail base, caudal vertebrae
not palpable.
Fig. 3.2 Schematic for scoring of the rat body condition. (Reprinted
with permission from AALAS. Hickman, DL, and Swan, M. 2010.
J Am Assoc Lab Anim Sci 49:155–159.)
• Reduced grooming or piloerection, leading to a “ruffled fur”

appearance
• Reduced level of spontaneous and exploratory (sniffing,
rearing) activity to the point that rats may not be moving

(“moribund condition”)
• Hunched posture with “guarding” of abdomen and reduced
mobility
• Squint-eyed appearance (either unilateral or bilateral)
• Increased aggressiveness on handling; may bite
• Porphyrin secretions (located around the eyes and nose); dis-
tinguish from bloody discharge by use of black light exposure
to the secretion type (porphyrin will “glow”; blood will not)
• Distanced from cage mates
• Reduced body condition, likely secondary to reduced nutri-
tional intake or experimental model resulting in muscle wast-
ing and weight loss
• Self-mutilation (excessive licking, biting, scratching) of the
painful area
• Abdominal writhing, increased back arching, falling or stag-
gering, poor gait, and twitching
• Palpation of unexpected masses
Monitoring Frequency
Similar to the laboratory mouse, a detailed and descriptive plan
for scheduled monitoring of rats both before and after any planned
experimental procedures, including the provision of therapeutic
treatments and supportive care, should be included in the IACUC
protocol submission. Investigators should be aware that as the poten-
tial for pain/distress in research animals rises, there should be an
increasing intensity of monitoring and frequency of observations.
Objective Scoring Systems

Professional and clinical judgments are essential for the evaluation
of an animal’s well-being and are critical to the ultimate decision
of euthanasia for humane reasons. As well, objective data-based
approaches to predicting imminent death, when developed for spe-
cific experimental models, should facilitate the implementation of
timely euthanasia before the onset of clinically overt signs of mori-

bund state (Toth and Gardiner, 2000). As described for mice, scoring
systems are one way in which rats can be monitored throughout an
experiment, and systems can be developed for individual experimen-
tal needs.
Novel approaches to pain assessment in laboratory rats have been
described based on coding of facial expressions, referred to as the
Rat Grimace Scale (RGS) (Langford et al., 2010, Sotocinal et al.,
2011). Rats are the most common animal model for preclinical pain
research (Mogil, 2009), and the RGS was used to improve quantifi-
cation of pain in three common algesiometric assays: intraplantar
instillation of complete Freund’s adjuvant, intra-articular kaolin/
carrageenan administration, and laparotomy. In contrast to the gri-
mace scale in mice, control rats display distinct bulging of the nose
and cheek regions; with pain, the bridge of the nose flattens and
elongates, further causing the whisker pads to flatten. This action
unit of “nose/cheek flattening” shows the highest correlation with the
presence of pain in the rat. The other action units, measured on the
0–2 scale, include orbital tightening and ear and whisker changes
(Figure 3.3).
Overall, quantifying pain by facial changes provides a practical
clinical assessment in that it can be performed in real time by trained
investigators, animal technicians, and veterinary staff (Sotocinal
et al., 2011).
veterinary care measures

Administration of Fluids
Dehydration is often present in rats that have pain or are unwell
and may be assessed by performing a skin tent or gentle pinch
of scruff over the scapulae of the rat and assessing the time that
passes for skin to return to normal placement. A prolonged return
time indicates a degree of dehydration that should be ameliorated.
Fluid administration through the subcutaneous (SC) route should
be the least-invasive way to provide supplemental fluid support to
the sick rat. Injections into the peritoneal cavity have been eval-
uated to determine how best to avoid accidental puncture of the
cecum, and it has been shown that by avoiding the left lower side
of the abdomen and injecting into the right lower side, the cecum
is not affected (Coria-Avila et al., 2007). Intravenous (IV) injections
Not present Moderate Obvious

“0” “1” “2”
Orbital tightening
Nose/cheek flattening
Fig. 3.3 Representative photographs of certain action units of the Rat

Grimace Scale for a rat at baseline (facial grimacing not present, 0;
a rat with moderate facial grimacing, 1; and a rat with obvious facial
grimacing, 2) (Reprinted with permission from Biomed central open
access. Sotocinal, SG, Sorge, RE, Zaloum, A, Tuttle, AH, Martin, LJ,
Wieskopf, JS, Mapplebeck, JC, Wei, P, Zhan, S, Zhang, S, McDougall,
JJ, King, OD, and Mogil, JS. 2011. Mol Pain 7:55.)
can be performed for rats using the femoral, jugular, or tail vein,
with animals appropriately sedated for access to the larger vessels
(Figure 3.4); incisions may be required to gain access to the vessel
of choice (Turner, Brabb, et al., 2011). Attempts at refinements for
smaller-volume dosing have identified the superficial penile vein of
the rat as an option for intravenous injections (Shapiro et al., 2010).
As a reminder, the beneficial effect of playful handling (tickling) for
rats is strongest when provided both immediately before and after
injection (Cloutier et al., 2010).
Water and fluid replacement sources are gaining in popularity,
expanding from products initially developed as sustainable fluid
sources for the duration of rodent shipping and transport. The provi-
sion of these water replacements, in disposable single-use containers,
is typically done on the cage floor for rapid access by those animals
in ill health. These supplementary fluid sources, when combined
Fig. 3.4 Fluids may be administered intravenously to anesthetized

rats using the lateral tail vein and a syringe infusion pump. (Reprinted
with permission from AALAS. Turner, PV, Pekow, C, Vasbinder, MA,
and Brabb, T. 2011. J Am Assoc Lab Anim Sci 50:614–627.)
with food, can maintain the health of rodents for several days in the
absence of routine water sources (Luo et al., 2003). Additional critical
care considerations for nutritional support, fluid administration, and
available products are provided in Chapter 4.
Blood Sampling
Blood sampling, or venipuncture, choices in rats may be influenced
by sampling site, anesthetic agent, and method of collection (Fitzner
Toft et al., 2006). Sampling allows for testing of serum chemistry
parameters, as well as complete blood counts. Suggested sampling
sites and further commentary are provided in Table 3.2.
As a guide, the volume of blood taken during a single survival col-
lection should be limited to that needed, not in excess of 10% total
blood volume (TBV) in rats; this also may be defined as a limit of
about 1.0 ml/100 g BW (Sharp and LaRegina, 1998). For example,
for 1% of BW to be withdrawn, 2.5 ml could be sampled at a single
time point from a 250-g rat. Following sampling of 1% BW volume,
replacement fluid therapy (0.5–1.0 ml SC or IP of sterile isotonic fluid)
should be provided.
Retro-orbital blood sampling may be performed with animals
under anesthesia but has been associated with subsequent lens
opacities and a higher outcome of clotted samples, as compared to
other methods (Mathieu, 2011). Other alternative sampling sites in
rats include the lateral saphenous vein (Figure 3.5), the sublingual
vein, and tail vessels.
Table 3.2: Recommended Sampling Sites and Related Information for

Blood Collection in R ats
Approximate Range
Anatomical Site Anesthesia? of Volume Collected Comments
Lateral Not required Up to 1% of BW
saphenous vein
Sublingual vein Not required 50–100 µl
Lateral tail vein Not required Up to 1% of BW
Tail clip Recommended Up to 1% of BW <2 mm of distal end of
tail should be clipped;
analgesia should be
considered
Jugular vein Recommended Up to 1% of BW
Submandibular Not required Up to 1% of BW
Retro-orbital Required Up to 1% of BW
vasculature
Cardiac Required 3+ ml Terminal procedure only
The sublingual vessel can be accessed with the animal unanes-

thetized and securely restrained, similar to a basic hold used for
performing an oral gavage. The mouth will open wide enough to

expose the sublingual vasculature, and the oral cavity should be
rinsed gently with saline or water and dried prior to sampling. Using
a 25- or 23-gauge needle, the vein is punctured, and blood is col-
lected via drip method into the appropriate collection tube. Gauze
can then be packed under the tongue to achieve hemostasis (Kohlert,
2012). Care must be taken with any method to ensure that struc-
tures surrounding the sampling site are not injured. As well, digital
pressure should be applied to achieve hemostasis following blood col-
lection. For critically ill animals, the tail sectioning method may be
used by making a transverse perpendicular incision at the tip of the
tail (Liu et al., 1996).
The submandibular technique initially performed in mice has
been adapted for use in rats. Rats should be lifted by the scruff
tightly behind the ears to include as much loose skin as possible;
alternatively, sedation will assist with the ability to limit mobility
for this procedure. Once lifted by the scruff, the insertion point for
the lancet should be located on the jawline, directly below the lateral
canthus of the eye. Lancet sizes vary, but a 5.5-mm lancet has been
used successfully in rats for this collection method. As with all sam-
pling approaches, hemostasis must be achieved following sampling,
typically through manual pressure over the lancet site of insertion
(Arzadon, 2011).
Fig. 3.5 Lateral saphenous vein sampling in the rat. The exterior

leg is shaved delicately with a scalpel blade, and lubricant is applied
over the vessel to allow blood to bead for collection (top); the vessel
is pierced with a medical lancet and blood allowed to pool over the
vessel (middle); the hematocrit tube is used to directly capture drops
of blood for later submission for serum chemistry and complete blood
counts (bottom).
Jugular venipuncture may also be utilized, and it has been per-

formed successfully in conscious and sedated animals using one-
handed restraint of the rat. If sedating the animal, the thorax should
be shaved to access the jugular vein and then swabbed with an alco-
hol pad. The collection needle should be inserted above the nipple
line at a 20° angle using a minute vacuum, until blood is drawn into
the collection tube. It is recommended to avoid continuous pressure
on the syringe plunger so the vessel will not collapse. Gauze should
be held in place over the blood draw site until hemostasis is achieved
(Zeleski et al., 2011).
Body Temperature Monitoring

Often, simple handling of ill rats will provide some indication of
whether they are excessively cool or warm to the touch, but body
temperature variations have to be extreme for manual detection.
Rectal thermometer probes require gentle placement and positioning
during procedures and may be more readily utilized in sedated rats,
given their larger body size compared to mice. Microchip transpon-
ders that provide identification as well as thermometry are also use-
ful for rodents (Bio Medic Data Systems, Seaford, DE).
Body temperature monitoring is critical for animals that are
scheduled to undergo prolonged anesthesia; the goal is to mitigate
hypothermia associated with experimental and surgical procedures.
Suggestions to ameliorate hypothermia would include incubators
and warm water bags, as well as Mylar-backed drapes, to reduce
radiant heat loss. In addition, warm water recirculation or forced-air
(Bair Hugger ®, Arizant Healthcare, Eden Prairie, MN) blankets may
be beneficial and synergistically effective when coupled with Mylar-
backed draping (Koch et al., 2008).
No matter the type of draping used, personnel should ensure that
draping allows for viewing of animals to ensure appropriate anes-
thetic administration and respiratory monitoring. Adverse incidents
involving unrecognized surgical fires occurring below the level of
a typical blue surgical drape have been described for the rat (Caro
et al., 2011). Drapes coupled with a heat-emitting gel pad (Figure 3.6)
can provide acceptable thermal support in the rat (Taylor, 2007).
Bone Marrow Access

Bone marrow aspiration from rats has been described as a method
to obtain antemortem cell samples. A minimally invasive approach
harvesting marrow from the femur (Figure 3.7) spares the knee joint
and serves to minimize potential damage to the musculature of the
quadriceps (Ordodi et al., 2006).
Endotracheal Intubation
Endotracheal intubation (Figure 3.8) can be readily performed in the
rat using either a method of blind access or a strong external light
Fig. 3.6 Reusable heating pad (SnuggleSafe® Microwave Heatpad,

West Sussex, UK) used for thermal support of anesthetized rats,
with manufacturer’s cover intact. (Reprinted with permission from
AALAS. Taylor, DK. 2007. J Am Assoc Lab Anim Sci 46:37–41.)
Fig. 3.7 Bone marrow harvesting in an anesthetized rat. In prepara-

tion for bone marrow harvesting, the rat should be anesthetized and
intubated; then, the thigh area should be shaved and disinfected for the
procedure (top). A 14-gauge needle and a 2-ml syringe are the required
instruments for harvest; the needle pierces the anterior face of the thigh
above the knee joint and is advanced into the femur prior to aspira-
tion of cell sample (bottom). (Reprinted by permission from Macmillan
Publishers Limited. Ordodi, VL, Mic, FA, Mic, AA, Tanasie, G, Ionac, M,
Sandesc, D, and Paunescu, V. 2006. Lab Anim (NY) 35:41–44.)
source that penetrates the skin to illuminate the larynx and facilitate
intubation. Other options for endotracheal tubing can be fashioned
from standard 2-ml syringes and a light source to illuminate the oro-
pharyngeal cavity, providing easy localization of the larynx (Molthen,
2006, Ordodi et al., 2005). If the rat is in respiratory distress, intuba-
tion should be undertaken with caution; however, it can be attempted
in animals weighing more than 100 g (Paul-Murphy, 1996). Note that
in the critically ill rat, intubation may be challenging and should be
A B
C D
Fig. 3.8 Intubation in the rat. (A) This photograph of a l aryngoscope

with a light source incident on the proximal end illustrates how
light is transmitted to the distal surfaces. (B) Image of laryngeal
opening of a rat showing the epiglottis (red a rrowhead), aryte-
noid cartilages (black arrows), and caudal margin of the soft pal-
ate (black arrowhead). Visual appearance is similar in the mouse.
(C) An anesthetized rat positioned and restrained on inclined
plane. (D) The laryngoscope is positioned in the oral cavity to pro-
vide visualization of the larynx. The tongue is grasped against the
shaft of the laryngoscope. The stylet and tracheal tube are shown
before being inserted into the oral cavity. Note the relative position
of stylet within the tracheal tube. (Reprinted with permission from
AALAS. Molthen, RC. 2006. J Am Assoc Lab Anim Sci 45:88–93;
and Rivera, B, Miller, S, Brown, E, and Price, R. 2005. Contemp Top
attempted only as a last resort to gain airway access if tracheostomy

cannot be performed (see relevant s ection in Chapter 4).
Injections and Oral Administration

Injections can be performed routinely using multiple routes (Table 3.3)
for the rat, including subcutaneous (SC), intradermal (ID), intraperi-
toneal (IP), intratracheal (IT), and intravenous (IV), as described in
Chapter 1. Intravascular access ports, typically placed surgically in
the subcutaneous space over the shoulder area for laboratory rats,
can be accessed using Huber needles for s ubstance administration
(Figure 3.9). The intramuscular route (IM) can also be more readily
used with small-volume injections, as compared to the mouse.
Table 3.3: Recommendations for Injection Dose Limits Based on

Weight of Laboratory R ats
Injection Limits (ml/kg)
Route PO SC IP IM IV (Bolusa) IV (Slow)
Dose (ml/kg) 10 5 10 0.1 5 20
Weight (kg)
0.200 2.0 mL 1.0 mL 2.0 mL 0.02 mL 1.0 mL 4.0 mL
0.225 2.2 1.1 2.2 0.02 1.12 4.5
0.250 2.5 1.2 2.5 0.02 1.25 5.0
0.275 2.7 1.3 2.7 0.02 1.35 5.5
0.300 3.0 1.5 3.0 0.03 1.5 6.0
0.325 3.2 1.6 3.2 0.03 1.6 6.5
0.350 3.5 1.7 3.5 0.03 1.75 7.0
0.375 3.7 1.8 3.7 0.03 1.85 7.5
0.400 4.0 2.0 4.0 0.04 2.0 8.0
0.425 4.2 2.1 4.2 0.04 2.1 8.5
0.450 4.5 2.2 4.5 0.04 2.25 9.0
0.475 4.7 2.3 4.7 0.04 2.35 9.5
0.500 5.0 2.5 5.0 0.05 2.50 10.0
0.525 5.2 2.6 5.2 0.05 2.6 10.5
0.550 5.5 2.7 5.5 0.05 2.75 11.0
0.575 5.7 2.8 5.7 0.05 2.85 11.5
0.600 6.0 3.0 6.0 0.06 3.0 12.0
0.625 6.2 3.1 6.2 0.06 3.1 12.5
0.650 6.5 3.2 6.5 0.06 3.25 13.0
0.675 6.7 3.3 6.7 0.06 3.35 13.5
0.700 7.0 3.5 7.0 0.07 3.50 14.0
a A bolus is a larger dose given over a shorter period of time.
Fig. 3.9 Access device for administration of substances in rats.

Vascular access port (top); noncoring Huber needle ( bottom).
(Reprinted with permission from AALAS. Turner, PV, Pekow,
C, Vasbinder, MA, and Brabb, T. 2011. J Am Assoc Lab Anim Sci
50:614–627.)
Administration of drugs and fluids can also be done by oral gavage;

however, animals must be healthy enough to drink from syringes.
This method of fluid/drug administration is less invasive and is
easily undertaken once rats are trained to the syringe (Figure 3.10)
(Atcha et al., 2010, Turner, Brabb, et al., 2011). If repeated oral (PO)
dosing is required, acclimation of rats to handling and to gavaging
(with up to 5 ml/kg of a control aqueous material) will help to dimin-
ish chronic stress from the procedures (Turner et al., 2012).
Creative approaches to disguising medications in palatable sub-
stances, like analgesics in gelatin desserts (Jell-O®), have been
described for rats (Flecknell, Orr, et al., 1999, Flecknell, Roughan,
et al., 1999). Attention should be paid to the differences in dosing
that may be required if delivering oral medications compared to sub-
cutaneous administrations; as well, rats may need to acclimate to
the novel substance prior to drug incorporation (Martin et al., 2001).
Urine Sampling
Clinically healthy rats will often dribble urine, which allows for a
free-catch sample (Klaphake, 2006). Slight pressure can be applied
Fig. 3.10 Oral dosing of laboratory rats. Rats readily drinking

galantamine (0.5 mg/kg) by the novel syringe-dosing method after
an acclimation training period (left); animal voluntarily consuming
nutritional supplement from a syringe (right) (Reprinted with permis-
sion from AALAS. Atcha, Z, Rourke, C, Neo, AH, Goh, CW, Lim, JS,
Aw, CC, Browne, ER, and Pemberton, DJ. 2010. J Am Assoc Lab Anim
Sci 49:335–343; and Turner, PV, Brabb, T, Pekow, C, and Vasbinder,
MA. 2011, J Am Assoc Lab Anim Sci 50:600–613.)
over the bladder to assist with expression of urine, and one should
ensure that an appropriate sterile receptacle is positioned to collect
the sample (Kurien et al., 2004). Critically ill rats should be stabi-
lized prior to attempting urinary catheterization if urine collection
by other methods has been unsuccessful. Urinary catheterization
should only be performed on anesthetized animals. Aseptic tech-
nique (see Chapter 4, “Perioperative Care Considerations”) and an
atraumatic approach should be used during placement of a urinary
catheter. Prior to insertion of the catheter, the external urinary orifice
should be gently cleansed using a disinfecting (e.g., chlorhexidine)
solution. The individual performing the catheterization is advised to
don sterile surgical gloves, use a sterile catheter, and apply a small
amount of sterile water-soluble lubricant on the external urinary ori-
fice. Additional sterile lubricant should be applied in a thin layer to
cover the surface of the urinary catheter for ease of insertion into
the urinary orifice, as described for mice (St. Claire et al., 1999). The
diameter of the urinary catheter should be the minimum that can
be inserted into the bladder and still prevent urinary leakage around
the catheter.
The anatomy of the female rat is unique in that the urinary orifice
is external and just anterior to the vaginal opening. Adult female
rats can be catheterized with number 50 polyethylene (PE) tubing
(2.9 French), a 3.5-French TomCat catheter, or a number 4 Coude

urethral catheter that has a bend to the tip of the catheter. This bend
facilitates passage of the catheter through the urethra. A guidewire
can be threaded through the PE tubing to increase the rigidity of the
catheter. Care should be taken that the tip of the guidewire does not
extend past the end of the catheter. Guidewires can be made of stain-
less steel surgical wire and coated with a water-soluble lubricant to
ease placement and removal from the PE tubing. The approximate
distance from the external urinary orifice to the neck of the bladder
for 200-g female rats is 17 mm (St. Claire et al., 1999).
If urine concentration tests are to be performed in rats, personnel
should be aware that dehydration may be secondary to any p rolonged
water deprivation necessary for this type of assay. Studies that
assessed clinical condition and BW, at a frequency of every 2 h begin-
ning 16 h after food and water deprivation were initiated, showed a
mean BW loss of 8% at 16 h and nearly 10% at 22 h. Clinical dehy-
dration was noted by 22 h, whereas appropriate urine concentration
was noted at 16 h. Therefore, it is recommended to complete the rat
urine concentration test within a 16-h period to maintain welfare of
the animals for this procedure (Kulick et al., 2005).
abnormal, critical, and emergent conditions

Categories of laboratory rodent health concerns are discussed in
alphabetical order to facilitate location by the reader. Under each
topic, “cause and impact” has been provided, and “potential treat-
ments” offer suggestions about procedures, therapeutic treatments,
or husbandry and environmental alterations. Every attempt has
been made to provide citations from the literature for evidence-based
medical outcomes.
It is essential to note that certain abnormal conditions
can be assessed and treated in laboratory rats by similar
methods to those done for laboratory mice; see relevant sec-
tions in Chapter 2 for the following:
• Abdominal swelling
• Abscessation
• Cage flooding
• Cannibalization
• Cross fostering
• Dystocia
• Fight wounds
• Fractures/orthopedic problems
• Hemorrhage
• Mortality (sudden death)
• Ocular lesions
• Respiratory distress
• Trauma
• Ulcerative dermatitis
For those health concerns that list drug therapy options, please
refer to the rodent formulary provided in Appendix C for additional
details on dosages and route of delivery.
Burns
• Cause and impact: Burns may be the unfortunate outcome
of improper surgical preparation of the animal with alcohol-
based disinfectants. Care should be taken to ensure that
animal skin that is prepped with alcohol is not then inadver-
tently ignited by cautery tools during surgery. Smoke inhala-
tion and superficial and partial-thickness burns have been
documented to result from this sort of accident (Figure 3.11),
the severity of which may be missed due to obstructive surgi-
cal draping (Caro et al., 2011). Burns may result in blistering
and skin necrosis, shock, and secondary bacterial infections.
• Potential treatments: Animals should be provided with oxy-
gen supplementation and stabilized following smoke inhala-
tion. The extent of burn damage should be assessed and any
wounds cleansed, debrided, and covered with a topical anti-
bacterial cream, like silver sulfadiazine. To prevent second-
ary bacterial infection, treatment with systemic antibiotics
should be considered. As well, warmed subcutaneous fluids
should be provided to offset shock and prevent dehydration.
Pain management should be a priority, with opioids or nonste-
roidal anti-inflammatory drugs (NSAIDs) provided through-
out the duration of the initial healing phases. The prognosis
will depend on the extent and thickness of the burned area;
aggressive management and monitoring are advised, and
euthanasia may be warranted.
Fig. 3.11 Surgical burn in a rat undergoing a procedure to emulate

rotator cuff injury. The animal was covered completely by the surgi-
cal drape, and the combination of alcohol applied to the surgical
site and cautery resulted in a surgical fire that singed the whiskers,
muzzle, and incision margins in the rat. Due to the degree of smoke
inhalation, this rat was ultimately euthanized. (Images courtesy of
Catheter Infections
• Cause and impact: Rats, more commonly than mice, are
catheterized using vascular access ports (Figure 3.9), typi-
cally into the jugular vein, for chronic administration of any
variety of test substances. Any indwelling catheter has the
potential to serve as a nidus of infection, leading to systemic
illness with clinical signs of decreased body condition and
activity and altered behavior. As well, localized inflammation
can occur at the skin surface and in the subcutaneous space
with development of pustular material and a threat to cath-
eter patency, particularly during chronic studies.
• Potential treatments: Frequent attention to catheter care
is the key prevention strategy against infections. Catheters
should be flushed at least twice weekly, once at the time of
treatment and again 3 days later. Skin overlying the vas-
cular access port can be cleansed with a chlorhexidine
scrub, alternated with dilute povidone–iodine solution.
Gentle manual restraint of the rats will permit access to the
port site; a noncoring Huber needle can then be inserted
through the skin and into the port reservoir. It is recom-
mended to flush the catheter with a volume of about 0.2 ml
saline, followed by 0.2 ml heparinized dextrose. Following
treatment administration, an additional 0.2 ml saline can
be injected to purge the catheter line. Catheters should be

“locked” with an anticoagulant to assist with patency; this
material can include heparinized dextrose or may involve
500 IU of heparinized glycerol (Wachtman et al., 2006,
Weiner et al., 2012).
Anecdotally, flushing the catheter with saline every 3 to 7
days has been evaluated and found to have negative impacts
on the ability to withdraw blood samples (Luo et al., 2004).
Should rats present with signs of systemic disease, anti-
biotics can be administered, along with supplemental fluids
and potential removal of the catheter from the animal. If the
rat is intended only for a study requiring chronic catheter-
ization, it may be best to euthanize the septic rat in lieu of
aggressive attempts at treatments.
Malocclusion (Incisors) and Caries

• Cause and impact: Incisor overgrowth may occur secondary
to congenital tooth patterns or may relate to lack of appropri-
ate caging materials for gnawing, softened foods, or a genetic
predisposition. Rats that have difficulty prehending food will
be anorectic, lose weight, and typically show a decreased BCS
within a relatively rapid (~24- to 48-h) time frame.
Dental caries may develop spontaneously in certain trans-
genic rat strains; animals should be monitored for signs of
anorexia and potential pain secondary to development of
caries (Nishijima et al., 2007).
• Potential treatments: Treatment includes trimming
overgrown teeth to a normal length and alignment. A dia-
mond blade, Dremel®, and dental burr are recommended
tools for use on rat dentition. Care should be taken not
to crack or split the teeth, which could potentially cause
pain and lead to tooth root infections. For valuable rats
with potential oral pain secondary to teeth abnormalities
and caries, anesthetic extractions of affected teeth may
be a potential treatment. Subsequent provision of softened
nutritional supplements and wetted chow may be necessary
to maintain body condition (see Chapter 4, “Nutritional
Therapy Considerations”).
Special attention should be given to the potential for mal-
occlusion in aged rats (especially noted in Wistar rats) during
long-term rodent studies as the increased incidence may be

detrimental to maintenance of health and general well-being
(Dontas et al., 2010).
Moribund/Weak/Paralyzed
• Cause and impact: Hind limb weakness (paresis) and
paralysis in laboratory rats are associated with trauma,

weakness, and dysfunction of the musculoskeletal and
nervous systems, neurologic disease models, adverse surgical
outcomes, or trauma that may be due to environmental or
experimental influence. Neoplasia and nonneoplastic dis-
eases, such as osteoarthritis, bone fractures, or peripheral
neuropathies, may also occur, particularly in aged rats
(Ceccarelli and Rozengurt, 2002).
Rat models of spinal cord injury are commonly implemented
for biomedical research; in addition to the induced spinal cord
lesions, the injured rats may experience alterations of the
liver, lung, bladder, and kidneys (Robinson et al., 2012).
Be aware that rats may also self-injure (autophagia or
autotomy) as a consequence of spinal cord or peripheral nerve
injury research, associated with altered mobility and pain
(Figure 3.12).
• Potential treatments: Rats found in a weakened and poten-
tially unresponsive state should be provided with ancillary
and supportive care of warmed subcutaneous fluids (2–4 ml
0.9% NaCl) and softened bedding substrates, nutritional sup-
plementation (including softened food pellets on the cage bot-
tom), and supplemental heat, until the level of responsiveness
is determined. It is critical to increase the frequency of moni-
toring and determine humane endpoints that eliminate pro-
longed suffering for paralyzed or moribund rats (see Chapter 4,
“Humane or ‘Clinical’ Endpoint Considerations”).
For those rat models of spinal cord injuries, research-
ers should be aware that suprapubic bladder catheteriza-
tions performed postinjury will not prevent development of
renal abnormalities in rats; therefore, manual expression of
the bladder should be performed two to three times daily to
eliminate urine accumulations (Robinson et al., 2012).
Increased observations and monitoring should be done for
those animals with self-inflicted lesions. If autophagia has
Fig. 3.12 Self-injury in an adult male Sprague-Dawley rat. Following

a left sciatic nerve transection and 48 h of postoperative analgesia,
this rat began to self-mutilate the left hind foot and toes (highlighted
in the enlarged image on the right) within 3 days, despite preventive
application of metronidazole/New Skin®. The animal was inhibited
from doing further damage by the application of a bitter-tasting
(Grannick’s Bitter Apple®) spray. (Images courtesy of University of
resulted in limb injury, then degree of lameness, amount

of swelling, and integrity of the wound should be assessed
(Geertsema and Lindsell, 2011). Treatments should include
analgesia, cleansing of the wound site, bandaging of
the area following application of local analgesics and t riple
antibiotic ointments, and placement of a restraint collar
to
prevent the rat from accessing the injured area (see
Chapter 4, “Restraint Collar Considerations”). Metronidazole
can be applied (“painted”) topically over the area where self-
injury has occurred due to its aversive taste; a NewSkin®
bandage can then be painted over the metronidazole to

prolong the presence of the drug on the skin and promote
healing (Zhang et al., 2001). If the self-injury is severe
to the point of severe welfare alterations, euthanasia is
recommended.
More often than not, moribund and paralyzed animals will
require euthanasia if there is no improvement or change in
activity status within 24 h of initial presentation.
Ocular Lesions Secondary to Anesthesia

• Cause and impact: Cloudiness of corneal tissue can be sec-
ondary to application of anesthesia and omission of topical
eye lubrication while animals are under anesthesia. Corneal
lesions and keratoconjunctivitis sicca (Kufoy et al., 1989) can
be more severe in animals anesthetized with ketamine plus
xylazine; minimal ocular changes have been noted in rats fol-
lowing enflurane or isoflurane anesthesia. Corneal lesions can
be observed within 24 h after injectable anesthetic administra-
tion and may be irreversible. Compared with Sprague-Dawley
and Lewis rats, Wistar, Long-Evans, and Fischer 344 rats had
increased incidence and severity of corneal lesions after anes-
thesia with ketamine plus xylazine, suggesting that these three
strains are at increased risk for developing postanesthetic cor-
neal lesions with this regimen (Turner and Albassam, 2005).
Acute reversible corneal lesions, attributable to a side effect
of xylazine, have been documented in rats (Calderone et al.,
1986).
• Potential treatments: Treatment with a sterile ophthalmic
lubricant (e.g., PuralubeTM) can assist with the prevention
of dry eyes and will soothe irritation. Ophthalmic products
of this nature should be applied any time a rodent is under
anesthesia to provide a protective film over the ocular surface,
similar to what is done in routine veterinary clinical practice.
The severity of corneal changes has been diminished in rats
for which ketamine plus xylazine anesthesia was reversed
with yohimbine (Turner and Albassam, 2005).
Topical ophthalmic ointment, with or without added antibiot-
ics, is warranted as a first-line approach even for animals that
appear to be otherwise behaviorally normal, despite the ocular
lesion. Certain lesions will be painful, with notation of animals
scratching at the eye and face; these animals should receive
topical (proparacaine) drops and systemic analgesics (meloxi-
cam 5 mg/kg SC) for pain relief. Surgical enucleation may be
warranted for correction of severe ocular injuries in rats.
Poor Body Condition

• Cause and impact: Rats may present clinically with a thin,
alopecic, and hunched appearance without much forewarn-
ing (Figure 3.13). This may be attributed to malocclusion (see
Fig. 3.13 Rats may present clinically with a thin, alopecic, and

hunched appearance. Animals should be treated with supportive
care, and further diagnostics are warranted. (Image courtesy of
relevant section on this topic) that is preventing ingestion of

hard food items (e.g., pelleted chow) or as a result of a gastro-
intestinal abnormality, husbandry alterations, experimental
manipulation, or internal tumor burden (Mexas et al., 2011).
Differentials should include behavioral stereotypies that
are preventing the animal from grooming or causing the ani-
mal to overgroom. Infectious agents should be ruled out as a
cause of alopecia through the performance of skin scrapes
and fungal cultures. If experimental treatments are poten-
tially toxic or unpalatable, this should be further discussed
with the research team.
• Potential treatments: The logical and prioritized causes
should be treated first, typically including administration
of a subcutaneous bolus of fluids, nutritional support, and
heat supplementation if the animal is hypothermic. If the
animals were expected to succumb to experimental disease,
then aggressive treatment efforts should be aimed at assist-
ing with nutritional supplements and comforting the animal
with provision of cage enrichments and bedding substrates to
attempt to reach the experimental endpoint.
The rest of the affiliated colony should be evaluated further
to determine if the condition is endemic, and blood sampling
can be done for further diagnostic assessments of poten-
tial infectious pathogens. Body condition scoring should be
monitored daily and BWs checked routinely to track any
further decline. Animals that become quiet, less alert, and
unresponsive should be considered for euthanasia prior to a

moribund state and spontaneous death.
Ringtail
• Cause and impact: Ringtail is a pathologic condition of
the tail, and sometimes feet, characterized by dry skin and
annular constrictions that can result in necrosis and loss of
portions of the tail (Figure 3.14).
The cause of ringtail is not completely understood, although
the condition is typically noted in weanling animals and may
be caused by relative environmental humidity levels below
25%. Other contributing factors, such as dietary deficien-
cies, genetic susceptibility, environmental temperatures, and
degree of hydration, also have been proposed. The variety of
possible etiologic factors suggests that this syndrome might
be the clinical expression of more than one causative agent
or that more than one causative agent may be necessary to
induce ringtail (Crippa et al., 2000).
• Potential treatments: Treatment with over-the-counter lan-
olin ointment (a nontoxic, inexpensive, and effective moistur-
izer) has been successful when initiated prior to the condition
becoming severe enough that there is tail necrosis. It can also
be applied prophylactically to rats starting at 7 days of age for
groups that may have a history of disease (Taylor et al., 2006).
Ulcerative Dermatitis
• Causes and impact: Ulcerative dermatitis (UD) has been
noted in Zucker lean rats, especially distal to forelimbs, with
isolated lesions on the head and behind the ears. Determining
the appropriate sensitivity profile to cultured bacteria is
essential to provide an effective antibiotic treatment. Skin
lesions may be secondary to dietary deficiencies, such as lin-
oleic acid deficiency, with manifestation of focal areas of alo-
pecia to diffuse areas of moist dermatitis on the head, face,
ear pinnae, and neckline.
• Potential treatments: Administration of leptin topically at
5 µg daily to affected areas can provide reduction in wound
size and severity. As well, trimming of hind toenails to pre-
vent self-inflicted skin trauma is advisable (Oppelt, 2005).
Fig. 3.14 Examples of ringtail in preweanling rats. (A) Normal,

healthy rat pup tail. This tail was given a condition score of 0. Note
that the distal portion of the tail has been biopsied for genotyping.
Representative clinical cases of ringtail of varying severity: (B) Rat
pup tail showing some flaking of the skin with mild constrictions.
This tail was given a condition score of 1. (C) Rat pup tail clearly
exhibiting annular constrictions and some malformation of tail
t issue. This tail was given a condition score of 2. (D) Rat pup tail
exhibiting annular constrictions with some malformation of tis-
sue; the tail tip appears necrotic. This tail was given a condi-
tion score of 3. Topical application of lanolin to tails appearing
like those in Figure 3.14B and 3.14C returned them to a healthy
and clinical normal tail appearance. Due to the level of necrosis
in Figure 3.14D, one might consider amputation of the tail tip to
remove necrotic tissue. (Reprinted with permission from AALAS.
Taylor, DK, Rogers, MM, and Hankenson, FC. 2006. J Am Assoc
Concurrent correction of dietary imbalances, topical applica-

tion of betadine cleanses, triple antibiotic ointment, and zinc
oxide may be beneficial (Godfrey et al., 2005).
For a comprehensive listing of various treatments of UD in
laboratory mice that may be efficacious for UD in laboratory
rats, refer to the relevant section in Chapter 2.
Urolithiasis
• Cause and impact: Clinical signs indicative of urolithiasis
include combinations of hematuria, red-stained bedding with
abnormal urine, red-stained or wet pelage (especially over the
abdomen), sensitivity to touch in the abdominal area, swollen
or palpable kidney or bladder, unkempt fur, anorexia, reduced
urination, reduced water intake, and unexpected weight loss
or gain (due to fluid retention) (Newland et al., 2005).
Partial-to-complete obstruction of urinary outflow can
cause mild-to-severe pressure necrosis of the renal pelvis,
medulla, and eventually the cortex. In addition, urinary cal-
culi can inflame and cause degeneration and necrosis of the
epithelial lining of the urinary tract. Incomplete emptying of
the urinary system due to obstruction, coupled with the loss
of epithelial integrity, allows bacterial overgrowth and subse-
quently an ascending urinary tract infection. In the case of a
severe infection, bacteria can gain access to systemic circula-
tion and cause sepsis.
Urolithiasis has also been linked to a model of lymphocytic
choriomeningitis virus (LCMV) infection in Lewis rats (Mook
et al., 2004).
• Potential treatments: These factors indicated above, when
taken as a whole, make it clear that once potentially obstruc-
tive uroliths form, the future health of the rat is at considerable
risk, perhaps irreversibly, because calculi are highly persistent.
Diet may need to be altered if the rats are to be maintained
in the research colony. For example, of those rats maintained
on a purified American Institute of Nutrition (AIN)-93 diet,
males are considerably more at risk for urolithiasis and
develop the condition within a few months of eating the diet
(Newland et al., 2005). As rats on the AIN-93 diet aged, the
discrepancy in risk between males and females increased; in
fact, by 100 weeks, nearly 60% of male rats died of urolithiasis,
Fig. 3.15 Representation of severe urolithiasis in an adult Lewis rat

experimentally infected with lymphocytic choriomeningitis virus.
Bladder walls were markedly thick, and the bladder was enlarged
and filled with multiple small uroliths. (Reprinted with permission
from AALAS. Mook, DM, Painter, JA, Pullium, JK, Ford, TR, Dillehay,
DL, and Pearce, BD. 2004. Comp Med 54:318–323.)
three times the prevalence seen in female rats. Postmortem

analyses suggested that males were more likely to have blad-
der calculi than were females, who usually formed calculi in
the kidney. Euthanasia of rats with severe clinical signs of
urinary dysfunction, likely secondary to stone formation, is
warranted to limit continued discomfort and potential spon-
taneous mortality (Figure 3.15).
research-related medical issues

Additional topics concerning laboratory rat health felt to warrant
further information herein due to their prevalence in contempo-
rary research environments are provided next in alphabetical order.
Under each topic, “background information” is provided, and “poten-
tial treatments” offer suggestions about procedures, therapeutic
treatments, and further considerations.
Arthritis Models
• Background: Induction of arthritis to better investigate
the pathogenesis of inflammation and test the potential
for antiarthritic agents is a classic model in the laboratory

rodent. Differing models include adjuvant arthritis (typically
in male Lewis rats, with injection at the tail base or into the
foot pad); type II collagen arthritis (typically in female rats
given bovine type II collagen); antigen arthritis; and injec-
tion of substances like capsaicin and carrageenan (Bendele,
2001). Tail and paw swelling with edema is expected as an
acute inflammatory reaction, and the severity of paw swell-
ing may render the animal immobile. While experimental
treatments can be administered, these rats would typically
be scientifically justified not to receive analgesics for pain
management due to concern for impact on the development of
experimental inflammation.
• Potential treatments: Provision of soft bedding and feeding
of a softened food or nutritional supplement on the cage floor
are recommended (Flecknell, 2001). Positioning of the water
source, such that the animal can access fluids easily without
additional pressure on the inflamed joints, should be consid-
ered. If analgesic treatments can be administered and not
compromise the data, one could provide NSAIDs (indometha-
cin), dexamethasone or other corticosteroids (at low doses to
avoid toxicity seen with chronic use), methotrexate (low dose),
and biological agents like soluble tumor necrosis factor R2
(TNF-R2) that are currently marked for human treatments of
arthritis (Bendele, 2001). It is not recommended that animals
be handled daily as this may increase their stress and alter
the desired inflammatory outcomes; instead, every-other-day
(EOD) handling should be sufficient (Brand, 2005). Humane
endpoints must be established for this type of model to best
limit the duration and intensity of pain sensation (National
Research Council [NRC], 2009).
Cranial Implant Maintenance

• Background: Neuroscience research often requires surgical
implantation of an apparatus that permits direct manipula-
tion of brain tissue or measurement of neuronal activity in
conscious animals. Successful factors for longevity of cra-
nial implants in rats have been described (Gardiner and
Toth, 1999). Contributing factors include accurate targeting
of the location of interest, aseptic surgical technique, maxi-
mal adherence of acrylic cement to the bone through proper
preparation of the skull surface, and provision of ventilation

during the thermogenic phase of cement curing. For the skin
to heal properly around the implant site, apposition of skin
to the implant is essential to promote comfort and reduce the
likelihood of secondary bacterial infections.
• Potential treatments: Wound margins should be treated
topically and liberally with antibiotic ointment (daily for
7 days postsurgery) (Gardiner and Toth, 1999). Repositioning
of the skin to adhere better to the headpiece may be of use
for a nonhealing incision site, and skin retraction may need
to be employed to gain the coverage needed over the skull-
cap. Systemic antibiotics can also be administered pre- and
postsurgically.
Continued maintenance of the areas around the implant
will likely be necessary to avoid the buildup of crusts and
potential for secondary bacterial infections (Figure 3.16).
Using a nonirritating antiseptic/cleansing solution, gently
remove any scabbing and minimize disruption of wound mar-
gins. Trimming of hairs along the margin is useful to mini-
mize irritation. Antibiotic ointment may then be applied as
needed. Cultures should be routinely taken to track potential
bacterial infections and provide relevant systemic antibiotics,
if necessary.
Fig. 3.16 Cranial implant complications in an adult rat. Porphyrin

staining was noted around the eyes and nares, indicative of poor
health and stress. Although the wound margins appeared relatively
healthy, apposition was compromised. Due to declining health, this
animal was euthanized, and Pseudomonas aeruginosa was cultured
from the brainstem surface (arrow). (Images courtesy of University of
Incontinence Secondary to Spinal Cord Injury Models

• Background: Urine scald secondary to urinary incontinence
from spinal cord injury studies can pose significant clinical
problems. Urine scald is likely to cause discomfort, signified
by skin with severe redness and warmth and the presence
of urine or urine stains. Related complications can include
intractable skin ulceration, secondary bacterial dermatitis,
and self-trauma.
• Potential treatments: Amelioration of the discomfort has
been attempted by application of a commercially available
hexamethyldisiloxane (HMDS)-based skin protectant barrier
film, 3M No-Sting Barrier Film® (3M Corporation, St. Paul,
MN), which is used to treat diaper rash in human infants and
urine scald in incontinent adults (St. Claire et al., 1997).
For incontinent rats, urinary bladders can be manually
expressed every 8 h until rats are observed to urinate with-
out assistance. After each timed expression of the urinary
bladder, barrier film is applied to clean, dry skin by spray-
ing a uniform coat of film over the affected area. Animals
should be observed for signs of discomfort after application
of the barrier film. In addition, daily monitoring for paralysis,
signs of dehydration, food intake, and evidence and degree of
urine scald should be performed. Skin irritation can be rated
from minor (slight redness, cool to touch) to major (severe red-
ness, warm to touch) with or without moisture from urine
(St. Claire et al., 1997).
Treatment of spinal cord injury with minocycline, an anti-
biotic with neuroprotective effects, has been beneficial for
restoration of motor coordination and hind limb reflexes.
This antibiotic can be administered within 1 h after injury
(90 mg/kg IP), followed by doses (45 mg/kg IP) twice daily for
5 days (Teng et al., 2004).
Middle Cerebral Artery Occlusion in Rat Models of Stroke

• Background: The major complication of the stroke model is
the substantial morbidity and mortality that occurs postop-
eratively due to respiratory distress caused by stimulation
of the sympathetic nerve system. Prolonged occlusion of the
common carotid arteries can lead to prolonged tachycardia
and potential for arrhythmias.
• Potential treatments: Administration of bupivacaine (0.25%

solution 0.1–0.2 ml SC) can ameliorate left-side heart failure
that would otherwise lead to mortality and may improve the
outcome of the model (Wang Fischer et al., 2003). Scoring
sheets have been developed for evaluation of pain manage-
ment in this model as the administration of NSAIDs for pain
relief may not be permitted due to bias of data outcomes
(Kirsch et al., 2002).
Obese and Diabetic Rat Models

• Background: Rat models for obesity and diabetes research
are beneficial to the identification of surgical and therapeutic
interventions that can be translated to the related human
disease syndromes. Performing surgery in physiologically
compromised rats, particularly with their body conformation
of a thinner thoracic cavity and larger abdominal mass, can
result in adverse effects like hypoglycemia, difficulty with
dehiscence of incision sites, and anesthesia reactions that
can potentially lead to fatal outcomes.
• Potential treatments: For those rats undergoing surgery,
attention to the minimal time for fasting, both presurgi-
cally and postsurgically, is key (see Chapter 4, “Fasting
Considerations”). To prevent dehiscence of surgical sites, it
is recommended to avoid surgical skin clips for this model
and instead close incisions with continuous suture patterns,
with minimal suture size, and subcuticular closure patterns.
Maintaining the anesthetic dose of isoflurane to no more
than 1.5%, with an oxygen flow rate of 0.5 L/min, facilitates
recovery issues and maintains animals at a reasonable depth
of anesthesia. As well, supplemental heat should be provided
as described and endorsed for all surgical models in rodents.
Application of these refinements has been shown to contrib-
ute to a survival rate of approximately 90% for gastrointesti-
nal procedures performed in obese and diabetic rats (Baran
et al., 2011).
Providing supportive care and specialized environments
will be best for these obese models. For animals with evi-
dence of diabetes, more frequent cage changes and provision
of more absorbent bedding substrates should be used in the
rat cages to compensate for increased urine production.
Opportunistic Infections in Immunodeficient Rat Models

• Background: Similar to cases in laboratory mice that are
immunodeficient (see relevant section in Chapter 2), Klebsiella
oxytoca has been identified as a monoculture from urogenital
tract infections and abscesses, as well as serving as the etiol-
ogy for otitis, keratoconjunctivitis, meningitis, lymphadenitis,
and pneumonia (Bleich et al., 2008). Abnormal colonization
with K. pneumonia has also been documented following anti-
biotic treatment in nude rats (Hansen, 1995). Rats enrolled
in longevity studies may succumb to opportunistic infections
with age, prior to collection of desired data points.
• Potential treatments: Husbandry and environmental
changes have been useful in eradicating opportunists, like
Pneumocystis carini. Housing rats in autoclaved cages, with
autoclaved bedding, and provision of trimethorim-sulfa-
methoxazole-treated acidified water have minimized reported
heath issues. As well, providing a diet with 14% protein and
3.5% fat, along with pair housing of rats, has effectively
extended the lifespan and improved overall health in aged
rats (Zahorsky-Reeves et al., 2007). Also, nutritional supple-
mentation in the form of sterile solidified gels may be provided
(see Chapter 4, “Nutritional Therapy Considerations”).
Pododermatitis
• Background: Pododermatitis can be common in mature rats
(>300 g) chronically housed (>1 year) in wire-bottom cages but
is less commonly noted when animals are housed on bedding
(Carraway and Witt, 2003, Peace et al., 2001). The problem is
characterized by chronic, suppurative inflammatory lesions
(ulcers) on the plantar surfaces of the hind feet; lesions may be
reddened and raised, with keratinized growth developing into
crusts and scabs (Peace et al., 2001, Sharp and Villano, 2012).
• Potential treatments: Topical and systemic treatment
options may be limited by impacts on study data; however,
antibiotics and analgesics would be ideal for addressing the
infectious nature and associated pain from these lesions.
Placement of some sort of flattened and softer bedding sub-
strate or surface on the wire cage bottom, akin to sterile gauze
squares (4 x 4 inches), has a significant preventive benefit for
diminishing the potential for ulceration of noted foot sores
(Dimeo and Mitchell, 2005, Peace et al., 2001). Soaking of

affected feet (hydrotherapy) in Epsom salt solution (4 cups
of warm water to 1 teaspoon of salt) has anecdotally been
successful for resolution of surface infection and softening of
crusts covering foot wounds. Surgical debridement is rarely
successful, and prognosis for complete resolution is guarded
(Langlois, 2004). Closely monitor affected animals to ensure
that any rats experiencing severe pain and distress are
removed from the study and euthanized.
Spontaneously Hypertensive Rat Models

• Background: To promote maintenance of BW of senescent
female spontaneously hypertensive rats (SHRs), supplement-
ing powdered feed is useful to offset loss of appetite and
weight loss.
• Potential treatments: With age, SHR rats will benefit from
the addition of powdered food to ensure that BWs remain sta-
ble and to prevent malnutrition that could lead to premature
death. Rats were also given powdered rat chow in shallow
bowls to facilitate the eating and digestion of food. As the
female SHR matures, special care and handling are essen-
tial to help maintain BW and good health. With only modest
changes in routine (i.e., powdered food) and an attentive eye
on the rats’ daily activities, it is possible to maintain these
rats in a healthy condition until the termination of the study
(Belanger et al., 1999).
Tumor Burden in Rat Models

• Background: Rodent tumor models are quite common in
laboratory animal facilities. Institutional guidance should
be followed with respect to size of allowable tumors and
increased monitoring of animal health (see Chapter 4, “Tumor
Development and Monitoring Considerations”). Spontaneously
occurring tumors may also develop and should be managed
based on how the animal’s overall health and body condition
fares.
Tumors in rats may be secondary to foreign body reactions,
particularly for intra-abdominal telemetry devices in certain
strains (Popovic et al., 2004). Tumor incidence should be con-

sidered an adverse outcome in instrumented rats.
Subcutaneous masses involving the mammary chain are
usually benign fibroadenomas, with less than 10% being
malignant. Mammary tissue in rats is extensive, and masses
can occur anywhere from the neck to the inguinal region,
arising in locations as dorsal as the flank areas and across
the shoulders.
Paraneoplastic syndrome in young rats has been described
secondary to extensive mammary neoplasia (Figure 3.17)
(Mexas et al., 2011).
Fig. 3.17 Induced neoplasia model young (2-month-old) female rat.

(Top) The animal was very thin with a BCS on initial examination
at 1.5 of 5. (Bottom) Visible and firm palpable masses extended
bilaterally throughout regions of mammary tissue on the ventral
aspect of the mouse (highlighted in boxes). The animal had a 48-h
history of lethargy and dehydration; on physical examination,
the rat became extremely stressed, developed agonal breathing,
and was euthanized immediately. Necropsy identified widespread
mammary tumors (corresponding with palpated masses) and

multiple organ abnormalities, including calcification as a paraneo-
plastic syndrome. (Images courtesy of University of Pennsylvania,
ULAR.)
Fig. 3.18 Rat with a spontaneous ulcerated mammary tumor. Due

to the ulceration and location in the left axillary region (left, ven-
tral view; right, right-side recumbent view), this rat would require
heightened monitoring for alterations to mobility, hemorrhage of the
mass, and further irritation to the tumor site. (Images courtesy of
• Potential treatments: Surgical excision is the most com-

mon form of therapy and results in more cures than all other
modalities combined (Mehler et al., 2004). Mammary gland
tumor removal can be straightforward; in brief, the vascu-
lar supply to these tumors is limited and therefore can be
ligated using vascular clamps or suture material. Once the
neoplastic tissue is removed, the tissue space and subcutane-
ous tissue can be closed with 3–0 vicryl suture (using a sim-
ple-interrupted or continuous pattern). Overlying skin can be
closed with suture, wound clips, or tissue glue (Fisher, 2002).
It may be possible to have the tumor treated in some other
manner to continue using the animal in a study; however,
overall animal welfare should not be compromised if the tumor
is left untreated, affects mobility, or ulcerates (Figure 3.18).
Tumor development may affect animal welfare for those ani-
mals in long-term studies and decrease confidence in the
reliability of data outcomes from the model.
euthanasia
Euthanasia is the process of inducing painless death in animals. To
the greatest extent possible, animals being euthanized should not
experience pain, fear, or other significant stress prior to their death.
Carbon dioxide (CO2) exposure or narcosis is a frequently used

euthanasia method for small laboratory animals due to its rapid onset
of action, safety, low cost, and ready availability. Exposure times for
carbon dioxide differ dramatically depending on the age of the rat
to be euthanized; rats older than 21 days typically require 5 min
of exposure time (Pritchett-Corning, 2009). Injectable and inhalant
methods are therefore preferred unless individuals have received
hands-on training for physical methods of euthanasia. Further dis-
cussion is provided in Chapter 4, “Euthanasia Considerations” and
in the AVMA Guidelines for the Euthanasia of Animals (American
Veterinary Medical Association [AVMA], 2013).
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4
special considerations for
in laboratory rodents
introduction
Critical care monitoring for small rodent patients is notoriously
challenging, complicated further by physiologic idiosyncrasies and
limitations on applicable treatments and interventions (Hawkins
and Graham, 2007, Lichtenberger, 2007, Lichtenberger and Ko,
2007). This chapter offers important supplementary information
for critical care for the laboratory rodent patient, with specifics on
types of m odels with inherent potential for pain and distress. The
comprehensive subject matter herein was consolidated from a variety
of institutional guidance documents, publications, and abstracts to
highlight p articular regulatory, clinical, and experimental facets of
working with these species. Due to the diversity of material covered
here, the t opics are in alphabetical order, akin to the listings of con-
siderations in p revious chapters, for ease of location.
In addition, readers are strongly encouraged to review their own
institutional guidelines and policies on these topics to best address
animal welfare and critical care management issues, ideally in con-
sultation with available veterinary staff.
1
aging animal model considerations

For biogerontology research, animals with median life spans longer
than 20 months are usually acceptable to achieve experimental
objectives (Nadon, 2004). The most commonly used strain for aging
research in mice is the C57BL/6, and the most commonly used rat
strain is the F344. Aging is often accompanied by dysfunction or
disease of bodily systems, manifesting as cardiovascular system
disorders, renal and respiratory function decline, osteoarthritis,
osteoporosis, cataracts, disease-promoting mutations, amyloidosis,
leukemias, and other cancers (Dontas et al., 2010, Nadon, 2004).
In aging mice, enlargement of preputial glands is common, particu-
larly bilaterally. However, these mice can still be used in research
(potentially excluding reproductive studies) because duct ectasia is
painless and does not commonly require treatment (Donnelly and
Walberg, 2011). An increased incidence of malocclusion has been
noted in aging Wistar rats; therefore, attention to dental health, and
provision of hard substances on which to gnaw, should be heightened
in aging colonies (Dontas et al., 2010, Nadon, 2004). Older rodents
may develop arthritis, which may lead to chronic pain; this can be
treated with a course of anti-inflammatory medications (2–3 weeks),
followed by a 1-week break, then resumption of analgesic treatment
(Flecknell, 2001). Neoplasia is also more common in aged laboratory
rodents, and specific strain idiosyncrasies have been reviewed else-
where (Danneman et al., 2012, Nadon, 2004).
Although many of these disease syndromes are progressive, rarely
are they life threatening; these aged animals can serve as unique
models in which to evaluate age-related physiologic alterations.
Unfortunately, the ability to detect subtle alterations due to aging
and subclinical declines in health is challenging despite daily obser-
vations of laboratory rodents. Similar to objective scoring systems for
monitoring health during experimental procedures, these types of
scoring systems can also be utilized for evaluation of aging animals
(Figure 4.1) (Phillips et al., 2010). Efforts have been undertaken to
determine markers of imminent death in aging mice by looking at
temperature and body weight changes over time (Ray et al., 2010,
Trammell et al., 2012).
Drug therapy in aging animals should be evaluated with some
caution. Older (geriatric) rodents may likely have varying degrees of
organ dysfunction and altered drug metabolism. Hepatic and renal
dysfunction can have a negative impact on attempts at drug therapy
special considerations for critical care management 3
Date: May 28, 2009 Rat ID C49

Notes: Teeth need checking Weekly Experiment 300
Date of birth 2–27–0 7
Age 27 mo
Possible score Actual score

Appearance: observe rat in cage
Appearance is normal: no obvious skin or coat problems 1
Slightly abnormal: slight change in coat or skin—could be dirty coat, dander, or inflammation 2
Moderately abnormal: obvious change from normal—more dullness, dirty, skin inflammation 3 3
Severely abnormal: marked change from normal—skin and coat very dull, dirty, or inflamed 4
Extremely abnormal: extreme signs of deterioration of coat and skin 5
Posture: observe rat while still in the cage or walking

Normal posture: sitting, standing, or rearing normally 1
Slightly abnormal posture: slight flatness or hunch 2
Moderately abnormal: possibly dragging belly or hunched 3 3
Severely abnormal: mostly flat with little elevation off surface 4
Extremely abnormal: flat to surface, unable to elevate off floor of cage 5
Moblity: observe rat’s ability to move around the cage

No impairment: able to move normally 1
Slightly impaired: may have some ataxia or splay causing slight problems with movement 2
Moderately impaired: obvious mobility problems 3
Severely impaired: definite problems with being able to move easily about the cage 4 4
Extremely impaired: unable to move at all or without extreme hardship 5
Muscle Tone: hold rat in hand to assess tone of rear legs and abdomen
Normal muscle tone: muscle groups have normal tone or mass 1
Slightly abnormal: muscle mass slightly soft 2
Moderately abnormal: muscle mass less firm, abdomen slightly soft 3 3
Severely abnormal: muscle mass very thin, soft, undefined 4
Extremely abnormal: muscle mass has no tone or definition 5
Total Score 4–20 13
Current weight 350.0

Previous weight 352.0
Weight gain (+) or loss (–) –2.0
Fig. 4.1 Sample score sheet for observational assessment, with

brief descriptions of the four measures, for aged rats. For each
measure, the highest (most abnormal) score possible is 5. The

total c
omposite score possible for all measures combined is 20.
In
addition, body weight for the current and previous weeks is
included, along with whether a weight gain or loss was present.
(Reprinted with permission from AALAS. Phillips, PM, Jarema,
KA, Kurtz, DM, and MacPhail, RC. 2010. J Am Assoc Lab Anim Sci
49:792–799.)
because the liver and kidneys are involved in the function, b

reakdown,
and clearance of most drugs. Dosing of certain m edications may
need to be lowered and administered with less frequency in aged
patients. Renal disease can diminish the kidney’s ability to filter and
excrete drugs, leading to the prolonged presence of drug and drug
metabolite in the body system. Therefore, drug types like aminogly-
cosides (e.g., gentamicin, amikacin), which are primarily removed
by the kidney, should be used with caution. As well, administration
of potassium-containing fluids could lead to life-threatening serum
potassium levels in animals with renal disease.
If aged animals are overweight or obese, one should consider
dosing at their “ideal” adult body weight, especially when using

drugs that distribute to adipose tissues, such as thiobarbiturates. In
general, animals with an abnormal body condition may have under-
lying medical problems that could affect drug choices; every effort
should be made to identify the cause of the altered body condition
before starting drug treatment.
blood loss considerations

Blood loss, secondary to hemorrhage, hemophilia, immune disorders,
or experimental sampling, may lead to physiologic alterations that
result in a critical health concern. An animal with chronic a nemia
may tolerate a loss of 60% of blood volume before a transfusion is
necessary. Assessment of the patient is key to best determine if the
animal is tolerating the anemia as blood transfusion has inherent
risks (Morrisey, 2003b).
Telemetric monitoring has been used during repeated blood draw-
ing to observe systemic effects of blood sampling/loss on mice.
In these studies, measurement of electrocardiogram (ECG), body
temperature, and blood pressure were performed while imitating a
blood sampling loss of up to 40% of the animals’ total blood volume
(TBV). Results noted on the ECG included declines in heart rates
after 30–40% of the TBV was withdrawn. In general, transfusions
are warranted for acute blood losses of greater than 30% of an ani-
mal’s blood volume or, alternatively, if there is a 50% decrease from
a baseline packed cell volume level (Morrisey, 2003b). In a related
study, mean blood pressure declined after 20–30% withdrawal of
TBV. Body temperature showed declines after 20% blood withdrawal;
therefore, these quantitative assessments support policies on limit-
ing blood drawing to 20% of TBV (Mercogliano et al., 2002).
Whole blood can potentially be transfused as needed but should

be transfused from like species to like species. Blood typing does not
appear to be necessary for most small mammals, and transfusion
reactions are rare. Premedication with diphenhydramine at 1 mg/kg
(intravenously [IV] or intramuscularly [IM]) may be beneficial to limit
adverse transfusion reactions (Mader, 2002).
chronic indwelling device considerations

Catheters and vascular access ports are implanted routinely in
laboratory animal patients being treated for cardiovascular diseases.
Management of device-associated infections is critical; development of
secondary complications can pose a serious health risk, particularly
if the animals are immunodeficient. In animals, infections associated
with long-term catheterization not only are potentially d etrimental to
their well-being but also are likely to cause variation in the experi-
mental data obtained from such animals.
Animals should be treated perioperatively with antibiotics when
long-term catheters are placed, potentially for a period of 5 to 7 days.
Diagnosis of infections, should they develop, may be based on clinical
signs of infection, including purulent discharge from the catheter
exit site, swelling or erythema at the exit site or along the subcutane-
ous catheter route, or abscess at any point along the catheter tract.
Temperatures may be elevated in these animals, with a decrease in
appetite and activity and high white blood cell counts on blood tests.
Specimens from suspect infections should be submitted for bacterial
culture and antibiotic sensitivity testing.
Localized treatment may be performed with removal of purulent
material from the catheter tract and disinfection/flushing of the tract
with a povidone-iodine containing solution, similar to what is done
in other species (Taylor and Grady, 1998). If health concerns increase
(continued fever, anorexia, lethargy), with failure to respond to topi-
cal disinfection and systemic antibiotics, the decision to remove the
catheter should be made in the interest of animal welfare.
depilatory cream considerations

Chemical depilatory creams are often used for fur removal, prior to
experimental procedures or surgery, in laboratory rodents. It may
be desirable to remove hair over blood collection sites (saphenous
vessels), ECG lead-positioning points, for imaging purposes, prior to

tumor cell implantation, or for surgical field access. Depilatory creams
for rodents work well, but creams should be completely wiped away
with a cloth and warm sterile solutions to prevent inadvertent inges-
tion during grooming and potential for skin irritation/inflammation,
which can confound research outcomes.
Animals should remain under anesthesia during the contact time
with the depilatory. Depilatories include over-the-counter products
like Nair ® (with or without aloe for sensitive skin) and Veet®. They can
be applied directly to the fur of the rodent or applied after clipping
the fur. The product may be left on for several minutes (1–10 min)
then wiped off with a sterile gauze pad or tissue; if necessary, the
application process can be repeated until hair removal is complete
(Angel et al., 1992, Finlay et al., 2012). No animal should be left unat-
tended during the application of depilatory agents.
Manual hair removal is not advised as an alternative; instead,
electric shavers or handheld razors should be employed with the
a nimal under anesthesia.
equipment considerations for rodent

surgery and emergency procedures
Prior to the presentation of critical care rodent cases, it is advisable
to prepare an emergency kit and supply area that is stocked, main-
tained, and readily accessible. Having designated locations assigned
for equipment and kits, containing supplies that are replaced on a
periodic basis, will prevent loss of time looking for relevant materials
during an emergent situation (Bergdall and Green, 2004, Mader,
2002, Paul-Murphy, 1996).
Items to consider maintaining for rodent procedures and surgery
include the following:
• Gram scale with disposable weigh boats: For easy collec-

tion of body weights of individual rodents to then accurately
calculate anesthetic and analgesic doses.
• Surgical tables: Design should be easily sanitizable and
appropriate for routine rodent surgeries, which can be per-
formed in dedicated areas outside a formal surgical suite;
enhanced design features for a table could include adjustable
height for the surgeon and allowance for tilting.
• Heating sources: Warm water-recirculating heat b lankets,

warm-air devices like the Bair-Hugger ® blanket and
ThermoCare® isolators, microwaveable gels, pads, and pock-
ets may also be of use for rodents (please see Chapter 1 for
further details on supplemental heat provision). Electrical
heating pads should not be used due to the potential for dermal
burns caused by uneven heating.
• Anesthesia machines: Equipment should include a vapor-
izer, oxygen tank and holder (or some connection to a central
O2 source with flowmeter), and CO2 absorber. Vaporizers need
to be specific to the type of volatile anesthetic used. Machines
designed to permit simultaneous anesthesia and monitoring
of multiple rodents are available.
• Anesthesia ventilators: For procedures like imaging or
prolonged surgeries, mechanical ventilation is typically nec-
essary. Ventilators are generally pneumatic (with bellows);
however, electronic ventilators are available for rodents.
• Anesthesia monitoring: To assist with determining the
overall ventilation status of the patient, consider investment
in ECG monitors for visual and auditory outputs, pulse oxim-
eters, and capnographs for measurement of the expired CO2
concentration (Stanford, 2004).
• Blood pressure monitors: These provide information about
the hemodynamic status of the patient and are especially
useful for cardiovascular surgeries.
• Cautery: Cautery applies an electric current to cut through
tissues and seal vessels as a means of hemostasis; harmonic
scalpels may be an alternative, using ultrasound to cut
and coagulate tissues simultaneously. For rodents, cordless
cautery loop tips (e.g., MediChoice®) are extremely useful for
t issue amputations and hemostasis.
euthanasia considerations
Euthanasia is described by the American Veterinary Medical
Association (AVMA) as a method of killing that minimizes pain, dis-
tress, and anxiety experienced by the animal prior to the loss of
consciousness, and causes rapid loss of consciousness followed by
cardiac or respiratory arrest and, ultimately, a loss of brain func-
tion (AVMA, 2013). Unfortunately, e
uthanasia is often the elected
outcome for many critically ill laboratory rodents due to irreversibil-

ity of disease and request for diagnostic necropsy. To alleviate pro-
longed suffering, interventional euthanasia is always preferable to
spontaneous death of laboratory animals. The AVMA provides eutha-
nasia guidance for the entire veterinary profession; however, the lab-
oratory animal community is expected to adhere to this published
guidance and additional references (AVMA, 2013, Danneman et al.,
2012, Muir et al., 1989, National Research Council, 2011, Sharp and
Villano, 2012).
Timeliness of humane endpoints involving euthanasia will need
to be outlined for critical cases, in particular if research data are
dependent on the preservation of the animal tissues. It is recom-
mended that staff achieve a consensus for when rodents should
be euthanized and in what time frame after reporting on the criti-
cal health condition (immediate, within 2 h, within 24 h, etc.). The
selected euthanasia method should minimize sources of potential
distress (excessive handling, disruption of compatible housing
groups, etc.). Methods may be deemed as acceptable (considered to
reliably meet the requirements of euthanasia), as acceptable with
conditions (considered to reliably meet the requirements of euthana-
sia when specified conditions are met), or as unacceptable (does not
meet the requirements of euthanasia) (AVMA, 2013). Assurance that
animals are to be euthanized properly must be documented in the
animal use protocol approved by the Institutional Animal Care and
Use Committee (IACUC).
General considerations for selection of euthanasia method for
laboratory rodents include the following:
1. Size, weight, age
2. Need for and type of physical restraint
3. Time required to produce loss of consciousness and death
4. Reliability and irreversibility
5. Skill level of personnel
6. Availability of facilities, equipment, and drugs
Acceptable methods of euthanasia for laboratory rodents include
injection of barbiturates (typically intraperitoneally at a dosage of
three times what would be used for anesthesia), injection of bar-
biturates in combination with local anesthetics and anticonvul-
sants, and injection of lethal doses of dissociative agents (such as
ketamine) in combination with drugs like xylazine or diazepam
(AVMA, 2013).
Methods that are acceptable with conditions include the use of

inhaled anesthetics (isoflurane, sevoflurane, desflurane, with or
without nitrous oxide in combination); inhaled CO2 (with or with-
out inhaled anesthetics; provided at a flow rate to displace 10–30%
of the chamber or cage volume per minute); and inhaled carbon
monoxide (not commonly used in the laboratory animal setting).
Note that euthanasia chambers prefilled with carbon dioxide are
unacceptable. Noninhaled agents like tribromoethanol and etha-
nol are acceptable with conditions. Physical methods (e.g., cervical
dislocation for animals < 200 g BW, decapitation, and focused
beam microwave irradiation) are also acceptable with conditions
(AVMA, 2013).
Unacceptable methods include the exposure of conscious mam-
mals to inhaled nitrogen or argon; animals would have to be under
heavy sedation or anesthesia for these agents to be allowed, and
other euthanasia methods are preferable. Noninhaled agents that
are unacceptable as a sole agent of euthanasia would include potas-
sium chloride, neuromuscular blocking agents, opioid overdoses,
urethane, or α-chloralose. There may be circumstances when these
agents, in combination with other methods, may be permissible
(AVMA, 2013).
Commonly, CO2 overdose is utilized for euthanasia of labora-
tory mice and rats. Because there may be a possibility that rodents
can recover from a deeply anesthetized state following exposure to
CO2, there is a need to ensure a secondary (confirmatory) means of
euthanasia. Further guidance is provided by the Office of Laboratory
Animal Welfare (OLAW), through the Public Health Service arm of the
federal government (see relevant section in Chapter 5 “Resources and
Additional Information”). These confirmatory methods, which should
be performed after CO2 overdose, might include one of the following:
exsanguination, decapitation, cervical disarticulation, bilateral tho-
racotomy, or at least 50% additional time in the euthanasia chamber
filled with 100% CO2 (IACUC-UPENN, 2008).
A recommended method for euthanasia of a pregnant dam is
CO2 exposure followed by cervical disarticulation. If fetuses are not
required for study, the method chosen for euthanasia of a pregnant
female should ensure cerebral anoxia to the fetus and minimally
disturb the uterine milieu to minimize fetal arousal (Klaunberg et al.,
2004). Euthanasia of neonates requires greatly prolonged exposure
to inhalant anesthetics, including CO2 (Table 4.1). Resistance to
hypoxia results in a prolonged time to unconsciousness if inhalant
agents are used. Therefore, it is an acceptable method for euthanasia
Table 4.1: Recommended Euthanasia Times for Laboratory Mice

and R ats
Minimum Time in 100% CO2 (minutes)
Age Mice Rats
Nonhaired pups: 0–6 days 60 min 40 min
Haired pups, eyes closed: 7–13 days 20 min 20 min
Haired pups, eyes open, preweaning: 14–20 days 10 min 10 min
Weanlings and adults: 21+ days 5 min 10 min
Source: Data from Pritchett-Corning, KR. 2009. J Am Assoc Lab Anim Sci
48:23–27; and Pritchett, K, Corrow, D, Stockwell, J, and Smith, A.
2005. Comp Med 55:275–281.
of neonates to administer sufficient quantities of injectable chemi-

cal anesthetics (American College of Laboratory Animal Medicine
[ACLAM], 2005, AVMA, 2013). Special cases for euthanasia are fur-
ther described in the AVMA guidelines (AVMA, 2013).
experimental autoimmune encephalomyelitis

and demyelinating disease model considerations
Rodent models of experimental autoimmune encephalomyelitis (EAE)
and related demyelinating diseases may result in a complex spectrum
of acute, chronic, and relapsing-remitting disease courses that most
often result in varying degrees of progressive ascending paralysis.
Due to the extreme variability in the onset and progression of clinical
signs and disease course, close monitoring, assessment of body con-
dition score (BCS), and provision of supportive care are necessary for
EAE animals (IACUC-UPENN, 2013).
EAE Scoring
Clinical signs and ascending paralysis in EAE are commonly
assessed on a six-stage scale of 0–5, with 0 representing a clinically
normal condition and 5 representing paralysis of all limbs (quad-
riplegia). Other scoring systems may be preferable and should be
clearly defined in the protocol and made available to animal care
staff in close proximity to the animal housing room.
0: Clinically normal
1: Decreased tail tone or weak tail only
2: Hind limb weakness (paraparesis)
3: Hind limb paralysis (paraplegia) or urinary incontinence

4: Weakness of front limbs with paraparesis or paraplegia
(quadriparesis) or atonic bladder
5: Paralysis of all limbs (quadriplegia)
Animal Care
Verification that research personnel are properly trained in the pro-
cedures related to these disease models must be documented in the
IACUC protocol. It is preferable to keep a written record of the disease
progression with information including the start date of experiments,
the BW and overall condition, and the general appearance per the
EAE scoring scale presented in the preceding section. Enrichment
with cotton nesting material is not recommended for animals that
will develop weakness and paralysis as the fibers may entrap and
strangulate weakened limbs/tails. It is recommended that a soft
bedding substrate (versus corncob bedding) be utilized to minimize
skin trauma secondary to paralysis.
When clinical signs are expected to begin, laboratory staff should
monitor mice at least once daily. The following guidance is designed
to assist with increasing monitoring and measures of care:
• Score 1–2: Separate affected animals to another cage to avoid

injury by unaffected animals. Alternatively, house with simi-
larly affected animals (preferred) to maintain social housing
environment. If available, provide water bottles with elon-
gated sipper tubes, even if housed in autowatering cages.
Provide nutritional (pelleted chow, special diets) and fluid gel
supplements on the cage floor and replenish as needed or at
least two or three times each week. Animal weights and BCS
should be determined at least twice weekly.
Note: If the BCS is 2 or less or if the animal has lost over 10%
of baseline weight, administer sterile fluid (1 ml SC) daily.
• Score 3–4: Treatments of daily fluid support and nutritional
and gel supplements should continue. Continued weight
collection and BCS should be performed twice weekly.
Once hind limb paralysis or urinary incontinence is noted,
increased monitoring and assistance with bladder expres-
sion will be expected. Gentle palpation of the bladder should
be done daily to express urine; gently press on the caudal
abdomen to assist with urination. If the bladder loses tone
(atony), express twice daily. Due to the potential for urine

dribbling onto skin, it will be critical to assess the animal for
ventral and perineal dermatitis, urine scald, penile prolapse,
and tail lesions. If these clinical abnormalities appear, the
animal will require further treatment, made in consultation
with the veterinary staff, to minimize urine scald.
• Score 5: Once an animal becomes quadriplegic (paralyzed in
all four limbs), it should be euthanized promptly unless the
approved IACUC protocol states otherwise. Other criteria for
euthanasia include a loss of more than 20% BW, a BCS of 1,
or moribund state.
fasting considerations
Small mammals require an almost-continuous supply of food and
water; accordingly, fasting (withholding of food for a designated
period prior to testing, then return of food) or restricting (limiting
ration of food provided) should be minimized to the extent necessary
to achieve the scientific objectives while maintaining animal well-
being. It is notable that many research protocols will request a time
period for the fasting of laboratory mice prior to procedures; fasting
by this definition means that the animals will be allowed free access
to water but that food may be removed prior to a planned procedure.
Fasting may occur for a number of reasons, including minimizing
the variability of drug exposure time prior to necropsy and reducing
the contents of the gastrointestinal tract prior to intraperitoneal
injection, intragastric dosing, or gastrointestinal surgery. In toxicol-
ogy laboratories, food may be withheld from rodents prior to necropsy
to improve the ease of handling and fixation of the gastrointestinal
tract and to yield more uniform liver histology sections. Withholding
of food is nonphysiological and may compromise ease of collection of
biological samples and overall animal condition. This practice also
may contribute unnecessary stress to experimental animals (Turner
et al., 2001).
In general, presurgical (~16 h or longer, also known as “overnight”)
fasting is not recommended for mice and rats, particularly due to the
increase in food generally ingested by rodents at the beginning of the
dark cycle. As well, water should not be withheld prior to anesthesia
(Lester et al., 2012). Rodents do not vomit; therefore, the rationale for
fasting to prevent aspiration pneumonia is not pertinent to mice and
rats. As well, the high metabolic rate can lead to hypoglycemia and
liver changes if rodents are fasted for any length of time (Morrisey,
2003a). Certain institutions may have policies that prevent any fast-
ing of animals prior to surgery unless the feeding condition is a key
aspect to the experimental model (Toth and Gardiner, 2000). In gen-
eral, it is recommended that mice and rats be fasted for no more than
2 to 4 h prior to procedures that require an emptied stomach (Lester
et al., 2012).
The provision of a palatable, simple carbohydrate to rats over-
night, in the form of sucrose (sugar cubes), reduces the size of the
gastrointestinal tract while minimizing other side effects of food
withholding, such as alterations in serum biochemistry parameters
and body weights (Levine and Saltzman, 1998). Offering sugar cubes
represents an inexpensive, simple, and readily available alterna-
tive to overnight fasting. However, the overnight feeding of sucrose,
in lieu of limiting chow or complete fasting, can result in marked
changes in gastrointestinal tract weight and pancreatic and hepatic
structure and function, as described for laboratory rats (Turner
et al., 2001).
The experimental rationale for fasting of rodents is typically
related to behavioral motivation and assessments. Animals may
experience some discomfort during longer fasting periods, and the
IACUC would require scientific justification for a particular duration
of deprivation balanced against the induction of potential distress
or physiologic harm (Rowland, 2007). Restriction studies normally
are preformed on healthy animals; thus, the physiologic conse-
quences differ from those of anorexia caused by illness. A healthy
animal that has lost 15% of body weight by restriction is likely
to acclimate and become clinically stable, whereas one that has
lost the same weight due to illness is typically not stable (Rowland,
2007). Overall, rodents can acclimate to fasting for experimental
purposes, specifically by efficiently reducing further fluid or energy
losses through a combination of innate behavioral and physiologic
adjustments.
fluid therapy considerations

Administration of warmed (~37°C) fluids is a frontline interven-
tion to treat acutely or chronically ill animals with various dis-
eases, including electrolyte disorders, acid-base disorders, and
hypovolemia induced by blood loss. Administering subcutaneous
fluids in critically ill rodents is the most common, efficacious, and
least-invasive method of delivery (DiBartola, 2000, Hawkins and

Graham, 2007, Klaphake, 2006, Mader, 2002). Dehydration (a loss
of body fluid from intracellular, plasma, interstitial, or transcellular
compartments) is the key indicator for fluid therapy. Dehydration
may be noted in a nimals that are in poor body condition, those
that have disease, or those that have undergone prolonged surgery
performed under anesthesia. Prompt and appropriate fluid ther-
apy should be instituted prior to or at the time that dehydration is
documented.
Fluid requirements for dehydration (Hawkins and Graham, 2007)
are calculated as follows:
% Dehydration × kg × 1000 ml/L = Fluid deficit (L)
Published dosages for fluid therapy in rodents range from 30 to

90 ml/kg in the first hour; further, maintenance rates for critically
ill rodents have been calculated at 100 ml/kg/day SC or IP with
compensation for special losses (Mader, 2002, Paul-Murphy, 1996).
Crystalloids
Crystalloids are most often used for rodent fluid therapy and are
defined as isotonic solutions (with plasma) that contain both elec-
trolytes and nonelectrolytes and are capable of entering all of the
body fluid compartments. Crystalloids are equally as effective at
increasing blood volume as colloid fluids but must be administered
in greater amounts since they are absorbed within all fluid compart-
ments. Crystalloids can be classified as replacement fluids (if they
are similar to the extracellular fluid) or as maintenance solutions (if
they contain less sodium and more potassium).
• Ringer’s solution with (LRS) or without lactate. Ringer’s solution

contains sodium, chloride, potassium, and calcium. LRS is
useful in increasing tissue perfusion and in extracellular
blood expansion. In addition, the lactate counteracts meta-
bolic acidosis that may occur with kidney failure or acute
fluid loss. There may be unwanted complications if LRS,
which contains calcium, is administered concurrently with
other drugs (Hackett and Lehman, 2005).
• Normosol®-R. Normosol-R is a buffered solution similar to
LRS; it contains acetate and gluconate instead of lactate.
• Saline. Normal saline (0.9% NaCl) is an isotonic solution that

contains normal sodium concentrations but greater chloride
concentrations than body fluids. Saline is used for acute
extracellular volume expansion.
• Dextrose. Dextrose at 5% (diluted in either NaCl or LRS) pro-
vides about 200 kcal/L and may satisfy the maintenance
energy requirement for rodents.
Colloids
Colloids are much less commonly used in laboratory rodents and
are defined as fluids that contain large macromolecules that are
restricted to the plasma compartment and cannot enter any of the
body’s fluid compartments. Colloids should be used in patients
with shock and hypoalbuminemia, particularly if an intravenous or
intraosseous access route has been established, to achieve volume
expansion rapidly. Colloids include natural (e.g., plasma or whole
blood) or synthetic (dextran, hydroxyethyl starch [Hetastarch],
and stroma-free hemoglobin [Oxyglobin®]) formulations. There are
noted limitations with colloid fluid therapy, including anticoagula-
tion activity; little published evidence of their effectiveness in rodent
patients exists.
food and fluid regulation procedures

Food and fluid regulation (whether scheduling or restriction) for
rodents is typically used in the research setting for three main areas
of study: as a means to motivate animals to perform novel or learned
tasks; to analyze the motivated behaviors and physiologic mediators
of hunger and thirst; and to investigate homeostatic regulation of
energy metabolism or food balance. Scientific justification should be
provided for using food and fluid regulation, and the least regulation
necessary to achieve the scientific objective should be used.
Experiments that involve food or fluid regulation should evaluate
the following factors: the level of regulation (meaning how limited the
access will be to food and fluid), potential adverse consequences of
regulation, and methods for assessing the health and well-being of
animals undergoing regulation (National Research Council, 2011).
Investigators should provide to the IACUC plans for the implementa-
tion of sufficient and scheduled monitoring (routine weighing and
target weight) of animals during food and fluid regulation studies.
Rodents should be acclimated over time (a minimum of 3 days)

to food and fluid regulation paradigms. Consideration should be
made to allow food and water to be available concurrently as rodents
typically do not eat caloric requirements without available water.
Regulated levels of food should not be lower than 30% of ad libitum
values. Overall, experiments involving food and fluid regulation are
not recommended in rodents less than 14 weeks of age.
At times, the degree of restriction may actually be better described
as deprivation for up to several hours. Studies have shown that
fluid deprivation (with ad libitum food access) in mice for 12, 24,
and 48 h results in average weight losses of 9%, 12%, and 18% of
initial body weight, respectively. These animals (relative to nonde-
prived controls) have decreases in plasma volume and increases
in plasma renin activity and corticosterone. For mice chronically
restricted over the course of 7 days to a 50–75% water ration, mod-
est dehydration anorexia (food intake reduction of ~10%) has been
noted, with severe renal lesions identified at necropsy. Acute restric-
tion of fluid over 24 h is associated with significant physiologic
stress and is not recommended due to welfare impact (Bekkevold
et al., 2012).
Animals on regulation should be closely monitored and weighed
at least weekly and weighed more often if animals are undergoing
greater restrictions (National Research Council, 2011). Weight and
BCS should be compared to age- and strain-matched control animals.
For animals undergoing scheduling or regulation, written records
should be maintained daily to document food and fluid consump-
tion, hydration status, and any behavioral or clinical changes used
as c
riteria for temporary or permanent removal of an animal from a
protocol (National Research Council, 2011). It will be useful to have
these written records readily available for the animal care staff,
IACUC members, or any outside reviewers. Designated personnel
should document certain experimental and clinical information,
such as the following:
• Date (daily documentation is necessary for animals undergo-

ing regulation)
• Baseline weight (prior to initiation of restriction)
• Weight and BCS (twice weekly)
• Indication of schedule
• Indication that access to food and water was granted (daily)
• Overall health, behavior, and activity (daily)
Specific humane, experimental, and interventional endpoints

must be clearly stated in the IACUC protocol. For food regulation in
rodents, the animal should not lose more than 20% of control weight
or baseline BW (if adult) matched by age, strain, and sex unless
scientifically justified to do so. Veterinary staff should be involved in
evaluations of animals that have lost 20% or more of baseline weight.
As well, it is not recommended that a food- or fluid-regulated animal
have a BCS of 2 or less if the BCS was higher than this level at the
start of the regulation protocol. For fluid regulation, animals with a
weight loss of 10% from baseline weight should be considered clini-
cally dehydrated and be treated as outlined in the discussion that
follows. Any rodent appearing dehydrated (displaying listlessness or
inactivity, with an increased “skin tent” time) should have a measured
volume of fluid provided promptly; care should be taken not to fluid
overload an animal that has been acclimated to restriction. In addi-
tion, up to 2 ml of fluid can be administered subcutaneously to boost
hydration s tatus and improve animal well-being. Research person-
nel involved in water restriction studies should be trained appropri-
ately to identify dehydration and correctly a dminister s
ubcutaneous
fluids.
Aspects of this section were adapted from institutional documents
(IACUC-UPENN, 2011a).
humane or “clinical” endpoint considerations

According to the National Research Council guide, experiments
that may result in “severe or chronic pain or significant altera-
tions in the animals’ ability to maintain normal physiology, or
adequately respond to stressors, should include descriptions of
appropriate humane endpoints” (National Research Council, 2011,
p. 5). Humane interventions or endpoints are defined as the point
at which pain or distress in an experimental animal is prevented
(with therapeutic interventions), terminated (by cessation of partic-
ipation in the study), or relieved (t ypically by euthanasia). Selection
of appropriate interventions provides significant opportunities
for refinements; as well, these endpoints must be relevant to the
selected model. For example, tumorigenicity studies could be ter-
minated as soon as progressive tumor growth is documented; how-
ever, carcinogen-induced papillomas may require later endpoints
so that cells are able to transform into a malignant state (Workman
et al., 2010).
It is extremely useful to itemize and describe specific humane

endpoints in IACUC protocols, particularly for those procedures
that may involve potential pain and distress. Humane i nterventions
and endpoints should be determined by a collaborative effort of
research staff with laboratory animal veterinarians. Humane end-
points for a variety of models are further reviewed in an issue of
the Institute for Laboratory Animal Research Journal (“Humane
Endpoints,” 2000).
Studies that commonly require special consideration for endpoints
may include the following:
• Tumor development
• Infectious disease
• Vaccine challenge
• Pain and trauma modeling
• Monoclonal antibody production
• Assessment of toxicological effects
• Organ or systemic failure
• Models of cardiovascular shock
• Demyelinating diseases
• Generation of animals with abnormal phenotypes
To develop a humane endpoint, one should be aware of the clini-

cal progression that a particular (group of) animal(s) is likely to
experience as a result of experimental manipulation or any spon-
taneously occurring disease that might develop during the lifetime
of the animal(s). Personnel must be adequately trained in the basic
principles of laboratory animal science (National Research Council,
2011); in this case, staff need to be able to recognize species-specific
behaviors, as well as signs of pain, distress, and morbidity.
The selection of appropriate humane endpoints requires a
detailed knowledge of the impact of the procedure on the ani-
mal to allow for intervention before unpredicted distress or pain
develops. If the outcome of a particular experimental model is not
known, which may be the case if studies are novel or alternative
endpoint information is lacking, pilot studies can be an effective
method for identifying and defining humane endpoints and reach-
ing c
onsensus among the investigators, the IACUC, and veteri-
nary staff (National Research Council, 2011). Pilot studies can be
designed to assess both the procedure’s effects on the animal and

the skills of the research team and must be conducted with IACUC
approval.
Various clinical signs are indicative of a moribund condition in
laboratory animals. If any of these signs are noted, prompt consulta-
tion with veterinary staff or euthanasia should occur (Aldred et al.,
2002, IACUC-UPENN, 2011b, Madeddu et al., 2006, Nemzek et al.,
2004, Paster et al., 2009, Schenk et al., 2012):
• Any condition interfering with eating or drinking (e.g., dif-

ficulty with ambulation)
• Rapid weight loss or net weight loss of more than 20% of BW
• Prolonged inappetance
• Evidence of muscle atrophy
• Marked loss of body condition
• Diarrhea, if debilitating, or constipation
• Markedly discolored urine, polyuria, or anuria
• Roughened hair coat, hunched posture, lethargy, or persis-
tent recumbency
• Central nervous system disturbance: head tilt, seizures,
tremors, circling, paresis, paralysis
• Lack of mental alertness
• Coughing, labored breathing, nasal discharge, or respiratory
distress
• Jaundice or anemia (pallor)
• Bleeding from any site
• Excessive or prolonged hyperthermia or hypothermia
• Marked dehydration
• Measurable distention of abdomen or torso
Frequency of animal assessments and monitoring, as well as

the objective criteria to be used for health evaluations, should be
clearly described in any IACUC protocol. Collection of individual
assessments (scoring) on fur appearance, respiration rate and effort,
mobility and ambulation, behavior, and body condition will all con-
tribute to the overall decisions about enacting humane interventions
and e ndpoints for animals.
nutritional therapy considerations

Anorexia, while not always observed, typically manifests as a loss of
body condition and weight in critically ill rodents. Nutritional sup-
port is key in the care of rodents, as the healthy rodent requires a
maintenance rate of 150–350 kcal/kg/day. The nutritional require-
ment can be two to three times this maintenance rate when animals
are ill (Paul-Murphy, 1996).
Nutritional supplementation of high-quality, laboratory-certified
feedstuffs, when provided to debilitated rodents, is part of expected
supportive care. Modifications of the animal’s own diet (typically
rodent block chow) can be done with the addition of Nutri-Cal® (a high-
calorie, palatable veterinary gel supplement) or soaking/softening the
chow in Ensure® (a high-calorie, flavored human shake supplement).
For animals in poor health, food and fluid options are often provided
on the cage floor in disposable trays/dishes or can be delivered in
liquid format by oral gavage for direct instillation into the stomach.
Oral gavage of mice (Figure 4.2) using sucrose-coated n eedles has
been validated as a means to decrease stress and improve animal
welfare during the gavage process (Hoggatt et al., 2010).
An expansive selection of nutritional and fluid supplementation
products is available for laboratory rodents that are in subopti-
mal health. Syringe feeding will likely be tolerated if the animal
is not moribund. Nasogastric intubation is typically physiologi-
cally impossible in mice and may have limited applications in the
critically ill rat. The cachectic or critical patient may require more
calories, a more concentrated caloric diet, and high palatability
until the appetite is restored and the animal is eating on its own
(Klaphake, 2006). Supportive and routine feedings may be neces-
sary at high frequency until rodents are through the crisis phase
of their illness.
Specialty nutritional supplementation (e.g., Nutra-GelTM Diet, Bio-
Serv ®, Frenchtown, NJ, http://www.bio-serv.com/) can be provided
to boost palatability and caloric intake for rodents that are ill or have
undergone major surgery or for models with abnormal phenotypes
that affect mastication (Figure 4.3).
Independent studies conducted by the particular vendor compa-
nies have ascertained that rodents find flavors like bacon and molas-
ses particularly appealing for these products (e.g., Transgenic Dough
Diet TM, Bacon SoftiesTM, Supreme Mini-TreatsTM, Bio-Serv), and
they may also be used effectively in pregnant females (Love MashTM
Fig. 4.2 Oral gavage in mice. (A) A variety of plastic and stainless

steel gavage needles is available for dosing in rodents and can be
straight or curved. Flexible needles (left) reduce the likelihood of
inadvertent esophageal rupture but can be chewed by the animal.
Ball-tipped, curved (rat sized, middle left) and straight (mouse-size,
middle right and right) needles are readily sanitized but can induce
injury if their passage is forced. (B) Measuring the gavage needle
for appropriate length for dosing, from tip of nose to last rib. The
needle can be marked for easy visualization of the appropriate inser-
tion distance. (Reprinted with permission from AALAS. Turner, PV,
Pekow, C, Vasbinder, MA, and Brabb, T. 2011. J Am Assoc Lab Anim
Sci 50:614–627.)
Rodent Reproductive Diet, Bio-Serv) and for administration to dams

to p
revent cannibalism of litters (Figure 4.4).
Gel supplements can provide palatable choices to improve postsur-
gical weight gain and assist with boosting calories as well as hydration
and can be formulated to assist with oral analgesic delivery (e.g.,
DietGel® Recovery, DietGel® Boost, MediGel® Sucralose, ClearH2O®,
Portland, ME, http://clearh2o.com/) (Figure 4.5). If a nimals require
further or continuous fluid supplementation in addition to those
provided as a bolus subcutaneously or intraperitoneally, oral options
A B
Fig. 4.3 Supplementary nutritional products, including (A) Nutra-

GelTM Diet purified and (B) grain-based options. (Photos courtesy of
Bio-Serv ®.)
A B
C D
Fig. 4.4 Supplementary nutritional products, including (A)

chocolate-flavored Supreme Mini-TreatsTM, (B) Bacon SoftiesTM,

(C) Love MashTM Rodent Reproductive Diet, and (D) Transgenic Dough
Diet TM. (Photos courtesy of Bio-Serv ®.)
can include water provided in individual packages inside the cage

(HydroPac®, Seaford, DE, http://www.hydropac.net/) or as gel for-
mulations (e.g., HydroGelTM, ClearH2O®; and Napa Nector TM, Systems
Engineering, Napa, CA. http://www.selabgroup.com/welcome.htm).
Providing pelleted food in powdered form to young C57BL/6
mice has been shown to significantly affect ingestion; animals con-
sumed more food when presented in powdered form than when it
was presented as pellets. The significant difference was associated
Fig. 4.5 Fluid and diet supplements, including DietGel® Recovery,

HydroGel®, and DietGel® 76A. (Reprinted with permission from
ClearH2O®.)
with corresponding higher intake of nutrients, including calories,

from the powdered forms. Therefore, it is key to also consider the
physical form of the diet when providing supplementary nutrition to
minimize confounding influences and experimental variations (Yan
et al., 2011).
perioperative care considerations

The “Guide for the Care and Use of Laboratory Animals” emphasizes
that successful surgical outcomes require appropriate attention to
presurgical planning, personnel training, anesthesia, aseptic and
surgical techniques, assessment of animal well-being, appropriate
use of analgesics, and monitoring of the patient’s physiologic sta-
tus during the procedures and beyond. There is an expectation that
training aspects of perioperative care will be provided to inform
research personnel performing surgery about maintenance of aseptic
technique during survival surgery (National Research Council, 2011).
A dedicated rodent surgical facility is not required, yet a d esignated
animal procedure area is necessary and should be located where
it can be maintained in an orderly, noncluttered status at all times
during surgery to minimize potential contamination of the patient.
This dedicated area should be away from heavy personnel traffic flow
and other unrelated activities and should have a s urgical surface
constructed of a readily disinfected material.
Aseptic surgery is performed with sterile gloves and sterilized (auto-
claved) instruments and materials (Figure 4.6) and takes precaution
Fig. 4.6 Perioperative considerations include provision of a sterile

pack with an enhanced draped space to allow for placement of
instruments and a larger sterile working surface. (Images courtesy
of University of Pennsylvania; S. Volk.)
to avoid introduction of infectious microorganisms to the patient.

Assurance that research personnel are properly trained in surgi-
cal preparation and technique must be documented in the IACUC
a nimal use protocol; personnel should wear sterile gloves and masks
while performing surgery. Gloves must be replaced if aseptic tech-
nique is disrupted (e.g., touching the isoflurane vaporizer with sterile
gloves, moving the animal with sterile gloves). With proper planning,
simple survival rodent surgeries may be performed by one person. If
this cannot be accomplished because of the complexity of the proce-
dure, then to consistently maintain aseptic technique, there should
be a surgical assistant or anesthetist who is trained to perform such
tasks that would otherwise interfere with proper aseptic technique.
If it is necessary for the surgeon to leave the surgical area during a
procedure, then they must reglove before resuming surgery.
Ensuring surgical procedures are kept sterile will lead to a
decreased chance of subclinical infections postoperatively or other
adverse postoperative outcomes. In addition, tissue should be han-
dled gently, with appropriate size instruments for rodent surgery.
Rodent preparation prior to surgery typically includes the removal
of the hair coat in a wide area around the intended incision site.
Excess clipped or depilated hair and other gross debris can be wiped
from the area using sterile gauze or a small disposable alcohol pad;
however, dousing the animal with alcohol is not recommended due

to potential for hypothermia as the alcohol evaporates from the
skin. The area can then be disinfected with appropriate surgical
scrub. Alcohol alone is not an appropriate disinfectant. Iodophors
(e.g., betadine) or chlorohexidines may be used and then wiped away
with sterile warmed or room temperature saline. Preemptive anal-
gesia will help to suppress pain responses that may be experienced
postoperatively, particularly through administration of opioids prior
to initiation of surgery (Flecknell, 2001). As well, and as mentioned
throughout the text, ocular protection with artificial tears (Rugby®
Sterile Artificial Tears Ointment Lubricant–Ophthalmic Ointment),
vitamin A ophthalmic ointment, or sterile lubricant (e.g., PuralubeTM)
is r ecommended for any animal undergoing anesthesia (Figure 4.7).
Attention to appropriate draping can increase the potential to
maintain a sterile surgical field. It is recommended that transparent
surgical drapes be utilized to ensure the patients can be observed
and monitored under anesthesia (Figure 4.8). As well, adhesive
drapes are being used increasingly with rodent patients to help affix
the anesthetic-delivering nose cone to the animal and the patient to
the surgical surface.
Fig. 4.7 Protective ophthalmic ointment should always be applied to

mice and rats undergoing anesthesia for any length of nonsurgical
or surgical procedure. Once the animal is sedated, ointment (Rugby ®
Sterile Artificial Tears Ointment Lubricant–Ophthalmic Ointment)
can be dripped directly onto the eye from the application vial (left) or
transferred to a sterile swab and then touched to the eyes (middle).
The eyes typically remain open (right) during anesthesia, and the
ophthalmic film will provide a protective layer to prevent desiccation
of the globe and surrounding structures. (Images courtesy of the
Fig. 4.8 A rodent leg draped with transparent adhesive TegadermTM

dressing to permit an enhanced view of the surgical site (left); the
animal can be draped first with a standard disposable blue cloth
and the adhesive used to hold the drape in place (right) while permit-
ting a larger sterile work space for the surgeon. (Images courtesy of
University of Pennsylvania; S. Volk.)
Advantages of this level of restraint include insurance that the

animal will not become disconnected from the source of anesthesia
and that there will not be mobility of the patient during surgical
manipulations (Locke et al., 2011). Adhesive drapes (i.e., TegadermTM
Transparent Dressing, 3M, Minneapolis, MN) can be ordered in
varying sizes as they are typically applied to human wounds. These
sterile film dressings provide permeability for oxygen and moisture
exchange while limiting access of debris and pathogens to the inci-
sion site. For mice, drape sizes of 4 × 4¾ inches are appropriate for
an abdominal procedure; rats will typically require a larger adhesive
drape size of 6 × 8 inches. Anecdotally, sterile over-the-counter plas-
tic wrap (e.g., SaranTM wrap) has been used successfully as a trans-
parent rodent drape. For the novice surgeon, a larger s terile drape
can be placed first, with the adhesive transparent drape placed over
the surgical window to hold the larger drape in place, which provides
a wider area of sterility for instrument placement and suture stor-
age, yet the animal can still be visually monitored for respiration and
movement during the procedure. Due to the oxygen-rich environ-
ment of the surgical area, coupled with the potential to use cautery
or other electrical devices within close proximity, one should practice
fire safety measures to avoid hazards that could harm personnel and
animals (Caro et al., 2011, Klein, 2008).
Surgical monitoring includes confirmation of anesthetic depth
to an appropriate level for the procedure. For most rodents, a brief
and strong manual pressure on the toes (“toe pinch”) should not
elicit a withdrawal response if the animal is at a surgical plane of

a nesthesia. There should also be no palpebral response, confirmed
by gently t apping the medial aspect of the rodent eyelid without elic-
iting a blink response. If anesthesia becomes light (and the rodent
potentially conscious) during a surgical procedure, the animal may
move in response to skin incisions and tissue manipulations and
exhibit a rise in heart and respiratory rates. Further attempts to
complete the procedure must cease until the animal is returned to
a surgical plane of anesthesia. Anesthetized animals should never
be left alone during the surgical procedure. Tissue manipulations
should be performed gently to reduce the degree of postoperative
pain; stretching and pulling on sensitive structures triggers noci-
ceptive nerve endings, and these impulses may heighten pain levels
following recovery from a nesthesia (Flecknell, 2001).
Postsurgical care management should include collaborative assis-
tance between veterinary and research staff members. Sutures are
more likely to remain intact, without chance of evisceration, if absorb-
able sutures or skin staples are used. Incisional hernia is the most
common abdominal wall defect, usually formed by trauma or infection.
Defects can be patched with synthetic materials, such as p olypropylene
mesh (SeprameshTM) inserted in a “sandwich” form between injured
peritoneum and abdominal muscles. The mesh can be combined
with Seprafilm®, a bioresorbable membrane that adheres and p romotes
the normal healing process. Adhesions and inflammation have been
documented to be reduced when these m embrane products were used
for abdominal wall repairs (Esfandiari and Nowrouzian, 2006).
Animals should be placed into a clean cage space, notably without
other awake animals, until fully recovered from anesthesia. Housing
animals individually after surgery may be necessary to promote
anesthetic recovery and to prevent potential for injury from cage
mates; however, both the physical and the social environment may
affect the way in which an animal copes with the stress associated
with postoperative recovery.
The recovery cage can be placed on top of a heat source, or heat
supplementation can be provided until full recovery from anesthesia
is observed. Fluid gel and soft food (standard food pellets soaked in
sucrose water) for up to 4 days postoperatively can decrease poten-
tial for dehydration and extreme weight loss; supplemental subcuta-
neous fluids may also be useful to support postprocedural recovery.
Observations and notations of all monitoring and interventions are rec-
ommended to occur twice daily for at least 72 h following c
ompletion of
surgery and recovery from anesthesia (Kalishman et al., 2004). During
the postsurgical period, animals should be monitored for signs of pain

or distress, as described previously (see Chapter 1). In most species,
signs of pain include decreased activity, abnormal posture, increased
attention to surgical site, and gait abnormalities. Analgesia and pain
relief regimens should be based on the species, the type of procedure
performed, the pharmacokinetics of available agents, and any known
adverse effects of the specific drugs (Lester et al., 2012). Further infor-
mation is available in the rodent formulary in Appendix C, along with
the additional insights gleaned from p ublished analgesic regimens.
Maintenance of a surgical record for each patient will be h elpful
to track patient health relative to the presurgical baseline data.
Postsurgical documentation (medical records) should include the
protocol number, animal identification, observations, date of obser-
vation, comments on the general condition and health of animal,
and analgesics or other medications given. The specific date, time,
and dose of the administered a nalgesics should be written into the
postoperative record. As is expected for a medical record, the surgical
record should also include initials of individuals writing the entries.
regulatory considerations
Currently, laboratory mice (of the genus Mus) and rats (of the genus
Rattus) used in biomedical research are exempt from oversight by the
Animal Welfare Act and Regulations. However, aspects of their use,
care, and treatment are covered in both the Guide for the Care and
Use of Laboratory Animals (National Research Council, 2011) and the
Public Health Service Policy on Humane Care and Use of Laboratory
Animals, which is specific to coverage of animals used in research
funded by the Public Health Service through the National Institutes
of Health in the United States (Public Health Service, 1996).
Descriptions of animal models, group sizes, experimental time-
lines, outcomes, adverse events, and humane endpoints are requested
in typical IACUC protocol templates and must be approved by the
IACUC prior to initiation of any experiments. As well, descriptions of
drug types and dosages are required to adhere to veterinary stan-
dards of practice. The selection of appropriate sedatives, analgesics,
and anesthetics is a moral imperative to minimize, if not eliminate,
animal sensation of pain or distress (National Research Council,
2011). Finally, all drugs must be “in date,” meaning not used past the
expiration date stamped on the vial or package, to adhere with man-
ufacturer recommendations and federal regulations and guidance.
Clinical treatments selected in response to an emergency or c ritical

care situation may not always be included in approved protocols, yet
consultation with veterinary staff will permit the limited application
of off-protocol drugs and therapeutics for the benefit of overall animal
welfare. However, once a clinical treatment has been applied, and if
it will be used for additional animals in an experimental cohort, this
relevant information must be amended into the existing animal care
protocol for the IACUC to review and approve.
restraint collar considerations

Collars can be used for prevention of undesirable behaviors (Brown,
2006a, 2006b). Restraint or neck safety collars, most commonly
referred to as “Elizabethan collars,” have been applied in numer-
ous animal species to prevent coprophagy, self-grooming, licking
test compounds off skin, and access to and potential self-injury of
postsurgical sites (Figure 4.9).
Considerations in the use of restraint collars include the following:
• Stress induction with placement of the collar, which can result

in transient depression in eating and subsequent weight loss
or potential for hyperthermia in small rodents.
A B C
Fig. 4.9 Restraint collars for rodents: (A) low-density polyethylene

Elizabethan collars; notice how the neck openings are either bound
with cotton jersey for light padding or padded with soft vinyl foam
(B). A rat with a caudally directed e-collar (C). Transparent collars
that allow an animal to see through them allow for better adjustment
than if the collar is made of a solid material or color that obstructs
their view. Rodents also perform coprophagy and will need access to
consume fecal pellets on the floor of their caging. On a nutritionally
complete diet, rats will eat about 10% of their feces. (Reprinted by
permission from Macmillan Publishers Limited. Brown, C. 2006a. Lab
Anim (NY) 35:23–25; and Brown, C. 2006b. Lab Anim (NY) 35:25–27.)
• Acclimation period, with increasing time wearing the collar

prior to its actual experimental use in the animal, monitored
by an observer.
• Fit must be appropriate: not too large to avoid entrapment or
further injury and not too small so that the a nimal chokes
or that the skin becomes abraded and ulcerated. Rule: Any
Elizabethan collar should be able to rotate 360° with minimal
difficulty.
• No sharp edges should be left on the collar; attempts should
be made to pad the inside edge thoroughly or place a thick
layer of tape so the edges are blunted.
• Consider individual housing of the animal during the time
when the animal is wearing the collar; there may be intraspe-
cies aggression from the noncollared animals toward the one
with the collar.
• It is critical that the animals be able to access food and water
sources in their housing/holding areas. Changes to confor-
mation of food and water receptacles may need to be under-
taken so that animals can manipulate their faces toward the
sustenance without removing the collar.
• Application of aversive nontoxic compounds (metronidazole
500 mg powder mixed with 1 ml New Skin® liquid bandage)
to susceptible body parts, in particular following spinal cord
injury, has assisted with maintaining animal welfare through
prevention of self-injury and autophagia (Zhang et al., 2001).
tracheostomy considerations
As a final or “salvage” procedure for a moribund yet invaluable rodent
model, one may attempt to correct respiratory arrest and loss of con-
sciousness by inserting a tracheostomy tube. Due to the critical
state of the moribund patient and need to intervene immediately,
the tube can be placed under local (topical) anesthesia even in a
conscious animal. One can use individual sterile alcohol swabs to
part the hair over the ventral neck and visualize the trachea through
the skin. A skin incision, using sterile instruments or scalpel blade,
should be carefully and gently made just over the area of the trachea,
immediately distal to the throat (larynx). Accessing the trachea is
best done by gently retracting it into the incision and placing a stay
suture underneath; this will help to keep the trachea everted prior to
incising between the cartilaginous rings. Once the trachea is isolated,

proceed to exert minimal force and gently guide a s terile 16-gauge
catheter or 1.0- to 2.0-mm endotracheal tube (if rodent > 100 g) into
the distal trachea. Oxygen should then be provided and the animal
closely monitored for a return to respiratory function (Paul-Murphy,
1996). The prognosis for this type of procedure is likely poor, given
the impaired condition of the patient and any additional physiologic
stress experienced by way of this invasive event.
tumor development and

monitoring considerations
For those experiments involving tumors, overall tumor burden
should be limited to the minimum required to meet the study objec-
tives. The general health and overall condition of the animal is to
be assessed with increasing frequency as expected tumors develop.
Adverse effects on the rodent will depend on the biology, site, mode
of growth of the tumor, and any additional procedures or treatments
(Workman et al., 2010). If tumors grow unexpectedly and are not
directly related to proposed experiments, research staff should con-
sult with a veterinarian to determine the best course of treatment
and interventions. It may be possible to have the tumor excised or
treated in some other manner to continue maintaining the animal
in a study. Awareness of the types of cancer that may cause inher-
ent pain to the animal should be considered with respect to humane
interventions and endpoints (Pacharinsak and Beitz, 2008).
Tumor implantation sites should be chosen to minimize damage to
adjacent normal structures; in particular, implantation of tumors on
the dorsum or flank of the rodent will likely have limited site-related
morbidity. Implantation of tumor cells to the face, limbs, or perineum
should be avoided as there is little to no space for tumor growth and
expansion. Intramuscular implantation should be avoided as this
may be painful due to the distension of the muscle by the tumor.
Tumor implantation on the ventral surface of the body should also
be avoided due to the risk of irritation to the tumor site in contact
with the bedding and floor of the cage. Tumor development should
not interfere with normal gait or postures; should not interfere with
vital functions (eating, drinking, breathing); should not result in
painful responses when palpated; and should not lead to persistent
self-trauma (likely secondary to pain). Determining the tumor bur-
den of internal cancers, lymphoreticular tumors, and metastatic
disease will be challenging and may require pilot studies in a smaller

number of animals to better characterize disease patterns (Workman
et al., 2010).
Evaluating Tumor Growth

• Animals that are in a tumor production study should be mon-
itored by the laboratory at least once weekly during the time
when the tumor is not yet detectable to observe when tumor
growth begins. After a visual or palpable tumor is evident,
animals should be monitored by the laboratory at least twice
weekly. More frequent observations may be necessary based
on tumor growth rate, study parameters, and general condi-
tion of the animal.
• Evaluating tumors involving nonsurface areas of the rodent
(e.g., bone, brain, lungs, internal organs) can be challeng-
ing. Objective tumor size cannot be routinely assessed, and
a limited tumor burden (well below the recommended maxi-
mum size) may cause impairment and other clinical signs.
For tumor models studying non-surface-area tumors, BCS
and clinical evaluations of the animals take priority over the
measured size of the tumor (Paster et al., 2009). The expected
clinical signs and the humane endpoints of those signs must
be clearly described in the protocol.
• Evaluating tumor growth on surface area, on the basis of a
percentage of BW, is generally inaccurate. While the grow-
ing tumor likely will cause an increase in body weight, the
general condition of the rodent may be decreased (losing lean
body mass), resulting in a relatively stable body weight but
a progressively more unhealthy animal. Therefore, tumor
growth should be monitored in the context of the evolving
BCS, objective dimensional criteria (size) of the tumor bur-
den measured by caliper or other mechanism, tumor location,
number of tumors, and tumor ulceration.
The guidance that follows assumes that a normal size adult rodent
will be studied (approximately a 25-g mouse or a 250-g rat). The
allowable sizes of tumors will be decreased if the tumors are injected
into immature or genetically small mice. When on the dorsum or
flank of an adult rodent, tumors may be allowed to grow as large
as diameters of about 2.0 cm (or 4.2 cm3) in mice and about 4.0 cm
(or 33.5 cm3) in rats as long as the rodent remains otherwise healthy.

The main concerns for permitting individual tumors to develop
beyond these recommended size limitations are related to the poten-
tial for ulceration of the tumor, with central necrosis of the skin over-
lying the mass, and potential further injury and poor health.
Tumor Ulceration
Ulceration of a tumor does not necessarily correlate with tumor size
or require euthanasia of the animal, but it does typically require more
frequent monitoring and treatment of the ulcerated site. Ulceration
can lead to discomfort related to the loss of skin integrity or local-
ized infection; as well, hemorrhage at the site of ulceration may
occur, and the site may become prone to infection (Narver, 2013).
The level of follow-up care for ulcerated tumors is based on both
the size of the ulceration and clinical judgment by the veterinarian.
Recommendations for monitoring of ulcerated tumors include the
following:
• Pinpoint (< 1 mm) ulcerations at the site of tumor injection

should be monitored at least two times per week for worsen-
ing of the ulceration site.
• Ulcerations (> 1 mm) of the surface area of the tumor should
be monitored at least three times per week and should be
reported to veterinary staff for evaluation and potential
treatment.
Multiple Tumors
Multiple tumors that are individually smaller than the single tumor
limit may not have the same negative sequellae as a single tumor.
Multiple tumors may be allowed to grow up to 150% of the volume
compared with the volume of a single tumor. The size limitation of
the diameter of any single tumor (2.0 cm in mice or 4.0 cm in rats)
should still be applied. Institutional allowance on permissible sizes
of tumors typically will be decreased if the tumors are transplanted
into immature or genetically runted mice.
Ascites Produced by Tumors

If tumors are expected to grow with accumulation of fluid in the peri-
toneal cavity (ascites), rodents must be weighed prior to inoculation
and subsequently be weighed at regular intervals. All monitoring

should be thoroughly described in the IACUC protocol and based
on the expected rate of fluid accumulation. Ascites pressure should
be relieved before abdominal distension is great enough to cause
discomfort or interfere with normal activity. When the BW exceeds
120% of baseline weight, the animal should be euthanized or the
fluid removed (“tapped”) from the abdominal cavity. The abdominal
tap should be performed by trained personnel using proper a septic
technique, with manual restraint or anesthesia, and by using
the smallest needle (18–22 gauge) possible that still permits fluid
removal. Certain institutions may have limitations on the number of
abdominal taps that can be performed prior to the animal reaching
a humane endpoint.
Aspects of this section were adapted from institutional documents
(IACUC-CORNELL, 2013, IACUC-UPENN, 2010).
wound management considerations

General anesthesia may be necessary for initial wound assessments
in rodents, depending on severity. Wounds typically undergo four
stages of healing: inflammatory, debridement, repair, and matura-
tion. Appropriate wound therapy is related to determination of the
stage of healing (Langlois, 2004). Many factors have an adverse
impact on wound healing of the critical patient, including nutritional
status, immune status, concurrent disease (e.g., diabetes mellitus),
neoplasia and paraneoplastic syndromes, and location of the wound.
Wound management involves culture and antibiotic sensitiv-
ity testing, decontamination (clipping of surrounding hairs after
covering wound with a water-soluble lubricant), wound lavage with
warm sterile saline, application of antiseptic ointment, and wound
debridement.
Topical treatments can be applied, with the realization that the
grooming behaviors of rodents may lead to inadvertent ingestion of the
substances. However, similar treatments to those listed for murine
ulcerative dermatitis (UD) (see relevant section in Chapter 2) can
be applied, including Neosporin®, Silvadene®, Preparation H® oint-
ment to stimulate collagen synthesis, aloe vera, CarraVet® wound gel,
and sugar and unpasteurized honey (Langlois, 2004, Mathews and
Binnington, 2002a, 2002b). Toenails should be trimmed to diminish
ability to self-injure by scratching at the wound site.
Ideally, wounds should be closed (whether by suture, steel skin

clips/staples, or tissue glue like VetbondTM), taking care to avoid
creation of tension across tissues, and protected to avoid disrup-

tion of the granulation tissue formation (Hernandez-Divers, 2004). If
suture closure is preferred, a suture size of 3–0 thickness or smaller
is p referred in rodents; individually packaged commercial suture
materials, with attached tapered needles, are commonly used.
Rodents will have a propensity to chew at incision sites, often as a
manifestation of discomfort; therefore, care should be taken to try to
bury the suture line into the subcuticular layer and provide appro-
priate dosages of analgesics. If skin clips or sutures are placed, they
should be removed 10 to 14 days after placement. Restraint collars
(see relevant section on this topic) can also be applied during the
first 7–10 days after wounds are dressed. Additional comprehensive
information about critical care management of wounds is reviewed
extensively in other texts (Garzotto, 2009, Langlois, 2004).
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5
resources and additional
information
introduction
Additional resources and helpful references are provided in this
chapter for both the general specialty of laboratory animal science
and specifics related to clinical laboratory animal medicine.
organizations
Professional organizations that provide clinical laboratory animal
medicine information are limited; however, those that exist provide
access to resources for their members and promote a network of col-
laboration between professionals in the field.
American Association for Laboratory

Animal Medicine (AALAS)
http://www.aalas.org/
AALAS serves a diverse professional group, ranging from
research investigators to animal care technicians to vet-
erinarians. AALAS publishes relevant specialty journals
and materials, from which the majority of materials for this
text were derived. These publications include the Journal of
41
AALAS (formerly Contemporary Topics in Laboratory Animal

Science), Comparative Medicine (formerly Laboratory Animal
Science), and Tech Talk. AALAS began to publish an additional
journal, Laboratory Animal Science Professional, in 2013. This
organization offers an online comprehensive training m odule
library, the AALAS Learning Library, which offers members
a diverse listing of courses relevant to laboratory animal
science. The AALAS website provides a link (http://national-
meeting.aalas.org/past_meeting_abstracts.asp) for members
to access those abstracts (dating through 1992) presented
at the annual national AALAS meetings; these abstracts
provide key suggestions and considerations for
laboratory
animal care.
American College of Laboratory

Animal Medicine (ACLAM)
http://www.aclam.org/
The American College of Laboratory Animal Medicine is
comprised of veterinarians certified in the specialty of

laboratory animal medicine. This group conducts an annual
ACLAM Forum for continuing education to advance the
humane care and responsible use of laboratory animals.
ACLAM posts published position statements and reports
(http://w w w.aclam.org/education-and-training/position-
statements-and-reports) on topics including veterinary care,
animal experimentation, pain and distress, rodent surgery,
and rodent euthanasia.
American Society of Laboratory

Animal Practitioners (ASLAP)
http://www.aslap.org/
ASLAP membership includes veterinary professionals, train-
ees, and students with an interest in laboratory animal
practice. ASLAP supports educational sessions aimed to
promote knowledge, ideas, and information for the benefit
of animals and society at both the national AALAS and the
annual American Veterinary Medical Association (AVMA)
conferences.
resources and additional information 43
Institute for Laboratory Animal Research (ILAR)
http://dels.nas.edu/ilar/
The mission of ILAR is to evaluate and disseminate information
on issues related to the scientific, technological, and
ethical use of animals and related biological resources in

research, testing, and education. The organization p ublishes
comprehensive topical issues of the ILAR Journal on relevant
subjects for the care and use of laboratory animal species and
those individuals that work with them. ILAR functions as a
component of the National Academies to p rovide e
xpertise to
the federal government, the international biomedical research
community, and the public.
publications
Published materials, books, journals, and other documents are
extremely valuable resources for clinical laboratory animal informa-
tion and discussion of relevant experimental models.
Books
Banks, RE, Sharp, JM, Doss, SD, and Vanderford, DA. Exotic Small
Mammal Care and Husbandry. Wiley-Blackwell, Ames, IA, 2010.
Birchard, SJ, and Sherding, RG. Saunders Manual of Small Animal
Practice, 3rd edition. Saunders, Philadelphia, 2005.
Danneman, P, Suckow, MA, and Brayton, C. The Laboratory Mouse,
2nd edition. CRC Press, Boca Raton, FL, 2012.
Ford, RB, and Mazzaferro, E. Kirt & Bistner’s Handbook of Veterinary
Procedures and Emergency Treatment, 9th edition. Saunders,
Philadelphia, 2012.
Fox, JG, Barthold, SW, Davisson, MT, Newcomer, CE, Quimby, FW,
Smith, AL. The Mouse in Biomedical Research. American College
of Laboratory Animal Medicine Series. Elsevier via Academic
Press, New York, 2007.
Gaertner, DJ, Hankenson, FC, Hallman, T, and Batchelder, MA.
Anesthesia and analgesia in rodents, Chap. 10. In Fish RE, Brown,
MJ, Danneman PJ, and Karas, AZ (eds.), Anesthesia and Analgesia
for Laboratory Animals. Academic Press, San Diego, CA, 2008.
Heinlich, H, Bullock, GR, and Petrusz, P. The Laboratory Mouse.

Academic Press, San Diego, CA, 2004.
Hrapkiewicz, K, and Medina, L. Clinical Laboratory Animal Medicine: An
Introduction, 3rd edition. Wiley-Blackwell, Ames, IA, 2006.
Johnson-Delaney, C.A. Exotic Companion Medicine Handbook for
Veterinarians. Wingers, Lake Worth, FL, 1996.
Macintire, DK, Drobatz, KJ, Haskins, SC, and Saxon, WD. Manual of
Small Animal Emergency and Critical Care Medicine. Lippincott
Williams & Wilkins, New York, 2005.
Murtaugh, RJ, and Kaplan, PM. Veterinary Emergency and Critical
Care Medicine. Mosby-Year Book, St. Louis, MO, 1992.
National Research Council (NRC). Recognition and Alleviation of
Distress in Laboratory Animals. National Academies Press,
Washington, DC, 2008.
National Research Council (NRC). Recognition and Alleviation of Pain
in Laboratory Animals. National Academies Press, Washington,
DC, 2009.
National Research Council (NRC). Guide for the Care and Use of
Laboratory Animals, 8th edition. National Academies Press,
Washington, DC, 2011.
Oglesbee, BL. Blackwell’s Five-Minute Veterinary Consult: Small
Mammal, 2nd edition. Wiley-Blackwell, Ames, IA, 2011. [Note:
Contains a section on rodents.]
Percy, DH, and Barthold, SW. Pathology of Laboratory Animal
Rodents, 3rd edition. Wiley-Blackwell, Ames, IA, 2007.
Plunkett, SJ. Emergency Procedures for the Small Animal Veterinarian,
2nd edition. Saunders, Philadelphia, 2000. [Note: Contains a
section on exotics.]
Pritchett-Corning, KR, Girod, A, Avellaneda, G, Fritz, PE, Chou, S, and
Brown, MJ. Handbook of Clinical Signs in Rodents and Rabbits.
Charles River Laboratories, Wilmington, MA, 2010.
Quesenberry, K, and Carpenter, JW. Ferrets, Rabbits and Rodents:
Clinical Medicine and Surgery, 2nd edition. Saunders,
Philadelphia, 2003.
Sharp, PE, and Villano, J. The Laboratory Rat, 2nd edition. CRC
Press, Boca Raton, FL, 2012.
Silverstein, D, and Hopper, K. Small Animal Critical Care Medicine.
Saunders, Philadelphia, 2008.
Suckow, MA, Weisbroth, SH, and Franklin, CL. The Laboratory Rat, 2nd
edition. ACLAM series. American College of Laboratory Animal
Medicine Series. Elsevier via Academic Press, New York, 2006.
Suckow, MA, Stevens, KA, and Wilson, RP. The Laboratory Rabbit,
Guinea Pig, Hamster and Other Rodents. American College of
Laboratory Animal Medicine Series. Elsevier via Academic Press,
New York, 2012.
Periodicals
ALN® (Animal Lab News) Magazine [published by Vicon Publishing,

Inc.], http://www.alnmag.com/
Comparative Medicine [published by AALAS] (formerly Laboratory
Animal Science)
ILAR Journal [published by Oxford Journals], http://ilarjournal.
oxfordjournals.org/
Journal of the American Association for Laboratory Animal Science
[
published by AALAS] (formerly Contemporary Topics in
Laboratory Animal Science)
Journal of Exotic Pet Medicine [published by Elsevier, Inc.], http://
www.exoticpetmedicine.com/home
Lab Animal magazine [published by Nature Publishing Group], http://
www.labanimal.com/laban/index.html
Laboratory Animals [published by the Royal Society of Medicine jour-
nals], http://la.rsmjournals.com/
Laboratory Animal Science Professional [published by AALAS]
Tech Talk [published by AALAS]
Veterinary Clinics of North America—Exotic Animal Practice [published
by Elsevier, Inc.], http://www.vetexotic.theclinics.com/
electronic resources
AALAS Learning Library
https://www.aalaslearninglibrary.org/default.asp
The AALAS Learning Library provides training modules of
benefit for technicians, veterinarians, managers, IACUC

embers, and investigators working with animals in a

m
research or educational setting.
Animal Care Training Services (ACTS)
http://actstraining.com/
ACTS is a training company that specializes in the daily opera-
tions of lab animal research institutions. They provide “job-
specific skill training” program modules, assist with training
of staff to achieve AALAS certification levels for technicians,
and support on-site seminars on a variety of customized
training topics at local and regional sites.
AVMA Guidelines for the Euthanasia

of Animals: 2013 Edition
https://www.avma.org/KB/Policies/Documents/euthanasia.
pdf
The 2013 guidelines, established by membership of the Panel
on Euthanasia, set criteria for euthanasia, specified appro-
priate euthanasia methods and agents, and are intended to
assist veterinarians. In this version, methods, techniques,
and agents of e uthanasia have been updated, and detailed
descriptions have been included to assist veterinarians in
applying their professional and clinical judgment.
CompMedTM listserv
h t t p :// w w w. a a l a s . o r g /o n l i n e _ r e s o u r c e s/ l i s t s e r v e s .
aspx#compmed
CompMed is an e-mail list for discussion of comparative medi-
cine, laboratory animals, and topics related to biomedi-
cal research. CompMed is limited to participants who are
involved in some aspect of biomedical research or veterinary
medicine, including veterinarians, technicians, animal facil-
ity managers, researchers, and graduate/veterinary stu-
dents. AALAS membership is not required to subscribe to
this group.
To subscribe:
Send e-mail to: LISTSERV@LISTSERV.AALAS.ORG
Message body: SUBSCRIBE COMPMED Yourfirstname Yourlast

name (Example: SUBSCRIBE COMPMED John Doe)
Drug Enforcement Agency (DEA),

Office of Diversion Control
http://www.deadiversion.usdoj.gov/index.html
This website provides information about registering to obtain
licensure for appropriating controlled substances (drugs)
for use in veterinary medicine in the United States and its
territories.
IACUC-Forum listserv
http://www.aalas.org/online_resources/listserves.aspx#IACUC-
Forum
IACUC-Forum is a member benefit for current AALAS insti-
tutional members. There are no fees for this service; it is
included as part of institutional membership dues. Current
institutional contact persons may enroll their IACUC mem-
bers and IACUC staff on IACUC-Forum; the IACUC members
and IACUC staff who have access to the list are not required
to be members of AALAS for the purposes of this list. Only
individuals directly related to the IACUC are eligible to have
access to the list.
To subscribe, complete and submit the application form found
on the web link.
International Council for Laboratory

Animal Science (ICLAS)
http://iclas.org/
ICLAS is the international scientific organization dedicated to
advancing human and animal health by promoting the ethi-
cal care and use of laboratory animals in research world-
wide. From the ICLAS membership page (http://iclas.org/
members/member-list), the following international laboratory
animal science groups can be accessed:
Asociación Argentina de Ciencia y Tecnología de Animales de
Laboratorio (AACyTAL)
Asociación Chilena de Ciencias del Animal de Laboratorio

(ASOCHICAL)
Asociacion Mexicana de la Ciencia de los Animales de
Laboratorio (AMCAL)
Asociación Uruguaya de Ciencia y Tecnología de Animales de
Laboratorio (AUCyTAL)
Association Française des Sciences et Techniques de l’ Animal
de Laboratoire (AFSTAL)
Associations of Central America, Caribbean and Mexico
Laboratory Animal Science (ACCMAL)
Associazione Italiana per le Scienze degli Animali da
Laboratorio (AISAL)
Australia and New Zealand Laboratory Animal Association
(ANZLAA)
Belgian Council for Laboratory Animal Science (BCLAS)
Canadian Association for Laboratory Animal Science (CALAS/
ACSAL)
Chinese Association for Laboratory Animal Science (CALAS,
China)
Chinese–Taipei Society of Laboratory Animal Sciences
(CSLAS)
Finland Laboratory Animal Science (FinLAS)
German Society for Laboratory Animal Science (GV-SOLAS)
Israeli Laboratory Animal Forum (ILAF)
Japanese Association for Laboratory Animal Science (JALAS)
Japanese Society for Laboratory Animal Resources (JSLAR)
Korea Research Institute of Bioscience and Biotechnology
(KRIBB)
Korean Association for Laboratory Animal Science (KALAS,
Korea)
Laboratory Animal Science Association (LASA, United
Kingdom)
Laboratory Animal Scientist’s Association (LASA, India)
Laboratuvar Hayvanları Bilimi Derneği (TURKEY)/
LASA-Turkey
Nederlandse Vereniging voor Proefdierkunde, (NVP)/
Biotechnische Vereniging (BV)
Scandinavian Society for Laboratory Animal Science

(SCAND-LAS)
Sociedad Española Para las Ciencias del Animal De
Laboratorio, (SECAL)
Sociedade Brasileira de Ciencia de Animais de Laboratorio
(SBCAL)
South-African Association for Laboratory Animal Science
(SAALAS, South Africa)
Swiss Laboratory Animal Science Association–Schweizerische
Gesellschaft für Versuchstierkunde (SGV)
Thai Association for Laboratory Animal Science (TALAS,
Thailand)
International Mouse Strain Resource (IMSR)
http://www.findmice.org/
The IMSR is a multi-institutional international collaboration
supporting the use of the mouse as a model system for study-
ing human biology and disease. The primary goal of the
IMSR is to provide a web-searchable catalog that will assist
the international research community in finding the mouse
resources needed.
The IMSR began with an initial collaboration between the Mouse
Genome Informatics (MGI) group at the Jackson Laboratory
and the Medical Research Council Mammalian Genetics Unit
at Harwell, United Kingdom. Many institutions and collabo-
rators are now contributing mouse resource information to
the IMSR catalog.
Mouse Genome Database
http://www.informatics.jax.org/
The U.S. National Institutes of Health provide support for this
reference database maintained through the website of the
Jackson Laboratory. This database provides a resource for
mouse genetic, genomic, and biological information, such
as gene characterization, characteristics of inbred strains,
descriptions of mutant phenotypes, and additional related
subjects.
National Institutes of Health Office of Research

Infrastructure Programs (ORIP): Rodent Resources
http://dpcpsi.nih.gov/orip/cm/rodents_index.aspx
ORIP’s laboratory rodents program funds development of
genetically engineered rodents and research rodent colonies,
facilities that distribute rodents and related biological materi-
als, and new ways to study, diagnose, and eliminate labora-
tory rodent disease. Related links from this page include the
following:
Rodent Resources for Researchers, a listing of hyperlinks
to various mutant mouse resource centers, phenotyping
programs, mutant rat resources, and resources for rat

genetic models (http://dpcpsi.nih.gov/orip/cm/rodent_
resource_researchers.aspx).
Office of Laboratory Animal Welfare
http://grants.nih.gov/grants/olaw/olaw.htm
The Office of Laboratory Animal Welfare (OLAW) provides guid-
ance and interpretation of the Public Health Service (PHS)
Policy on Humane Care and Use of Laboratory Animals, sup-
ports educational programs, and monitors compliance with the
policy by assured institutions and PHS funding components to
ensure the humane care and use of animals in PHS-supported
research, testing, and training, thereby contributing to the
quality of PHS-supported activities. The site contains an exten-
sive listing of answers to frequently asked questions, providing
further commentary on topics related to research animal wel-
fare (e.g., pharmaceutical-grade drug definitions, euthanasia,
housing expectations per the National Resource Council’s 2011
Guide for the Care and Use of Laboratory Animals [National
Academies Press, Washington, DC]).
Pubmed
http://www.ncbi.nlm.nih.gov/pubmed
PubMed is an electronic database supported by the U.S. National
Library of Medicine and National Institutes of Health; it com-
prises more than 22 million citations for biomedical litera-
ture from MEDLINE, life science journals, and online books.
Citations may include links to full-text content from PubMed

Central and publisher websites.
Rat Genome Database
http://rgd.mcw.edu/
The Rat Genome Database is a collaborative effort between lead-
ing research institutions involved in rat genetic and genomic
research. This resource is monitored and supported by grant
HL64541. “Rat Genome Database,” awarded to Dr. Howard
J. Jacob at the Medical College of Wisconsin by the National
Heart Lung and Blood Institute (NHLBI) of the National
Institutes of Health (NIH). The Rat Genome Database was cre-
ated to serve as a repository of rat genetic and genomic data,
as well as mapping, strain, and physiological information.
It also facilitates investigators’ research efforts by providing
tools to search, mine, and analyze these data.
TechLink listserv
http://www.aalas.org/online_resources/listserves.aspx
TechLink is an electronic mailing list (listserve) created espe-
cially for animal care technicians in the field of labora-
tory animal science. Open to any AALAS national member,
TechLink serves as a method for laboratory animal techni-
cians to exchange information and conduct discussions of
common interest via e-mail messages with technicians in the
United States and other countries around the world.
To subscribe:
Send e-mail to: LISTSERV@LISTSERV.AALAS.ORG
Message body: SUBSCRIBE TECHLINK Yourfirstname
Yourlastname
(Example: SUBSCRIBE TECHLINK John Doe)
Veterinary Bioscience Institute (VBI)
http://www.vetbiotech.com/
VBI offers training modules for experimental and veterinary
surgical and biomethodology training for technical and medi-
cal staff. VBI provides online training with hands-on training
modules for a variety of rodent surgical procedures, including

innovative approaches like laparoscopy. The site provides fee-
for-service access to webinars and learning modules online.
Veterinary Emergency and Critical Care Society (VECCS)
http://www.veccs.org/
VECCS aims to raise the level of patient care for seriously ill or
injured animals through quality education and communica-
tion programs. The society works closely with the American
College of Veterinary Emergency and Critical Care (ACVECC)
to provide information related to life-threatening and acute
disease conditions in pet medicine.
Veterinary Information Network (VIN)
http://www.vin.com/VIN.plx
VIN serves as an online resource for veterinarians with content
submitted by veterinarians from various specialties in clinical
practice. Membership to the site, which supports conference
proceedings from a variety of veterinary annual conferences,
is for a fee; however, veterinary students and academicians
are allowed access at no charge.
Topics of interest can be searched for input from colleagues, and
continuing education courses and lectures are available.
commercial resources
ALN Buyer’s Guide
http://www.alnmag.com/buyers-guide [also available in hard-

copy format]
The ALN® Magazine and ALN World™ Buyer’s Guides are com-
prehensive sources of resources, products, and information
to design, build, and equip animal research facilities. Direct
links are provided to vendor and commercial information
concerning products for laboratory animals; animal care and
maintenance; facility design, materials, and equipment; labo-
ratory and research equipment and supplies; organizations;
surgical and medical equipment and supplies; veterinary
and research services; and other services, materials, and

equipment.
Lab Animal Buyer’s Guide
http://guide.labanimal.com/guide/index.html [also available in

hard-copy format]
A comprehensive database of suppliers, products, and services
in laboratory animal care. The source can be searched by the
name of a specific supplier or product category. Searchable
categories include animal care, animals, food and water,
housing, husbandry, information resources and manage-
ment, plant, research, services, surgery, and veterinary med-
ical care.
appendix A: glossary of
acronyms and terms
Abbreviation/Word Definition
Ad libitum Continuous access to food and fluid sources
Autochthonous Tumor burden originating within the host animal
Ascites Fluid in the peritoneal cavity
BAR Bright, alert, responsive
BCS Body condition score
BID Twice daily treatment
BT Body temperature
BUN Blood urea nitrogen
BW Body weight
CFA Complete Freund’s adjuvant
CO2 Carbon dioxide
CS Controlled substance
DEA Drug Enforcement Agency; provides registrations
to entities that intend to use any CS in the United
States and its territories
DMSO Dimethyl sulfoxide; solvent with anti-inflammatory
properties, typically used as a drug carrier
EAE Experimental allergic encephalitis
ECG (EKG) Electrocardiogram
Ectopic Site of tumor growth different from the tissue of
origin (i.e., liver tumor cells transplanted under
the renal capsule); opposite of orthotopic
EDTA Ethylenediaminetetraacetic acid
EMLA® Eutectic Mixture of Local Anesthetics (topical
eutectic mixture of 2.5% prilocaine and 2.5%
lidocaine cream)
EOD Every other day
Erythema Reddened skin, may be thickened
ET Endotracheal
(Continued)
55
56 appendix A: glossary of acronyms and terms
Abbreviation/Word Definition
Fasting Food access is removed, yet animals have ad
libitum access to fluid (i.e., water)
FNA Fine-needle aspirate
g Gram (unit of weight)
GY Gray (unit of radiation)
HCT Hematocrit
IACUC Institutional Animal Care and Use Committee
ICU Intensive care unit
ID Intradermal
IM Intramuscular
IP Intraperitoneal
IT Intratracheal
IV Intravenous
LRS Lactated Ringer’s solution
Metastasis Spread of tumor cells from primary site to distant
sites in the body
MUS Mouse urologic syndrome
NOD Nonobese diabetic (model for type 1 diabetes)
NSAID Nonsteroidal anti-inflammatory drug
Orthotopic Anatomically correct site for tumor transplantation
(i.e., liver tumor cells transplanted into the liver);
opposite of ectopic
PE Polyethylene
PO Per os (by mouth)
Restriction (of food/fluid) Total volume of food or fluid is strictly monitored
and controlled
RO Retro-orbital
SC Subcutaneous
Scheduling (of food/fluid) Animal consumes as much food or fluid as desired
at regular intervals
SCID Severe combined immunodeficiency (mutation)
SID Once daily treatment
Syngeneic Tumor cells transplanted between animals of same
inbred strain
TBI Total body irradiation
TBV Total blood volume
UD Ulcerative dermatitis
Ulceration Circumscribed, inflamed, and “open” skin lesion
with death (necrosis) of surrounding tissues
Xenogeneic Tumor cells transplanted between different species
of animals (i.e., human cells transplanted into a
mouse)
appendix B: suggested
medical supplies for
rodent critical care
• Alcohol swabs
• Antiseptics (Betadine [povidone-iodine] swabs, chlorhexidine
solution)
• Bacterial culturettes/blood culture medium
• Blood analyzer (portable hand-held or table-top)
• Blood collection tubes (red top, green top [heparin], purple top
[EDTA])
• Catheters (IV)
• Cotton-tipped applicators (single use, sterile) for topical oint-
ment applications
• Disposable hypodermic needles (23 to 26 gauge for size range)
• Disposable syringes (1 to 3 ml)
• Endotracheal tubes (uncuffed 1.0–2.0 mm for rodents > 100 g)
• Epsom salts (to treat pododermatitis, etc.)
• Feeding needles (for orogastric gavage; 22 gauge, ball tipped)
• Fluids (see Chapter 4)
• Fluorescein stain
• Gauze (4 × 4) sponges
• Glucometer
• Lanolin ointment
57
58 Appendix B: Suggested Medical Supplies
• Meloxicam
• Nail clippers for teeth and nail trimming (rats)
• Nose cones for anesthesia
• Nutritional supplements (see Chapter 4)
• Ophthalmoscope
• Otoscope
• Refractometer
• Scalpel blades/handles
• Scissors for teeth and nail trimming (mice)
• Silver sulfadiazine ointment
• Stethoscope (pediatric)
• Surgery instrument packs
• Surgical draping materials (see Chapter 4)
• Suture with attached needles
• Tape
• Tissue glue
• Topical antibiotic ointment ± steroid
• Tweezers
• Vitamin E ointment
• Warm-water recirculating blankets
appendix C: rodent
formulary
introduction
Selection of appropriate drug and therapeutic regimens requires
careful consideration of multiple factors, including published adverse
effects, to maximize effectiveness and minimize risks. Consideration
of the selected species, the intended procedure, and the practicality
of available agents contributes to the choice of treatments utilized
in a given clinical case. Procedures in animals that may cause more
than slight pain or distress should be performed with appropriate
sedation, analgesia, or anesthesia; one should assume that any pro-
cedures deemed painful to humans are therefore able to cause pain
to animals (Interagency Research Animal Committee [IRAC], 1985).
Pharmaceutical-grade drugs should be used, whenever available, for
animal procedures. These agents are defined in detail by the Office
of Laboratory Animal Welfare (see relevant section in Chapter 5);
as well, expired agents may not be used in any laboratory animals
(National Research Council [NRC], 2011).
Dosing in rodents is typically off label and will vary depend-
ing on age, gender, strain, and condition of the animals. Pregnant
rodents will require special consideration depending on the stage
of pregnancy, whether the agent under consideration crosses the
placenta, and whether potential effects on the fetus will alter experi-
mental data. Avoiding drug problems during therapy of the criti-
cal patient takes preplanning and foresight (Hackett and Lehman,
2005, Meador, 1998). Guidance is available for review of drug inter-
actions, adverse effects, and indications and contraindications for
59
60 appendix C: rodent formulary
multimodal t herapy or “balanced anesthesia” in veterinary patients

(Flecknell, 2001, Plumb, 2005). Balanced anesthesia is defined as
the “administration of a mixture of sedatives, analgesics, and anes-
thetics to produce anesthesia with lower doses than would be nec-
essary if each component were used individually” (He et al., 2010
p. 45). It will be critical to obtain an accurate body weight (BW) for
each animal prior to administration of a calculated drug dose to limit
the potential of adverse effects related to either over-or underdosing.
The following abridged formulary is adapted from numerous
rodent references (Danneman et al., 2012, Gaertner et al., 2008,
I ACUC-UPENN, 2010, Oglesbee, 2011).
induction agents
Induction agents and premedications can calm the patient, smooth
anesthetic induction and recovery, and reduce the dose of anesthetic
agent needed. Preemptive analgesia should be administered with the
induction agents.
Dosage (mg/kg) Route of

Induction Agent Species (Unless Specified) Administration
Atropine Rodents 0.05 SC, IP, IV
Diazepam Rodents 1–3 IP, SC
Isoflurane Mice 4% Inhaled
Neonatal mice 2–4% Inhaled
Rats 5% Inhaled
Midazolam Rodents 0.5–2.0 IP, SC
Propofol Mice 26 IV
Rats 10 IV
IP = intraperitoneal; IV = intravenous; SC = subcutaneous.
anesthetics
Anesthesia should be provided to animals undergoing p rocedures
that cause more than momentary or slight pain or distress.
Anesthetics render the animal unconscious without loss of vital
functions. Inhalant anesthetics provide a reliable and reversible
means of rendering rodents unconscious in order to perform surger-
ies and other intricate or potentially painful procedures. Injectable
a nesthetics may not be as predictable in efficacy between animals;
appendix C: rodent formulary 61
however, they are documented to provide sedation and even anes-

thesia at a surgical plane. For those drugs that are controlled
substances (designated in the formulary tables by CS), and if
used in the United States and its territories, a Drug Enforcement
Administration (DEA) license will be required to obtain these drugs
for use in animals.

Anesthetic Species (Unless Specified) Administration
Chloral hydrate Mice 370–400 IP
Rats 300–450 IP
Rats 400–600 SC
Hypothermia Rodent pups Placed on crushed ice, Contact
(neonatal pups separated by a thin
only) layer to avoid direct
contact with ice
Isoflurane Mice 0.08–1.5% Inhaled
Neonatal mice 0.25–2.5% Inhaled
Rats 0.25–2.5% Inhaled
Ketamine (CS)/ Mice 100 K/5 D IP
diazepam (CS) Rats 40 K/5 D IP
Ketamine (CS)/ Mice 50–75 K/1–10 M IP
midazolam (CS) Rats 60 K/0.4 M IP
Ketamine (CS)/ Mice 90–150 K/7.5–16 X IP
xylazine Rats 40–80 K/5–10 X IM, IP
Ketamine (CS)/ Mice 70–100 K/5–10 X/1–3 A IP
xylazine/ Rats 40 K/8.0 X/4.0 A IM, IP
acepromazine
(necessary for
surgical plane of
anesthesia)
Medetomidine/ Rats 200–300 μg/kg M/300 IP
fentanyl (CS) μg/kg F
Sevoflurane Rats 2–2.4% Inhaled
Sodium Mice 30–90 IP
pentobarbital (CS) Rats 30–60 IP
Tiletamine (CS)/ Rats 20–40 IP
zolazepam (CS)
Thiobarbital Mice 80 IP
(Inactin) (CS)
Tribromoethanol Mice 125–300 IP
(TBE or Avertin)
Tribromoethanol/ Rats 150 T/0.5 M (reversal 2.5 IP
medetomidine mg/kg atipamezole)
CS = controlled substance; IM = intramuscular; IP = intraperitoneal; IV = intravenous;
SC = subcutaneous.
analgesics
Analgesia should be provided to animals undergoing procedures that
cause more than momentary or slight pain or distress. Analgesics
reduce or relieve pain without loss of consciousness. Systemic or
local analgesics may also reduce the anesthetic requirements and
have a preemptive effect on pain perception that persists into the
recovery period. Preemptive, but also immediate postoperative, anal-
gesic administration is important for adequate pain relief in postsur-
gical rodents.
Note: Please see further published considerations about specific anal-
gesics at the end of the formulary tables.
Dosage (mg/kg)
(Unless Route of
Analgesics Species Specified) Administration
Acetaminophen Rodents 100–300 PO, SC, IP
(Tylenol®)
Aspirin (acetyl Rats 50–100 PO
salicylic acid) Mice 50–100 PO
(administer every Rodents 20 SC
4–24 h)
Rodents 100–120 IP
Buprenorphine Mice 0.5–2.0 SC,IP
(Buprenex®) (CS) Rats 0.01–0.10 SC, IP, IM
(administer every May need doses
6–8 h) up to 5.0–10.0
mg/kg if dosed
orally
Butorphanol (CS) Mice 1–5 SC
Rats 1–5 SC
Carprofen Mice (for acute 5 SC
(Rimadyl®) incisional pain)
(administer at Rats 5–15 PO, SC
least every 6–12
h)
Celecoxib Rats 10–20 PO
Clonidine Mice 0.25–0.5 PO
Mice 0.001–0.1 IP
Diclofenac Mice 9.0–28 IP
Dipyrone Rats 50–600 SC, IP, IV
Dipyrone/ Rats 177–600 D/3.1– SC, IV
morphine (CS) 3.2 M
Fentanyl (CS) Mice 0.025–0.6 SC
Rats 0.01–1.0 SC
Rats 2.0–4.0 g/day PO
(Continued)
Dosage (mg/kg)
(Unless Route of
Analgesics Species Specified) Administration
Flunixin Mice 4.0–11 SC
meglumine Rats 2.5 every 12–14 h SC
(Banamine®)
Ibuprofen (Advil®), Mice 40–100 PO, SC
Motrin®, Nuprin®)
Ibuprofen/ Rats 200 I/2.3 H PO, SC
hydrocodone (CS)
Ketoprofen Rats 5–15 every SC, IP
(Ketofen®) 12–24 h
Mice 5 every 24 h SC, IP
Lidocaine Rats 0.67–1.3 mg/ SC-pump
(Xylocaine®) kg/h CRI
Lidocaine/ Mice 0.44 mM L/0.18 Topical
buprenorphine mM
(CS) B in DMSO
Meloxicam Mice 5.0 SC
(Metacam®) Mice 5.0 (oral PO
(administer once suspension)
daily) Rats 2.0 SC
Morphine (CS) Mice 10 SC
Mice 6.1 mM in DMSO Topical
Rats 2.0–10 SC
Rats 2.8 SC-Liposome
Naproxen/ Rats 200 N/1.3 H SC
hydrocodone (CS)
Oxymorphone (CS) Mice 0.2–0.5 SC-Liposome
Rats 0.1 IV
1.2–1.6 SC-Liposome
Tramadol Rats, mice 5–12.5 SC, IP
(administer every
12 h)
CRI = constant rate infusion; CS = controlled substance; DMSO = dimethyl sulfoxide;
IP = intraperitoneal; IV = intravenous; PO = by mouth; SC = subcutaneous.
local and topical anesthetics

Local anesthetics can reduce the perception of pain at the surgi-
cal site; these should be infiltrated around the incision site prior to
recovery of the animal from general anesthesia and preferably prior
to initiation of the incision. In conjunction with other agents, their
use may allow reduced levels of general anesthetics, which may speed
recovery and minimize potential for adverse outcomes.
Local or Topical Dosage (mg/kg) Route of

Anesthetics Species (Unless Specified) Administration
Bupivacaine Rodents Local infiltration up Local SC around
to 5 mg/kg incision site
EMLA® (may take Rodents Application of a layer Topical application
30 min for effect) up to 1-mm thick
Lidocaine Rodents Local infiltration up Local SC around
to 10 mg/kg incision site
Proparacaine Rodents 1–2 drops per eye Topical onto eye
(0.5%) prior to staining
the cornea or prior
to RO sampling
Note: Lidocaine and bupivacaine may be mixed at a 1:1 ratio for subcutaneous
instillation at the incision site.
RO = retro-orbital; SC = subcutaneous.
reversal agents
Reversal of certain drugs leads to early termination of anesthe-
sia, which may reduce adverse events and allow rapid return of the
rodents to the home cage environment. If reversal agents are used,
both the anesthetic and the analgesic properties of the drug may be
terminated; thus, alternative sources of analgesia should be provided.

Reversal Agents Species (Unless Specified) Administration
Yohimbine Mice 0.5–1.0 SC, IP
Atipamezole Rats 0.5 SC
(Antisedan®)
Naloxone Rodents 20 IP
IP = intraperitoneal; SC = subcutaneous.
antibiotics
Antibiotics should be selected based on sensitivity and culture results,
when available. Typically, more common and broad-spectrum drugs
are preferred to begin the treatments prior to culture results.

Antibiotics Species (Unless Specified) Administration
Amoxicillin/clavulanic Mice 12.5–15 every 12 h PO
acid (Clavamox®) Rats 150 every 12 h IM
(Continued)

Antibiotics Species (Unless Specified) Administration
Azithromycin Rodents 10–30 every 24 h PO
Chloramphenicol Mice 30–50 every 8h PO
Enrofloxacin Rodents 5–10 every 12h PO, SC
Dilution in
300-ml water
bottle with 1 mg/
ml drug = 300
mg/300-ml bottle
Gentamicin Rodents 5–10 every 8–12 h SC, IM
Penicillin Rodents 22,000–100,000 SC
IU/kg daily
Polymyxin B sulfate- Rodents Over-the-counter Topical
neomycin sulfate- formulations;
bacitracin zinc apply every
(Neosporin® or similar 12–24 h to cover
antibiotic ointment) the affected area
Trimethoprim-sulfa Rodents 15–30 every 12 h PO in drinking
Dilute in water water
source
IM = intramuscular; PO = by mouth; SC = subcutaneous.
further commentary on specific

anesthetics and analgesics
Multimodal or balanced anesthesia and analgesia (Parker
et al., 2011) are the veterinary standards for procedures that are
invasive, penetrate a major body cavity, or are predicted to result
in m
oderate-to-severe intensity of impact on the animal. The ideal
a nalgesic regimen will manage patient pain without creation of

unwanted side effects or bias to the research model outcomes.
Specific commentary about published concerns and consider-
ations for various analgesic regimens in laboratory rodents are listed
alphabetically, with species designations as indicated.
Acetaminophen
Rats
• Acetaminophen should not be dosed in rats above
300 mg/kg PO due to potential for hepatic necrosis and
impact on research studies (Hausamann et al., 2002).
• Rats should be acclimated to the novel taste prior to actual

administration of flavored suspensions to avoid dramatic
reductions in fluid intake due to neophobic tendencies (Bauer
et al., 2003, Speth et al., 2001).
Bupivacaine
• Bupivacaine may sting on injection and infusion around the
planned incision site; therefore, it should be injected after the
patient is anesthetized. It should provide pain management
at the site of injection for up to 4–6 h.
Buprenorphine
Mice
• Buprenorphine is appropriate for management of acute
incisional pain at doses of 0.5 to 2.0 mg/kg SC (Yamada

et al., 2009).
• This drug can influence behavior (when dosed up to
1.0 mg/kg) and lead to increased spontaneous locomo-
tor a
ctivity, which may adversely affect research outcomes
(Cowan et al., 1977).
• In comparative experiments, when mice were dosed (2.0 mg/
kg SC) before ova implantation surgery (whereby incisions
were made over the flank area with ovary isolation and retrac-
tion) and then dosed twice at 6-h intervals after surgery,
buprenorphine did not offer superior pain relief compared
to one dose of drug given presurgically. Postoperative heart
rate and blood pressure parameters were recorded telemetri-
cally and found to have no significant differences between
the three doses versus one dose. However, those animals
that were given three doses had significant weight loss due
to diminished food consumption, which was deemed to be an
adverse outcome of the study (Goecke et al., 2005).
• Following intraperitoneal surgery under isoflurane, mice
have been shown to better tolerate recovery with the addition
of a line block at the incision site (bupivacaine, lidocaine, etc.).
Buprenorphine can be given intraoperatively at 1.0–2.0 mg/
kg IP, then administered twice daily for day 1 after abdominal
surgery and subsequently replaced with meloxicam at 5 mg/
kg given SID on days 2–3 postprocedure.
• Analgesic efficacy of sustained-release buprenorphine

(Bup-SR) dosed at 1.0 mg/kg has been shown to last at least
12 h (compared to 3–5 h for injectable buprenorphine HCl) in
male BALB/cJ and SWR/J mice (Carbone et al., 2012).
Rats
• Buprenorphine can increase activity when dosed at 0.1–
3.0 mg/kg and lead to abnormal behaviors like repetitive
licking and biting of limbs and biting of aspects of the cag-
ing environment, along with incidents of fighting, at 4 to 5 h
postadministration (Cowan et al., 1977).
• Respiratory depression has been noted in conscious rats
following injections (Cowan et al., 1977).
• Oral dosing of buprenorphine is discouraged in rats and has
been shown only to be effective for 6- to 8-h intervals for
mild-to-moderate pain levels (assessed by hot-water tail-flick
assays) at doses approaching 5 mg/kg PO (Gades et al., 2000,
Martin et al., 2001, Thompson et al., 2004).
• Buprenorphine offered in flavored gelatin is not readily con-
sumed by rats at doses (5.0–10.0 mg/kg) necessary to induce
significant increases in pain threshold, which necessitates
orogastric administration by gavage (Martin et al., 2001).
For rats undergoing flank laparotomy, 0.3 mg/kg in gelatin
provided analgesia and limited postprocedural anorexia and
weight loss (Flecknell, Roughan, et al., 1999).
• In a multimodal regimen, specifically for hypophysectomy
surgery in rats, following anesthesia with pentobarbital
(30–50 mg/kg), animals were provided with buprenorphine
(0.05 mg/kg), carprofen (5 mg/kg SC), and fluid therapy
(30 ml/kg).
• Rats injected subcutaneously with a 1.2-mg/kg sustained-
release formulation (Bup-SR) were tested in thermal nocicep-
tion and surgical postoperative pain models. In both, Bup-SR
showed evidence of analgesia for 2 to 3 days (Foley et al., 2011).
• For management of visceral pain in rats, buprenorphine is
less effective than oxymorphone (Gillingham et al., 2001).
• Buprenorphine administration in rats has been linked to
side effects of weight loss (Brennan et al., 2009) followed by
hyperphagia and weight gain due to pica (Clark et al., 1997,
Thompson et al., 2004).
• Pica, manifesting typically as ingestion of large amounts of

bedding materials within 1–2 days following buprenorphine
administration, has been noted especially in Sprague Dawley
rats. Pica can lead to gastric and esophageal impaction,
may indicate nausea in rats, and is potentially analogous to
emesis in other species (Bender, 1998, Bosgraaf et al., 2004,
Clark et al., 1997, Takeda et al., 1993). Pica can also lead to
esophageal obstruction, or choking, and may require lavage
with hydropropulsion using endoscopy via the mouth to best
attempt to advance the obstruction into the stomach (Ovadia
and Zeiss, 2002).
Carprofen
Carprofen should be considered as an adjunctive therapy to refine
analgesic regimens for rodent surgery and to improve postopera-
tive care (diminish instances of ataxia, bleeding, and weight loss);
to increase survival rates; and to maintain animal welfare (Weiner
et al., 2011).
Mice
• Carprofen is appropriate for management of acute incisional
pain at doses of 5 mg/kg SC every 6 h (Yamada et al., 2009).
• Postlaparotomy, mice can be administered carprofen
(5 mg/kg subcutaneously, twice daily for 3 days) with pro-
phylactic antibiotics, like enrofloxacin (30 mg/kg SC SID for
4 days). Some reports have noted that 5 mg/kg is the mini-
mum dose, and that doses up to 10–20 mg/kg carprofen
may provide a more effective analgesic dose for mice (Clark
et al., 2002).
Rats
• In a multimodal regimen, specifically for hypophysectomy
surgery in rats, following anesthesia with pentobarbital
(30–50 mg/kg), animals were provided with buprenorphine
(0.05 mg/kg), carprofen (5 mg/kg SC), and fluid therapy
(30 ml/kg).
• For rats undergoing laparotomy, carprofen (5 mg/kg SC)
minimized a postoperative reduction in food and water con-
sumption; however, if dosed orally, higher dose rates should
be provided (Flecknell, Roughan, et al., 1999).
Fentanyl
• Transdermal delivery of fentanyl for analgesia has benefits,
including more consistent systemic concentrations, reduced
dosing frequency, and reduced handling stress. The choice of
application site is influenced by the ability of the animal to
remove the patch, difficulty of maintaining skin contact by
the presence of hair or movement of the animal, and interfer-
ence with the medical or surgical procedure being performed.
• The interscapular region is a common application site; how-
ever, drawbacks of this location include the need to shave the
area (which may impair skin integrity) and movement of skin
in the conscious animal.
Hypothermia
Mice
• Neonatal rodents typically are resistant to inhalant anes-
thesia and may best be anesthetized using hypothermia, in
essence by placement of altricial pups on ice, separated from
direct contact by a thin layer of plastic wrap, parafilm, or
paper towel (Phifer and Terry, 1986). Neonates can remain
exposed to the ice for 3–10 min to induce torpor for injections
or sampling.
• Following the procedure, pups should be slowly rewarmed
using a heating source (e.g., incubator ~33°C) or through
manual warmth and gentle stimulation and then returned to
maternal dams for care. With rewarming, pups become active
and responsive within 20–30 min.
Rats
• Anesthesia of neonatal rats (12–14 days old) using hypother-
mia (by placement of pups on a draped ice pack) combined
with inhalant isoflurane anesthesia has been shown to be
more effective for subcutaneous implantation procedures
than anesthesia with injectable agents, like ketamine and
xylazine (dosed 100 and 10 mg/kg, respectively). Rat pups
have been documented to crawl back to the dam and com-
mence suckling after hypothermic anesthesia (Libbin and
Person, 1979); they are readily accepted by the dam with no
long-term side effects noted (Molloy et al., 2004).
Ibuprofen
Mice
• Mice prefer the palatability of oral ibuprofen liquid-gel (at
1 mg/ml) over children’s berry-flavored ibuprofen elixir (at
1 mg/ml) as determined in a study in which mice with var-
ious size wounds were given either of the two nonsteroidal
anti-inflammatory drug (NSAID) options and were further
monitored over a 9-day period. Mice consumed significantly
more of the liquid-gel formulation. In addition, the mice on
liquid-gel consumed twice the amount of food and were more
alert, active, and groomed than when given the elixir formu-
lation (Ezell et al., 2012).
• Commercially available cherry-flavored ibuprofen elixir (at
2 mg/ml concentration) has been shown to promote postsur-
gical recuperation in mice; however, mice may consume this
fluid solution in excess of normal and to the detriment of food
intake (Bosgraaf et al., 2006).
Ketamine Plus Xylazine

Mice
• Body temperature in rodents may decrease by several
degrees following administration of ketamine plus xylazine,
and this decrease may be exacerbated by increased urination,
defecation, and salivation (Wixson and Smiler, 1997, Wixson
et al., 1987).
• Ketamine plus xylazine, combined further with acepromazine
to achieve surgical anesthesia for 45 min, can have substan-
tial cardiovascular effects, manifested by low pulse rates and
hypotension (Buitrago et al., 2008).
• Ketamine plus xylazine offers sedation but does not routinely
enable the animal to reach a surgical plane of anesthesia
(Arras et al., 2001).
• Addition of acepromazine in the anesthetic regimen with ket-
amine can prolong recovery times, as determined by righting
reflex and time to walking (Baker et al., 2011).
Rats
• It has been documented that rats anesthetized with ketamine
plus xylazine may develop ocular lesions, including kera-
toconjunctivitis sicca (Kufoy et al., 1989) and irreversible
orneal lesions, despite perioperative eye lubrication (Turner

c
and Albassam, 2005).
• Profound reductions in rectal and core body temperatures
have been noted in rats, demonstrated by a decrease of up to
4°C over 60 min of anesthesia (Wixson et al., 1987). This side
effect emphasizes the overwhelming need to minimize hypo-
thermia in rodents undergoing anesthesia (Lin et al., 1978,
Simpson, 1997).
Ketoprofen
Rats
• In 2- to 3-month-old female Crl:CD[SD] rats, perioperative
treatment with ketoprofen (5 mg/kg SC) led to marked gastro-
intestinal bleeding, erosions, and small intestinal ulcers, which
worsened in intensity of clinical signs if the drug was coupled
with inhalant isoflurane anesthesia (Shientag et al., 2012).
Oxymorphone
• Oxymorphone has been shown to be a superior analgesic
for visceral pain management over buprenorphine in rats
(Gillingham et al., 2001).
Tramadol
• Tramadol is an approved, opioid-like analgesic; the optimum
dosage and route of administration were determined to be
12.5 mg/kg IP for provision of long-lasting and effective
a nalgesia (Zerge Cannon et al., 2009).
• In comparative studies, tramadol (5 mg/kg SC dosed twice
daily on the day of surgery and 24 h after surgery, then
SID through day 3 postoperatively) has provided superior
pain relief in rat models of endometriosis, as compared to
buprenorphine (Debrue, 2011).
• For incisional models of pain, tramadol alone (at 10 mg/kg prior
to skin incision and 10 mg/kg IP twice daily) does not provide
sufficient analgesia; instead, buprenorphine (0.05 mg/kg SC)
and tramadol plus gabapentin (80 mg/kg) were deemed to be
appropriate (when administered preemptively and for 2 days
postoperatively) (McKeon et al., 2011).
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Life Sciences
for Laboratory
MICE
and RATS
F. Claire Hankenson
For critical care of laboratory rodents, there is a scarcity of sources for comprehen-
sive, feasible, and response-oriented information on clinical interventions specific
to spontaneous and induced models of disease. With the more complex cases that
need critical care management, many treatment approaches to veterinary emergen-
cies cannot be applied directly to the laboratory rodent. The first text of its kind
devoted to the challenges of critical care management for laboratory rodents,
Critical Care Management for Laboratory Mice and Rats provides a special-
ized resource for all veterinary, husbandry, technical, and research professionals
who utilize rodent models for biomedical research.
The book covers the varied approaches to laboratory rodent patient care, health
assessments, characteristics of specific disease models, monitoring and scoring
of disease parameters, and humane interventions. Giving primary consideration
to preservation of animal health and welfare, the text also considers how best to
balance welfare with the achievement of proposed scientific objectives. Organized
into five chapters, this full-color book covers the following topics:
• General Approaches for Critical Care
• Critical Care Management for Laboratory Mice
• Critical Care Management for Laboratory Rats
• Special Considerations for Critical Care Management in Laboratory Rodents
• Resources and Additional Information
The author provides treatment guidelines with the expectation that they will
be applied with apt professional judgment, allowing for further modification
of clinical recommendations for improved patient-based care and welfare
for research animals.
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1 general approaches for critical care.........................................1

2 critical care management for laboratory mice........................25

veterinary care measures......................................................33

Spontaneously Hypertensive Rat Models........................ 148

4 special considerations for critical care management

5 resources and additional information.................................. 199

American Association for Laboratory Animal

Within the field of laboratory animal practice, guidance and

F. Claire Hankenson, DVM, MS, DACLAM

influences on the functionality of the animal’s immune system must

obtaining a clinical history and medical records

• Genetic background, strain, or stock

• Administration of anesthesia or analgesia (type and time

Assessments of the critical patient must be comprehensive, orga-

Species: Sex: Strain: Vendor:

Brief Clinical History:

Problem including differential diagnoses:

Sick Animal Report

***REPORT EMERGENCIES TO THE CLINICAL STAFF IMMEDIATELY***

body condition scoring

relocation for physical examinations

monitoring critically ill rodents

• Blood or blood-stained discharges from any orifice

Table 1.1: Diagnostic Differentials Based on Clinical Conditions

Table 1.2: Rodent Health A bnormalities, with Emergencies

In a critical care situation, the initial goal is to keep the rodent

• Is the patient conscious?

• Auscultation can be performed using a pediatric stethoscope.

pain recognition and assessments

Table 1.3: Categorization of Painful Procedures Performed in the

pain may be challenging; in particular, one must be aware that each

supportive care for surgical procedures

• Depth of anesthesia: Assess by direct visualization, gauging

Table 1.4: Template A ssessment for Pain E valuation in Rodents

Table 1.4: (Continued) Template A ssessment for Pain E valuation in

to stimulation (toe pinch); note that the palpebral response is

Fig. 1.3 (A) An anesthetized adult mouse in preparation for a pro-

Pre-, peri-, and postoperative supportive care should include per-

Following surgery, it is imperative to manage potential pain and

Analgesics may be necessary at scheduled intervals for several

supplemental heat provision

can be placed near animals to provide thermal support. Care should

Routes of Drug and Fluid Administration

Fig. 1.4 Differing routes of administration of substances, including

• Subcutaneous (SC). This is one of the most common sites for

to successfully place and secure (to prevent chewing by the

• Intramuscular (IM). Intramuscular injections of bolus ­ fluids

Alternative drug routes for mice and rats include intratracheal

REPORT EMERGENCIES TO THE CLINICAL STAFF IMMEDIATELY

• Intramuscular (IM). Intramuscular injections of bolus fluids

Physiological aspects, like body weight, activity, and b ehavior

Fine-needle aspiration can be performed to d etermine if the

Fig. 2.14 An example of how a flooded mouse cage appears following a