Essentials For Animal Research:
A Primer For Research Personnel
By
B.T. Bennet
M.J. Brown and
J.C. Schofield
Beltsville, Maryland
National Agricultural Library
April 1990
Chapter 1. . . . . . . . . . . . Regulations and Requirements
by B. Taylor Bennett
Chapter 2. . . . . . . . . . . . . . Alternative Methodologies
by B. Taylor Bennett
Chapter 3. . . . . . . . . . . . . Animal Care and Use: A
Experimental Variable
by John C. Schofield and
Marilyn J. Brown
Chapter 4. . . . . . . . . . . . Principles of Anesthesia and
Analgesia
by Marilyn J. Brown
Chapter 5. . . . . . . . . . . . . Principles of Aseptic
Technique
by John C. Schofield
Chapter 6. . . . . . . . . . . . . . . Perioperative Care
by Marilyn J. Brown and
John C. Schofield
Chapter 7. . . . . . . . . . . . . . . . Euthanasia
by B. Taylor Bennett
Chapter 8. . . . . . . . . . . . . . . The Animal Welfare
Information Center
by Jean A. Larson,
Kevin P. Engler and
B. Taylor Bennett
Chapter 9. . . . . . . . . . . . Organizations, Associations
and Societies
by Marilyn J. Brown,
John C. Schofield and
B. Taylor Bennett
Chapter 10 . . . . . . . . . . . . . General References
by John C. Schofield,
Marilyn J. Brown and
B. Taylor Bennett
INTRODUCTION
This manual was developed from the outlines of a course
entitled Essentials for Animal Research, originally developed at
the University of Illinois at Chicago for graduate students who
wanted to learn more about the use of animals in research than
generally covered in the training received in their chosen area
of concentration. From its inception, the course has constantly
evolved to remain current with everchanging regulations and an
increasing awareness by graduate students of the issues
concerning the use of animals in biomedical research, teaching
and testing. The course introduces those elements which have
become essential requirements for using animals in research,
teaching or testing programs. These requirements primarily center
around the responsibilities one assumes when they intend to use
animals in their work. The ultimate responsibility lies with the
Principle Investigator who must have a working knowledge of the
regulations, be familiar with the factors that affect the
selection, acquisition and maintenance of experimental animals
and be aware of the ethical and social issues involved with the
use of animals in biomedical research.
The goals and objectives established for developing the
class lectures are applicable to the material presented in this
manual. With these goals in mind, the authors developed the ten
chapters included in this manual. Remember it was not the
authors' intentions to present an exhaustive treatise on key
elements essential for conducting animal research in a manner
which assures individual and institutional compliance with
pertinent regulatory requirements, but rather an introduction to
the subject matter in a manner which will hopefully encourage
additional reading where appropriate.
In writing this manual it was the author's intent to provide
the reader with:
An appreciation and basic understanding of the regulatory
process and the means by which compliance can be assured. An
overview of those factors which can affect the selection,
acquisition and maintenance of animals used in biomedical
research.
An understanding of the basic principles of controlling
pain and distress, preventing intraoperative infection and
assuring a humane death in the animals used.
An awareness of the responsibilities that one assumes when
choosing to use laboratory animals. These responsibilities would
include, but not be limited to, those which involve an obligation
to the institution, regulatory and funding agencies, the public
and the animals.
The manual has been organized into ten chapters, the first
seven are intended to cover the specific objectives described
above. The last three chapters contain resource information on
the Animal Welfare Information Center of the National
Agricultural Library, a list of organizations from which
additional information can be obtained and a list of general
references covering topics of interest to the investigator who
utilizes animals in research, teaching and testing programs.
The authors would like to thank Drs. James Harwell, Louis
Ramazzotto and Richard Simmonds for their input and support
during the review process of this project. To the staff of the
AWIC, we send our appreciation for their enthusiastic support
throughout the project and for their assistance in the final
stages of transferring the electronic version of the manual to
the library.
This manual was produced as joint effort of the USDA
National Library of Agriculture and the University of Illinois at
Chicago and supported by cooperative agreement number
58-32U4-7-070.
Chapter 1
Regulations and Requirements
B. Taylor Bennett, D.V.M., Ph.D.
INTRODUCTION
Since the ultimate responsibility for compliance with
regulations that affect the care and use of animals lies with the
investigator, it is important that he/she have a working
knowledge of the basic regulatory requirements. In this manual,
the types of regulations will be discussed under two broad
general headings:
1. Involuntary
2. Voluntary
Involuntary regulations can be defined as those required by
law or set forth as a condition of funding. There are four types
of regulatory controls which can be considered as involuntary:
1. The Animal Welfare Act (AWA)
2. The Public Health Service Policy
3. The Good Laboratory Practices Act
4. The Requirements of Private Funding Agencies
Voluntary regulations can be defined as those that an
individual or institution adheres to as part of their overall
commitment to research and academic excellence. There are two
types of regulatory controls which can be considered as
voluntary:
1. Accreditation by the American Association for the
Accreditation of Laboratory Animal Care (AAALAC)
2. Requirements of Individual Users
INVOLUNTARY REGULATIONS
Animal Welfare Act
The Animal Welfare Act was first passed August 24, 1966, as
PL-89-544. It was entitled the "Laboratory Animal Welfare Act"
and authorized, "The Secretary of Agriculture to promulgate such
rules and regulations, and orders as he may deem necessary to
effectuate the purposes of this Act." The purposes of the
original act were to:
1. Protect the owners of dogs and cats from theft of such
pets.
2. Prevent the sale or use of dogs and cats which had been
stolen.
3. Insure that certain animals intended for use in
research facilities were provided humane care and treatment.
In charging the Secretary, Congress specifically prohibited
the promulgation of rules, regulations, or orders which would
interfere with the conduct of actual research. Determination of
what constituted actual research was left to the discretion of
the research facility.
The original Act covered non-human primates, guinea pigs,
hamsters, rabbits, dogs and cats. Humane treatment was required
while they were at the dealers or research facility and while
being transported by dealers. Dealers were required to be
licensed. Research facilities which used, or intended to use,
dogs or cats and either purchased them in commerce or received
any federal funds were required to be registered.
The Secretary also established regulations and standards for
the implementation of unannounced facility inspections and for
the maintenance of specific records by dealers and research
institutions. Responsibility for administering the Act was
delegated within the United States Department of Agriculture
(USDA) to the Administrator of the Animal and Plant Health
Inspection Service (APHIS). Enforcement duties are the
responsibility of the APHIS Deputy Administrator for Regulatory
Enforcement and Animal Care (REAC). The actual inspections are
conducted by 42 Veterinary Medical Officers working under one of
the five REAC Sector Supervisors. The Sector offices are located
in Fort Worth, Texas, Tampa, Florida, Annapolis, Maryland,
Minneapolis, Minnesota and Sacramento, California.
In 1970 the original Act was amended (PL- 91-579) and
renamed the Animal Welfare Act. The amended Act covered broader
classes of animals and included those used in exhibitions and
sold at auction and regulated anyone involved in these
activities. The definition of an animal was expanded to include
all warmblooded animals. The definition of a research facility
was expanded to include those institutions using covered live
animals and not just dogs and cats. These facilities were
required to file an annual report. Civil penalties were also
added for refusing to obey a valid cease and desist order from
the Secretary. The term "handling" was added to the basic
categories for which standards were to be created and the
phrase "adequate veterinary care" was broadened to include the
appropriate use of anesthetics, analgesics and tranquilizers.
The intent of the original Act to prohibit interference with
research was clarified and the Secretary was enjoined from
directly or indirectly interfering with, or harassing in any
manner, research facilities during the conduct of actual research
or experimentation. The determination of when actual research
was being done was still left to the discretion of the research
facility itself.
In 1976, the Animal Welfare Act was further amended to
enlarge and redefine the regulation of animals during
transportation and to combat the use of animals for fighting.
Essentially the Act was broadened to include all forms of
commercial transportation of animals and required all carriers
and intermediate handlers who were not required to be licensed
under the Act to register with the USDA. It also expanded the
definition of a dealer and extended the record keeping
requirements to carriers and intermediate handlers.
In 1976, the Secretary also promulgated regulations which
specifically excluded rats, mice, birds, horses and farm animals
from the definition of an animal. This exclusionary language
effectively excludes over 80 percent of the animals currently
used in research, teaching and testing from coverage under the
Animal Welfare Act.
In 1985 the Act was further amended with the passage of the
Food Security Act of 1985 (PL-99-198) which contained an
amendment entitled the "Improved Standards for Laboratory Animals
Act." This amendment when fully implemented will strengthen the
standards for providing laboratory animal care, increase
enforcement of the Act, provide for collection and dissemination
of information to reduce unintended duplication of experiments
using animals and mandate training for those who handle animals.
The most recent amendment to the AWA also includes
development of standards: for the "exercise of dogs," for
"provision of a physical environment which promotes the
psychological well-being of primates," for limitation of multiple
survival surgeries, and to require the investigator to consult
with a veterinarian in the design of experiments which have the
potential for causing pain to insure the proper use of
anesthetics, analgesics and tranquilizers. Each research
facility will have to show upon inspection, and include in their
annual report, assurances that professionally acceptable
standards for the care, treatment and use of animals are
being used during the actual research or experimentation. As
part of these standards, the investigator is required to consider
alternative techniques to those which might cause pain or
distress in the experimental animals.
The 1985 amendment requires the Chief Executive Officer of
each research facility to appoint an Institutional Animal
Committee consisting of at least three members including a doctor
of veterinary medicine and one member who is not affiliated with
the institution. The regulations promulgated to implement the
amendment designate this committee as the Institutional Animal
Care and Use Committee (IACUC) and charge it to act as an agent
of the research facility in assuring compliance with the Act.
The Committee is required to inspect all animal facilities and
study areas at least once every six months, and to review the
condition of the animals and the practices involving pain to the
animals to insure compliance with the regulations and standards
promulgated under the Act. The Committee is also required to
review once every six months the research facility's program to
assure that the care and use of the animals conforms with the
regulations and standards.
The Committee must file a report of its inspection with the
Institutional official of the research facility. If significant
deficiencies or deviations are not corrected in accordance with
the specific plan approved by the Committee, the USDA and any
Federal funding agencies must be notified in writing.
The Committee must also review and approve all proposed
activities involving the care and use of animals in research,
testing or teaching procedures and all subsequent significant
changes of ongoing activities. As part of this review, the
Committee must evaluate procedures which minimize discomfort,
distress and pain and that when an activity is likely to cause
pain that a veterinarian has been consulted in planning for the
administration of anesthetics, analgesics and tranquilizers and
that paralytic agents are not employed except in the anesthetized
animal. The IACUC must also determine that animals which
experience severe or chronic pain are euthanized consistent with
the design of study, that the living conditions meet the species
needs, that necessary medical care will be provided, that all
procedures will be performed by qualified individuals, that
survival surgery will be performed aseptically and that no animal
will undergo more than one operative procedure that is not
justified and approved. Methods of euthanasia must be consistent
with the definition contained in the regulations.
The IACUC must also assure on behalf of the research
facility that the principal investigator considered alternatives
to painful procedure and that the work being proposed does not
unnecessarily duplicate previous experiments. To provide this
assurance the Committee must review the written narrative
description provided by the investigator. This description must
include the methods and sources used in determining that
alternatives were not available.
In reviewing proposed activities and modifications, the
IACUC can grant exceptions to the regulations and standards, if
they have been justified in writing by the principal
investigator.
In addition to the above requirements, the research facility
is required to provide training in the following areas to
scientists, animal technicians and other personnel involved with
animal care and treatment:
1. Humane practice of animal maintenance and
experimentation.
2. Research or testing methods that minimize or eliminate
the use of animals or limit pain or distress.
3. Utilization of the information service of the National
Agricultural Library.
4. Methods whereby deficiencies in animal care and treatment
should be reported.
The regulations require that each research facility establish
a program of adequate veterinary care that includes: appropriate
facilities, personnel and equipment; methods to control, diagnose
and treat diseases; daily observation and provision of care;
guidance to personnel on the use of anesthetic, analgesic and
euthanasia procedures and pre and post-procedural care. Specific
requirements for maintaining records and filing annual reports
are included in the regulations along with a miscellaneous
section containing a variety of requirements to which a research
facility must adhere.
Public Health Service Policy
The Public Health Service Policy on Humane Care and Use of
Laboratory Animals can be found in Chapter 4206 of the NIH Manual
and Chapter 1-43 of the PHS Manual. The NIH originally initiated
the Policy in 1971. It was extended to all PHS activities
January 1, 1979, and was revised in the spring of 1985 with
implementation to be effective January 1, 1986. With the passage
of the Health Research Extension Act of 1985 (PL-99-158), the
Policy was further revised and the Director of the NIH was
required by law to establish guidelines which heretofore had only
been a matter of PHS policy. An additional revision was released
in September 1986 which reflected the changes required by this
Act.
Under the PHS policy, each institution using animals in
PHS-sponsored projects must provide acceptable written assurance
of its compliance with the Policy. In this Letter of Assurance
the institutions must describe:
1. The Institutional Program for the Care and Use of
Animals.
2. The Institutional Status.
3. The Institutional Animal Care and Use Committee (IACUC).
The Institutional Program must include a list of every branch
and major component, the lines of authority for administering the
program; the qualifications, authority and responsibility of the
veterinarian(s), the membership of the Institutional Animal Care
and Use Committee and the procedures which they follow must be
stated. The employee health program must be described for those
who have frequent animal contact. A training or instruction
program in the humane practices of animal care and use must be
available to scientists, animal technicians and other personnel
involved in animal care, treatment and use. The gross square
footage, average daily census and annual usage of each animal
facility must be listed.
The Institutional Status must be stated as either Category one
(1) (AAALAC accredited) or Category two (2) (nonaccredited).
Institutions in Category two (2) must establish a reasonable plan
with a specific timetable for correcting any departures from the
recommendations in the Guide for the Care and Use of Laboratory
Animals (86-23).
The IACUC must be appointed by the Chief Executive Officer and
consist of at least five members; one of whom is a veterinarian
with program responsibility, a practicing scientist, an
individual whose expertise is in a non-biological science and an
individual who is not affiliated with the institution. This
Committee must use the Guide to review the animal facilities and
the institutional program for humane care and use of animals at
least once every six months and prepare reports of these
evaluations for the responsible institutional official. The
Committee must review and approve animal-related components of
proposals and significant modifications made in ongoing
activities involving the care and use of animals. The Committee
is responsible for reviewing concerns involving the care and use
of animals and making recommendations to the institutional
official regarding any aspect of the animal program, the
facilities, or the personnel training. They are also
authorized to suspend activity involving the care and use of
animals as set forth in the PHS Policy.
In reviewing the animal care and use component of a proposal,
the IACUC must confirm that the project will be conducted in
accordance with the AWA and consistent with the recommendations
in the Guide. In addition, all procedures are reviewed to assure
that pain or distress will be minimized and that (when necessary)
appropriate anesthetics, analgesics and tranquilizers will be
used. The living conditions and medical care available must be
appropriate for the species used, and personnel conducting the
procedures must be appropriately trained and qualified. Methods
of euthanasia should be consistent with the recommendations of
the American Veterinary Medical Association Panel on Euthanasia.
The investigator is responsible for completing a proposal in
accordance with recommendations in the PHS Policy and the
instructions contained in the PHS 398 application packet. As of
October 1988, the instructions for completing 398 can be found in
two locations within the application package. On page 6 the
research investigator's responsibilities for assuring the humane
care and use of animals are clearly addressed. Detailed
instructions for completing Section F of the Research Plan which
describes the use of Vertebrate Animals can be found on page 21.
The institution is responsible for maintaining all the
necessary records to document compliance with the PHS Policy and
for filing annual reports which detail any changes in the program
and indicate the dates of the semi-annual inspections and
programmatic reviews.
The PHS Policy described above is intended to implement and
supplement the "U.S. Government Principles for the Utilization
and Care of Vertebrate Animals in Testing, Research and
Training." The nine principles are published in the PHS Policy
and in the Appendix of the Guide. All those responsible for the
design, supervision and review of the animal care and use
component of a proposal should be familiar with this document.
Good Laboratory Practices
In 1978 the Food and Drug Administration adopted the Good
Laboratory Practices rules which applied to all regulated parties
who conduct nonclinical safety assessment studies. The rules
require the creation of Standard Operating Procedures for all
aspects of the study including animal care and use. A Quality
Assurance Unit must be established to conduct internal inspection
of practices and records to insure compliance with established
policies and procedures. In general the recommendations
contained in the Guide would suffice in terms of animal care when
adherence is properly documented.
Private Funding Agencies
In recent years the requirements of many private funding
agencies which fund research projects involving the care and use
of laboratory animals have changed. It is important to obtain
the requirements from the agency before spending time preparing a
proposal. Some of these agencies not only require review of the
proposal by the IACUC, but require proof of accreditation by
AAALAC. In many instances, the proposals must be reviewed and
approved prior to submission.
VOLUNTARY
American Association for the Accreditation of
Laboratory Animal Care (AAALAC)
AAALAC was originally chartered April 30, 1965, as a voluntary
organization that accredited institutional programs of animal
care and use. AAALAC is governed by a Board of Trustees composed
of representatives of 32 professional organizations. An
18-member Board-appointed Council on Accreditation makes
recommendations based on the results of site visits to evaluate
an institution's compliance with the recommendations contained in
the Guide. This is a peer review process in which standards are
being continually upgraded to reflect current knowledge in
laboratory animal medicine and science. In its accreditation
program the AAALAC Council uses the Guide more as a compilation
of regulatory "standards" and not as a set of "recommendations."
Since the AAALAC accreditation program and the Guide are so
closely linked, a brief review of the Guide's history and its
current contents are warranted. In 1963 the first Guide for
Laboratory Animal Facilities and Care was published by the
Institute for Laboratory Animal Resources (ILAR) under a contract
from NIH. Since its original release the Guide has been revised
in 1965, 1968, 1972 (when the title was changed to the Guide for
the Care and Use of Laboratory Animals, 1978 and 1985. In the
most recent revision, the organization of the chapters was
changed to reflect the increasing role and responsibility of the
institutional program in establishing acceptable standards for
the care and use of laboratory animals. The first chapter is now
Institutional Policies. The remaining four chapters are
Laboratory Animal Husbandry, Veterinary Care, Physical Plant and
Special Considerations. Prior to an AAALAC site visit, each
institution is required to prepare a description of the
institutional facilities and programs using the AAALAC Outline
for Description of The Institutional Animal Care and Use Program,
which follows the Guide's chapter headings.
Once accredited, an institution must submit an annual report
describing changes in the program and facilities and documenting
the annual usage of animals. Site visits occur at least every
three years and these visits consist of an inspection and review
of policies, procedures and facilities which comprise the animal
care and use programs inclusive of selected animal usage areas.
Should deficiencies be identified in a previously accredited
program, the institution is either granted a probationary period
in which to make specified changes, or if the deficiencies are
major, accreditation could be withdrawn.
Individual Users
The instructions for completing PHS 398 clearly define the
roles and responsibilities of the investigator in assuring proper
care and usage of laboratory animals. In addition to this
requirement, it should be understood that any type of care or use
of an animal which results in the creation of nonexperimental
variables can potentially compromise the integrity of an entire
project. As part of their commitment to scientific excellence,
the
users should provide the impetus for setting and maintaining high
standards for the care and use of laboratory animals within their
individual and collective institutions. Failure to do so invites
increased internal and external regulatory requirements which can
drain limited institutional research resources. Good animal care
is good science; the practice of good science should be the
primary goal of all who have chosen careers in the scientific
community.
SUMMARY
In summary, the regulations that affect the use of animals in
research, teaching and testing programs are numerous. A working
knowledge of the applicable regulations is necessary if the
principal investigator is to insure that proposals for funding
contain the necessary information and to assure that the conduct
of all research proposals is in compliance with the requirements
of the regulatory and funding agencies. While the ultimate
responsibility for compliance rests with the principal
investigator, institutional policies should be designed to
provide those responsible for compliance with the necessary
resources to do so.
REFERENCES
Application for Public Health Service Grant, PHS, 398.
Revised October, 1988. OMB No. 0925-001.
Animal Welfare Act (Title 7 U.S.C. 2131-2156), as amended by
PL-99-198, December 12, 1986.
Guide for the Care and Use of Laboratory Animals, NIH
Publication No. 86-23.
Public Health Service Policy on Humane Care and Use of
Laboratory Animals. Revised as of September 1986.
Non-Clinical Laboratory Studies. Good Laboratory Practice
Regulations. Register, December 22, l978, Part II, pp.
59986-60026.
Public Law 99-198. Code of Federal Regulations, Title 9,
Subchapter A, Animal Welfare l989.
Townes, J. Federal Regulations an Overview, Lab Animal,
July-August (9)4:l6-22 (1980).
Chapter 2
Alternative Methodologies
B. Taylor Bennett, D.V.M., Ph.D.
INTRODUCTION
In the regulations promulgated to implement the Animal Welfare
Act as amended in 1985, the research facility must provide
assurances that the principal investigators considered
alternatives techniques to painful procedures and to provide
guidance concerning research and testing methods that limit the
use of animals or minimize the animals' distress. In this
chapter the reader will be introduced to the classical concept of
alternatives with a brief discussion of each major category
including a limited number of examples. For more indepth
coverage of the subject, the reader is encouraged to obtain the
latest bibliography on alternative techniques available from the
Animal Welfare Information Center of the National Agricultural
Library (see Chapter 8).
In recent years the term alternative techniques has come into
common usage in the current controversy involving the use of
animals in research, teaching and testing. It is a term that has
different meanings to different people and this difference
largely depends on which side of the issue one is found. To many
biomedical researchers, alternative techniques refer to those
which can be used in addition to the more traditional animal
models. These techniques can focus on specific biological
functions and in many cases reduce the numbers of animals used.
Therefore these
methods are an adjunct to the more commonly used animal models.
To
the so-called abolitionist who seeks the immediate end to all
animal research, teaching and testing, the term alternative
refers
to those techniques which can entirely replace the use of
animals.
The dictionary, defines alternative as: "offering or expressing a
choice." The dictionary also defines technique as "a method of
accomplishing a desired aim." By combining these definitions,
the
term alternative technique becomes "one which offers a choice in
accomplishing a desired aim."
In designing an experiment which involves the use of animals
to confirm or refute a theory, one should consider all the
possible
techniques that could be used to gather the necessary data. From
this review, choose the method which offers the best chance of
generating the necessary information in the most economical
manner. Economy, in this context, refers to time, actual cost
and the number of animals used. By considering the choices that
are available for accomplishing the desired aim of the experiment
and choosing the one that offers the best chance for success, one
has met the requirements of this literal definition of
alternative techniques.
Since a literal definition provides a rather simplistic
approach to dealing with our responsibility for reducing the
potential pain and suffering of animals that must be used, it is
necessary to develop a working definition of the term. In Dr.
Rowan's book, Of Mice, Models and Men, he defines the term
alternatives to refer to those techniques or methods that replace
the use of laboratory animals altogether, reduce the numbers of
animals required, or refine an existing procedure or technique so
as to minimize the level of stress endured by the animal. Since
stress can be difficult to describe and quantitate, for the
purpose of this manual it will be replaced by the term distress.
The working definition of alternative techniques thus evolves to
"those techniques which replace the actual use of animals, reduce
the numbers used, and/or refine the techniques to minimize the
potential for the animal to experience pain or distress."
This concept of the 3 R's is not new. It first appeared in a
book by Russell and Burch published in 1959 entitled The
Principles of Humane Experimental Technique. In the original
work, the authors defined the 3 R's as follows:
"Replacement means the substitution for conscious living higher
animals of insentient material. Reduction means reduction in the
numbers of animals used to obtain information of given amount and
precision. Refinement means any decrease in the incidence or
severity of in-humane procedures applied to those animals which
still have to be used."
In this text the authors included nonrecovery techniques in
anesthetized animals, as well as tissue culture, as replacement
methods. Reduction included statistical techniques which were
designed to reduce the actual numbers needed in the study. The
use of better animals was also encouraged as a means of reducing
actual numbers used. Refinement referred to techniques that
reduced the potential for pain and distress. This approach still
holds today. It is the principles of Replacement, Reduction and
Refinement that will be covered in this chapter. To attempt to
address these issues for all the uses of animals that fall under
the general rubric of research, teaching and testing is far
beyond the scope of this manual. Therefore the comments that
follow will address only broad issues with some specific examples
for the purpose of clarification.
Prior to discussing the replacement of animals with non-animal
models, the word animal must be defined. On the surface this
appears an easy task. Common sense would tell us that an animal
is one of the two major kingdoms of living organisms. The
dictionary defines an animal as "any of a kingdom of living
beings typically differing from plants in capacity for
spontaneous movement and rapid motor response to stimulation."
In the Definition of Terms promulgated to implement the amended
Animal Welfare Act an animal is defined as:
"any live or dead dog, cat, nonhuman primate, guinea pig,
hamster, rabbit, or any other warm blooded animal, which is being
used or is intended for use for research, testing,
experimentation, or exhibition purposes or as a pet. This term
excludes: birds, rats of the genus Rattus and mice of genus Mus
bred for use in research, and horses and other farm animals such
as but not limited to livestock or poultry used or intended for
use as food or fiber, or livestock, or poultry used or intended
for use for improving animal nutrition, breeding, management, or
production efficiency, or for improving the quality of food and
fiber."
The PHS Policy defines an animal as "Any live, vertebrate
animal used or intended for use in research, research training,
experimentation, or biological testing or for related purposes."
On the other hand the Guide defines an animal as "any warm
blooded vertebrate animal." For the purposes of this manual, and
to be consistent with most approaches to discussing alternative
techniques, an animal will be any living vertebrate, with the
caveat that any model system which moves down the phylogenetic
scale from the generally acceptable animal model will be
considered an alternative.
REPLACEMENT
Alternatives which replace animal models can be classified
into the following broad general categories:
Use of Living Systems
Use of Nonliving Systems
Use of Computer Simulation
Use of Living Systems
In Vitro Techniques - The most commonly recognized nonanimal
living systems are those which fall into the broad category of in
vitro methods such as organ, tissue and cell culture. These
techniques afford the investigator the greatest control of the
"test subject's" environment. Since these systems will not work
when the incorrect combination of atmosphere, humidity,
temperature, pH and nutrients are provided, they tend to minimize
the effects that nonexperimental variables can have on the final
outcome of a study.
Generally, when suboptimal environments are provided for an in
vitro system, the problem becomes one of loss of all experimental
results and not just the production of compromised results. The
most commonly used of the in vitro methods are cell culture
techniques for monoclonal antibody production, virus vaccine
production, vaccine potency testing, screening for the cytopathic
effects of various compounds and studying the function and make
up of cell membranes. The potential uses of in vitro techniques
are almost limitless and will continue to expand as more is
learned about the various organs and their component tissues and
cells, and as the technology of maintaining in vitro environments
improves.
Invertebrate Animals - Invertebrates are another type of living
system which can be used to replace the more commonly used
laboratory animals. Over 90 percent of the animal species thus
far identified are invertebrates. An invertebrate which has long
been used in biomedical research is the fruit fly, Drosophila
melanogaster -- a classic model for the study of genetics. This
species also can be used for detecting mutagenicity,
teratogenicity and reproductive toxicity. The marine
invertebrates represent different species which have not been
widely investigated. However in neurobiology a number of
different marine species have been well characterized and used to
study the physiology of the nervous system.
Micro-Organisms - The micro-organisms represent a third living
system which has been used to replace traditional animal models.
The Ames mutagenicity/carcinogenicity test uses Salmonella-
typhimurium cultures to screen compounds that formerly required
the use of animals. Such systems allow for an almost limitless
number of compounds to be tested which can create an interesting
dilemma. The more compounds that can undergo screening, the more
compounds that will be potentially available to test in animals.
Alternative techniques can replace the number of animals at a
given step in the screening process. However, use of
alternatives may increase the number of compounds that must be
finally tested in intact animals.
Plants - Plants offer another alternative living system which can
be used to replace animals in studies of basic molecular
mechanisms. There is very little morphological and functional
difference between the organelles isolated from plants and those
isolated from animals. The rigid cell wall of plants, however
limits their applicability for use as undisrupted cells.
Use of Nonliving Systems
Chemical Techniques - The most widely used nonliving model system
involves the use of modern chemical techniques. This is
particularly true of the analytical techniques which can be used
to identify substances and to determine their concentration or
potency. Immunochemical techniques use the binding capacity of
highly specific antibodies to seek out minute quantities of
antigen. A classical example of this technique can be
demonstrated by the currently used techniques for identifying
bacterial toxins. Toxin identification previously required the
injection of as many as several hundred mice with supernatant
from cultures of suspected contaminating bacteria. These new
antibody techniques save animals and speed up confirmation of a
tentative diagnosis. By adding a color marker to the Enzyme
Linked Immunosorbent Assay system (ELISA), the whole process
becomes a commercially available test kit such as those used in
home pregnancy detection. A test that previously required the
use of a rabbit now can be performed using an over-the-counter
test kit. There are a variety of chemical techniques that can be
used to determine the presence of a particular chemical reaction
or the presence of an enzyme necessary for a specific reaction.
At the most basic level, the identification of a particular
chemical structure in a compound can provide a great deal of
insight into the potential reactivity and thus the resulting
toxicity of a given substance.
Physical and/or Mechanical Systems - The use of physical and/or
mechanical systems to replace living animals of even the highest
order has application in teaching specific skills and/or
reactions to a well defined set of predetermined circumstances.
The use of computer-linked mannequins in teaching basic
principles of medicine and applied techniques can be best
illustrated by the mannequins used to train people in
cardiopulmonary resuscitation.
Historical data can be used for analyses in a variety of
databases commonly used in the field of epidemiology. However,
while the body of potentially useful information that already
exists in a variety of sources is immense, it may not always be
in a format which permits ready accessibility for evaluation.
For this reason, retrospective epidemiological studies are often
the subject of fairly heated debates. Yet with the increasing
access to historical data available on existing computer
programs, this problem may to a large extent be overcome in the
future.
Use of Computer Simulation
The standout in the alternative techniques controversy is the
claim made for computer simulation as a means of virtually
replacing the use of living animals. In order for a biological
phenomena to be adapted to a computer model, the basic processes
must be expressed in a mathematical formula. Once a formula is
developed then an enormous number of variables can be introduced
and swiftly processed. The key element for success is the
generation of a program from the mathematical formula. The more
complete the formula, the more useful the program. The problem
is that many of the questions being asked of an animal model are
not defined well enough to develop the necessary mathematical
model. As the core knowledge of the biological processes expands
so will the opportunities to use computer simulation to replace
the number of live animals being used.
REDUCTION
In discussing the ways to reduce the numbers of animals used,
the definition of an animal and the principle of moving down the
phylogentic scale must also be kept in mind. The four broad
categories for reducing the number of animals used are:
Animal Sharing
Improved Statistical Design
Phylogenetic Reduction
Better Quality Animals
Animal Sharing
Sharing of animals can significantly reduce the number of
animals used within a given institution. Between institutions,
sharing is more difficult, but can be effective as demonstrated
with the Primate Supply Information Clearinghouse, Regional
Primate Research Center (SJ-50) University of Washington,
Seattle, WA 98195. This service has reduced the total number of
primates used by helping to optimize the usage of those already
in facilities throughout the country.
Sharing can be as simple as allowing someone to practice a
surgical approach on an animal that has been, or is to be
euthanized for other purposes, or providing organs or tissues at
the time of necropsy. Sharing becomes more complicated when
attempting to maximize the use of control animals, but it can
significantly reduce the number used within an institution. If
two studies involve the need to perform a sham operation, the
administration of compounds by identical routes, the use of
standard control diets or the need to condition animals to a
particular environment, control animals could be shared within
the institution. Animal sharing would require some form of
centralized clearing process within the Institutional Animal Care
Program to match the needs of the various investigators and their
studies.
Improved Statistical Design
Anyone who has ever taken a course in experimental design or
applied statistics has been bombarded with the importance of
consulting with the statistician during the design phase of the
experiment and not when the data collected needs to be analyzed.
Improper design of experimental protocols and/or the failure to
use appropriate statistical methods can result in the usage of an
inappropriate number of experimental animals. A variety of
design strategies are available which can reduce the number of
animals needed in a given study. Experimental protocols which
utilize serial sacrifice, group sequential testing and crossover
designs can significantly reduce the numbers of animals required.
The availability of low cost statistical packages for almost
every computer on the market permits more and more investigators
access to sophisticated data management and analysis. This
accessibility makes possible the use of design criteria and
complicated statistical analysis which heretofore have been
largely confined to institutions with large statistical support
units. With this ability at their finger tips, investigators
should be able to maximize the analysis of the data generated
from each animal used, thus reducing the total numbers of animals
necessary for a particular set of data.
Phylogenetic Reduction
Projects which can be designed to use one of the myriad of
invertebrate species instead of a non-human primate species
represent a type of phylogenetic reduction which was discussed as
a replacement technique. Such broad jumps across the phylogenetic
scale are not always possible, but less dramatic shifts can
significantly reduce the numbers of higher species being used in
research, teaching and testing. In many instances, the theory of
phylogentic reduction has been blurred by a species's use as a
companion animal with little regard for phylogenetic ranking.
The animals chosen for project usage should be the least advanced
from a phylogenetic standpoint that will provide the necessary
data.
The principle of phylogenetic reduction is generally well
accepted as a way to reduce the number of animals used, but it
often brings many hidden difficulties. As one descends the
phylogenetic scale, the available information on the maintenance
and use of these animals in a biomedical setting often becomes
difficult, if not impossible, to obtain. When choosing a study
model, it is critical that the principal investigator take into
account the ability of the institution to provide appropriate
care for the species chosen. Phylogenetic reduction is an
important means of decreasing the number of animals used, but
should be practiced carefully and with the full knowledge of the
requirements of the species chosen.
Better Quality Animals
It is a rare study in which the initial cost of the animals to
be used represents the single most expensive aspect of the study.
For this reason it can often be false economy to select the
source of the animal based on cost alone. When purchasing
laboratory animals, it is important to keep in mind that cost and
quality are usually directly correlated. By choosing the best
quality animal in terms of health status, the possibility that
animals will be lost or data compromised by the intrusion of a
concurrent disease condition is minimized, if not eliminated.
Choosing the best quality of animal, in terms of genetic status,
will virtually insure the consistency of animals from study to
study. This requires an institutional commitment to the use of
animals of defined health status and limits the investigators to
the animal sources approved by the institution. Mixing of
animals of different health status is a disaster waiting to
happen and may negate all the benefits derived from the use of
quality animals.
The role of the investigator and their staff in assuring the
integrity of an animal colony cannot be overemphasized. In
choosing a source of animals, a veterinarian should be consulted
to insure that the best animals that can be effectively
maintained in the institution are purchased. Animals of
different or unknown health status should never share the same
environment nor common equipment in the animal facility or in the
research laboratory.
REFINEMENT
Refinement refers to techniques which reduce the pain and
distress to which an animal is subjected. For the purpose of this
manual these techniques can be classified into the following
broad categories:
Decreased Invasiveness
Improved Instrumentation
Improved Control of Pain
Improved Control of Techniques
Decreased Invasiveness
A hallmark of most of the new diagnostic and therapeutic
techniques used in human medicine is the minimal degree of
invasiveness that is required to successfully perform a procedure
to obtain a given set of data. In many instances these techniques
are applicable in the research environment and can be adopted for
use in animals. A sophisticated example could be the use of
Magnetic Resonance Imaging for results that formerly required
euthanasia of multiple animals along a time curve to obtain assay
tissue. Today one animal can provide all the information along a
given curve. A less dramatic example is the vascular access
device which permits repeated samples or injections in a single
animal instead of using several animals. Invasiveness reduction
methods are available in almost every area of biomedical
research, and in project design, it is important to identify and
use these methods wherever possible. Not only do they represent
an alternative technique, but they generally provide much more
consistent and reproducible data.
Improved Instrumentation
Monitoring Animals -In this age of microelectronics, fiber optics
and laser instrumentation, the potential for refining techniques
used in animal experimentation seems almost limitless. Improved
instrumentation can minimize animal distress by reducing the
level of restraint and/or manipulation necessary to obtain
biological samples. Included in this category are the use of
tethers in a variety of species to allow continuous access to the
various organ systems, while permitting the animal virtually
unrestricted movement within its primary enclosure. The
advantages of these systems are numerous, not the least of which
is minimizing a variety of nonexperimental variables associated
with prolonged restraint.
Analyzing Samples -Once obtained, samples can be analyzed in very
small volumes for a multitude of parameters. Examples of this can
be found in the commercially available diagnostic laboratory
equipment which require only micro-liter blood samples to perform
a variety of diagnostic tests. The use of smaller sample sizes
permits the use of smaller animal species and prevents the need
to euthanize many of these species to obtain the necessary volume
of blood. It is now possible to obtain serial blood samples from
small laboratory rodents which reduces the number of animals
necessary to obtain data over the length of the study.
Improved Control of Pain
The Animal Welfare Act requires "that the principal
investigator consider alternatives to any procedure likely to
produce pain or distress in an experimental animal" and in any
practice which could cause pain to animals that a doctor of
veterinary medicine is consulted in the planning of such
procedures for the use of tranquilizers, analgesics and
anesthetics. Since appropriate anesthetic and analgesic agents
can minimize the potential pain and distress experienced by
animals, an entire chapter of this manual is devoted to the
principles of using these agents. Suffice it to say, that of all
the possible ways that the 3 R's can be utilized this is an area
where the laboratory animal veterinarian can often be of most
help to the investigator.
Improved Control of Techniques
Proficiency in the handling and restraint of animals makes it
easier to perform a variety of routine procedures with minimal or
no pain or distress to the animals involved. Animals are
creatures of habit and when proper handling is part of their
regular routine, the degree of distress caused by the procedures
is minimized. Animals can be trained or conditioned to accept a
variety of procedures which if suddenly forced upon them can be
distressful.
Almost every animal commonly used in the laboratory responds
positively to a little tender loving care. It's inexpensive,
readily portable, safe even at the highest doses and spreads
rapidly through the staff. To develop the proper techniques and
gain confidence in their use requires training by someone with
appropriate experience. This can be the veterinarian, a member of
the animal care staff or a fellow investigator. Whomever it may
be should be sought out before a new species or technique is
incorporated into the study. This will reduce the potential
distress of all animals involved in the study up to and including
the principal investigator.
SUMMARY
In this chapter, the use of alternative techniques has been
defined in terms of the present regulatory requirements and the
principles of Replacement, Reduction and Refinement were
introduced. In summary, the reader should consider a fourth
R--Responsibility. The use of animals in teaching and research
brings with it a responsibility to minimize animal pain and
distress. The adoption of the 3 Rs as part of the process of
planning and conducting projects using laboratory animals will go
a long way toward implementing Responsibility--the fourth R.
REFERENCES
Animal Welfare Act (Title 7 U.S.C. 2131-2156) as amended by
PL 99-198, December 23, 1985.
Guide for the Care and Use of Laboratory Animals, NIH
Publication No. 86-23.
Models for Biomedical Research: A New Perspective, l985.
National Academy Press, Washington, DC; l985.
Navian, J.B. Animal Models in Dental Research. The
University of Alabama Press.
Paton, William. Man and Mouse Animals in Medical Research.
Oxford University Press, New York, 1984.
Public Health Service Policy on Humane Care and Use of
Laboratory Animals. Revised as of September 1986.
Public Law 99-198. Code of Federal Regulations, Title 9,
Subchapter A, Animal Welfare, 1989.
Rowan, A.N. Of Mice, Models, and Men: A Critical Evaluation
of Animal Research. State University of New York, 1984.
Russel, W.M.S. and Burch, R.L. The Principles of Humane
Experimental Technique, Methuen and Co, Ltd., London, 1959.
U.S. Congress, Office of Technology Assessment. Alternatives
to Animal Use in Research, Testing, and Education. (OTA-BA-273,
Feb. 1986)
Webster's Ninth New Collegiate Dictionary, Merriam-Webster,
Inc., Springfield, MA; 1986.
Wessler, S. l976. Animal Models of Thrombosis and
Hemorrhagic Diseases, NIH Publication No. 76-982.
Chapter 3
Animal Care and Use: A Nonexperimental Variable
John C. Schofield, B.V.Sc., M.R.C.V.S.
and
Marilyn J. Brown, D.V.M., M.S.
INTRODUCTION
The response of a laboratory animal to an experimental
variable is influenced by a variety of genetic and environmental
factors. An understanding of these factors is necessary to
control their affects and minimize the potential influence of
nonexperimental variability on the final outcome of a given
experimental protocol. Minimizing nonexperimental variability
can optimize the use of animals in a given study.
Since the 1930's, the concept of genetic makeup, or genotype
of an animal, combining with the developmental environment to
produce the phenotypic expression of the animal had been well
accepted. A useful concept concerning the relationship of
genetic and environmental factors--dramatype--was proposed by
Russell and Burch in 1959. They defined dramatype to be the
pattern of performance in a single physiological response of
short duration relative to the animal's life time. It is
determined by phenotype and the immediate environment in which
the response is elicited. This concept distinguishes between the
developmental environment, which directly interacts with genetic
factors, and the proximate or immediate environment, which acts
upon the combined system. Simplified, genotype plus
developmental environment equals phenotype and phenotype plus the
immediate environment equals dramatype. This concept stresses
the interrelation of the genetic background of the animal, the
environment in which it is raised and housed and the laboratory
environment in which the animal is used or tested.
Genotype may be controlled through the use of genetically
defined animals produced in structured breeding systems or by
genetic engineering. This is easiest to accomplish through the
purchase of genetically defined animals from reputable suppliers.
In-house breeding programs are difficult and time consuming to
maintain in a manner which assures genotypic integrity. If such
colonies must be used, it is advisable to consult a geneticist to
design a breeding program that produces animals of defined
genetic characteristics. A genetic monitoring program might also
be required to define the genetic makeup of the animals produced.
This can be an expensive proposition and requires some expertise
to perform. The phenotype can be influenced by regulating
environmental conditions in which the animals are reared. For
uniform dramatype, the environmental conditions in which the
animals are tested must be controlled.
This chapter will deal with three broad categories of
nonexperimental variables: physical factors, chemical factors and
microbial factors. Physical factors which will be discussed
include: cage design and construction, temperature, humidity,
ventilation, light intensity and photoperiodicity, noise,
bedding, watering systems, feeding, housing systems, shipping and
handling. Chemical factors to be discussed will include
contaminants of food, water, bedding, and air. Microbial factors
will be discussed in terms of some of the common viral, bacterial
and parasitic diseases that can affect laboratory animals. The
total of all of the components included in these three broad
categories combines with the animal's genetic background to
constitute Russell and Burch's concept of phenotype and
dramatype. It is important to appreciate that our knowlege of
the effects of nonexperimental variables is rapidly expanding and
the purpose of this chapter is to introduce the reader to this
subject rather than present an exhaustive or complete treatise.
PHYSICAL FACTORS
The physical environment of laboratory animals may be
considered to consist of the animal room, or macroenvironment,
and the primary enclosure (cage), or microenvironment. Cage
design and composition influence the interaction between micro
and macro environment. Therefore the temperature, humidity,
airflow, concentration of waste gases, illumination and noise
levels within the cage may be quite different from that monitored
at the room level. Each of these factors represents an important
nonexperimental variable that will be discussed in more detail.
Cage Design
Cage design and construction material can influence the
study results. Galvanized caging material or rubber bottle
stoppers can serve as a source of trace minerals which could
affect the results of studies where the level of these compounds
is being controlled. Other important considerations include
whether contact bedding can be used or if animals must be housed
on a wire floor. The type of sample collection may require the
use of a metabolic cage, or observation studies may require the
use of clear rather than opaque caging. The behavioral
characteristics of the animal will also dictate the type of cage
design used. For example, some animals require perches, nesting
boxes or hiding places, and others require built-in restraint
devices such as the squeeze mechanisms often found in primate
caging. Reproductive needs may require specific caging features.
In some species the male must have a method of escape from an
overaggressive female. Many neonates have inadequate
homeothermic mechanisms and will become hypothermic if not
protected by contact bedding or nesting material placed in the
cages.
Temperature and Humidity
The temperature and humidity in the animal room
(macroenvironment) should be monitored and maintained within
published acceptable limits. The temperature and humidity in the
microenvironment is more difficult to monitor and control.
Variations in temperature and humidity are influenced by such
factors as filter tops, hanging wire or solid bottom caging,
population density, animal activity level, cage location, and
temperature and humidity in the animal room itself. Variations in
temperature and humidity can have a variety of effects. For
example, exposure to high temperatures will frequently cause
rabbits to lick their fur which can predispose them to the
formation of hairballs. Very low humidity has been associated
with a rodent lesion called ring tail which is characterized by
annular constrictions and can result in loss of the tail. More
subtle temperature and humidity effects include: altered drug
metabolism, increased disease susceptibility and decreased
reproductive efficiency. These examples serve to illustrate the
need for controlled temperature and humidity in the animals'
micro and macroenvironment and the vital role it plays in the
generation of consistent, reliable data.
Ventilation
Ventilation in animal rooms can have significant impact on the
health status of the occupants. Excessive odor is often the first
indication of a ventilation problem in an animal room. The
concentration of waste gases at the cage level is usually higher
than those detected at the room level. Furthermore, the
concentrations capable of causing pathology are much less than
human sensory threshold levels. Many design features affect room
ventilation including the location, number, and configuration of
supply and exhaust ducts. Cage-level ventilation is further
affected by the presence and/or type of filter top on the cage as
well as the design and location of the cage relative to the room
airflow pattern. Ventilation should be such that it keeps the
concentration of waste gases to a minimum, reduces the spread of
disease, provides a stable temperature and humidity and avoids
drafts.
Lighting
Light intensity and photoperiodicity in animal rooms can
affect reproductive function and animal vision. The
recommendation of the Guide for light intensity in animal rooms
is 75-125 footcandles (fc). However, prolonged exposure to such
levels can cause irreversible retinal degeneration in albino
rodents and 25 fc has been suggested as a more appropriate
intensity for these species. Variable light intensity control
devices such as dimmer switches or multiple bank lighting can be
installed to facilitate adequate light for observation and
husbandry yet provide lower intensity light for general animal
housing. Cage position on a rack can be an important factor and
an 80-fold difference in light intensity can exist between the
upper and lower shelf locations. Photoperiods or light/dark
cycles (usually given in hours as L:D) can modify reproductive
behavior and circadian rhythms. A daily light cycle which has 12
to 14 hours of light is usually recommended for most species. It
is important to keep the light intensity and periodicity
constant. Animal rooms should be equipped with automatic light
timers. The presence of windows, either to the outside or to the
corridor, can affect reproduction in some animals. Corridor
windows may be desirable for observational purposes, but they can
provide enough light to affect circadian rhythms in nocturnal
animals. As with all environmental factors, the special
characteristics of the animal should be taken into consideration
when planning light cycles. Duration and type of light can
affect estrus behavior. Animals can have their reproductive
cycles manipulated by changing the light cycle. This technique
has been used in several rodent species, cats, and farm animals.
Reversed light cycles can be used to accommodate circadian
rhythm, sleep and breeding studies within the normal working
hours in an institution. Individual room timers provide a
facility with more flexibility to meet a variety of experimental
requirements.
Noise
Excessive noise can also disrupt animal breeding behavior.
Noise at excessive levels can cause mechanical damage to the
auditory system in both animals and man. Some effects of noise
in animals include audiogenic seizures, eosinophilia, increased
serum cholesterol levels and increased adrenal weights. It is
recommended that noise levels in animal facilities not exceed 85
decibels (db).
Caging Accessories
In addition to the microenvironmental effects of the physical
configuration of the primary enclosure as discussed above, other
aspects of the cage environment should be considered. The
presence or absence of bedding material is dependent on the
species and situation. For example, many breeding programs
utilize some form of bedding to improve neonatal survival. An
ideal bedding material should be dustfree, nonpalatable,
absorbent, and free of microbial and toxic contaminants. The
choice of watering system depends on species, experimental
design, and management factors. Automatic watering systems are
expensive to install but can pay for themselves in labor savings
over time. Automatic watering systems should be flushed daily
when used with low flow rates, such as in rodent rooms, to avoid
stagnation and minimize bacterial buildup. When the study
protocol requires delivery of a compound in the water, or
measurement of daily intake is needed, water bottles or
pans are often used. Choice of feeder and type of food is also
species and situation dependent. Some species such as the
hamster are frequently fed on the floor of the cage because their
broad muzzle can make obtaining food from some rodent feeders
difficult. Some species such as the rabbit do not readily
tolerate sudden changes in diet composition or formulation. When
designing a study, it is important to consult someone
knowledgeable in the biology and husbandry requirements of the
species to be used, so that wherever possible, species variations
are taken into consideration.
Cage Size - Occupancy Standards
Consideration should also be given to the cage size. There
are specific cage size requirements set forth in the Guide for
the Care and Use of Laboratory Animals and by the Animal Welfare
Act. Cage size requirements depend upon the species, weight or
size of the animal(s), number of animals in the cage and breeding
status. In addition to the floor space requirements the
behavioral characteristics of the species, strain, and sex must
be considered when group-housing animals. For very social
animals, individual housing may cause stress. Even among social
animals, the formation of new groups can result in fatal trauma
from fighting. Male mice will often fight when group housed,
whereas male rats usually do not. Aggressive behavior can be
strain specific; for example, F344 male rats and C57BL mice are
generally considered to be more aggressive than other commonly
used strains. Even in docile animals, overcrowding can lead to
fighting, cannibalism and stress. Breeding activity can be
significantly modified by group housing arrangements. For
example, group-housing female mice can lead to anestrus with
subsequent estrus synchronization with the introduction of a male
mouse.
Shipping
The shipping of animals can be a significant physiological
stress. Studies have documented the effects that prolonged
transport, high ambient temperatures, lack of water and the
potential for microbial contamination may have on the research
data collected from animals exposed to such factors. The
provision of climate-controlled transport vehicles and filtered
crates decreases these stresses. Even under optimal shipping
conditions, it has been shown that it takes 1-5 days for the
immune system and body weights to return to normal. It is also
important to remember that changes in feed, water, and housing
conditions can markedly affect newly arrived animals. Animals
should be given an adequate period of time to equilibrate after
transport.
Handling
The frequency and type of handling an animal receives is
another nonexperimental variable. Investigators and technicians
should be familiar with and skilled in the correct techniques for
handling and restraining the species involved. This can prevent
injury to either the animal or the handler. Daily husbandry
routines may need to be scheduled around the research needs.
Close communication between the investigator and the animal care
staff can minimize handling stress. For example, collection of
biological samples may be performed during routine cage changing.
This is particularly useful when chemical restraint is required
for either function. Since many animals are creatures of habit,
regular handling may reduce stress.
CHEMICALS
Chemicals found in the animal's environment may be
inherently toxic or their metabolism may result in the formation
of toxic products. They may directly injure cells or interfere
with cellular homeostasis. The possible effect of a chemical
depends on the concentration, the agent's physiochemical
properties, as well as the duration, frequency and route of
exposure and potential interactions. These chemicals can
influence various body systems. For example, it has been
demonstrated that chemicals can affect hepatic microsomal enzymes
which have many functions, including the biotransformation of
drugs and chemicals and regulation of oxygen radical removal.
Such chemical sources include: softwood bedding, room
deodorizers, insecticides, and ammonia. Chemicals can also
target the immune system. Some insecticides cause lymphopenia.
Heavy metals can decrease resistance to disease by the reduction
of antibody formation, altered phagocytic capacity of
polymorphonuclear cells and macrophages, and suppression of
interferon production.
Food and Water
Food and water can serve as sources for chemical contamination
of research animals. Drinking water may be contaminated with
synthetic organic solutes such as pesticides. Trihalomethanes are
often found in water supplies as a result of the chlorination
process. Some facilities hyperchlorinate or acidify water to
decrease microbial contamination; however, these techniques can
affect the immune response. Inorganic contaminants may include
heavy metals and nitrites. Diets can also be a source of
contaminants such as estrogenic compounds, aflatoxins,
insecticides, and preservatives. These compounds may occur
naturally in plant materials, remain as residues from
agricultural use, or be the result of contamination in storage or
the processing procedures. Commercial diets assayed prior to
shipment are available and the results of this assay are printed
on the tag attached to each bag.
Drugs
Drug therapy, prior to or during a study, can compromise the
data obtained. For example, tetracycline alters the immune cell
function through its ability to depress chemotaxis and
phagocytosis. Aminoglycosides can have neuromuscular blocking
properties, and can have negative inotropic effects on cardiac
and arterial muscle. Other agents having neuromuscular
depressant activity include tetracycline, lincomycin, and the
polymyxins. It is important that investigators and the animal
care staff communicate about the effect that any medications may
have on study animals prior to the initiation of treatment.
Similarly, anthelmintics or insecticides given by the animal care
staff to treat parasitism problems, could affect research results
and must be considered in protocol design.
Anesthetic agents are frequently part of experimental
protocols. The researcher should balance appropriate levels of
analgesia, anesthesia, and chemical restraint with the possible
effects of these agents on the experimental results. For example,
the dissociative agent ketamine hydrochloride is widely used in
anesthesia and restraint because it is easy to administer, is
effective in a wide range of species and has a wide margin of
safety. Besides the better known cardiovascular effects of
ketamine hydrochloride, this drug also has been shown to affect
intracellular cyclic AMP, cellular permeability and calcium
channels. A pharmacologic knowledge of these drugs will aid in
selecting those best suited for each experimental protocol and
allow for more informed interpretation of results. Consultation
with the institutional veterinarian regarding the use of
anesthetics and analgesics during the planning of potentially
painful procedures is now a legal requirement.
MICROBIAL FACTORS
Pathogenic microbial agents can affect research by causing
clinical disease, lesions and death. However, in laboratory
animals, infection more frequently is asymptomatic with carriers
who develop overt disease when stressed by shipping or
experimental manipulation. Animals with latent infection may
show no overt disease but research results may be compromised
through subtle physiological, biochemical or histological
changes.
Bacterial Diseases
Species-specific mycoplasmal and bacterial diseases are well
documented. There are a number of these pathogens associated with
commonly used laboratory animal species. For example,
mycoplasmosis is an endemic disease in some conventional rodent
colonies. It can cause respiratory and genital tract infections
thereby affecting exercise tolerance, sensitivity to anesthetic
agents, increased susceptibility to other respiratory pathogens,
decreased reproductive efficiency and a variety of immune system
anomalies. The investigator using rabbits should be aware of the
incidence and significance of pasteurellosis as a cause of acute
and chronic disease. Pasteurella multocida is very common in
conventional rabbit colonies and can cause upper and lower
respiratory tract infections, subcutaneous abscesses, middle and
inner ear infection and reproductive tract infections. Some
species may serve as asymptomatic carriers of bacterial
infections which can cause severe clinical disease in other
species; therefore different species should not be mixed.
Bordetella bronchiseptica can often be isolated from clinically
normal rabbits and rarely causes disease in that species but it
can be a significant cause of respiratory disease in guinea pigs.
In addition to the species-specific organisms, post-operative
infections can be caused by a myriad of bacterial contaminants
normally present in the animal's environment. It is important
that invasive surgical procedures be done aseptically to minimize
the potential affects of these opportunistic organisms.
Although not experimental variables, there are several
bacterial diseases of laboratory animals which can be transmitted
to man and therefore are of possible concern to those using
animals in research. These may include tuberculosis,
salmonellosis, campylobacterosis, and shigellosis. The
investigators whose studies involve substantial animal contact
should be familiar with institutional guidelines and policies
regarding the prevention of zoonotic disease. These should
include a program of periodic physical examination, an
educational program for personnel, immunization where appropriate
and the use of protective clothing.
Viral Diseases
Viral infections in laboratory animals can often be
asymptomatic. As with bacterial and mycoplasmal infection,
clinical viral disease can occur when an animal is stressed.
These viruses can be particularly devastating because the effects
on research data may not be recognized, yet still be significant.
The effects of these latent viruses have been best defined in
rats and mice. Barrier housing of commercially available
specific pathogen-free rodents will help eliminate these viruses
from a colony. Contaminated tissues, particularly murine tumors,
have been implicated in many outbreaks of disease. Tissues
should be screened for the presence of contaminants prior to
their use in a research facility. It is beyond the scope of this
chapter to review all the research implications of viral
pathogens currently known; however, a few examples will be
briefly mentioned. There are key viral diseases of most common
laboratory animals and it is important for the investigator to
work with the institutional veterinarian to become familiar with
those viruses and learn how they might affect a particular
research project.
Sendai virus, a common viral contaminant in conventional mouse
and rat colonies, can cause histopathologic changes in the
respiratory tract, immunosuppression, and decreased reproductive
efficiency. It can also act synergistically with other
respiratory pathogens. A viral disease of mice which is often
asymptomatic but serious is Mouse Hepatitis Virus (MHV). This
virus has been implicated in wasting syndromes in nude mice. It
can cause respiratory, hepatic, and enteric disease. Even in
asymptomatic animals, it can cause profound immunological
disturbances. Some diseases of laboratory animals are often
associated with clinical disease and affect a research study due
to high morbidity and mortality rather than the subtle effects of
the latent viruses. Canine distemper, feline panleukopenia and
measles in macaques are examples of these types of viral
infections. Although not as prevalent as bacterial zoonoses,
some viruses of laboratory animals can be transmitted to man.
Examples of these include; lymphocytic choriomeningitis, Herpes
virus simiae and rabies.
Parasitic Diseases
Parasites of laboratory animals have also been implicated as
nonexperimental variables in research. Some parasites such as
Trichosomoides crassicauda of rats are capable of causing tumors
which could significantly obscure results of a carcinogenicity
study. Skin mites of mice have been shown to affect immune
parameters. Parasites are also capable of causing significant
clinical disease such as the rectal prolapses seen with pinworms
in rodents and bowel perforation seen with Prosthenorchis elegans
in non-human primates. Some parasites of laboratory animals can
also be transmitted to man. Examples of these parasites are
Hymenolepis nana and Entamoeba histolytica.
It is important to remember that while laboratory animals may
not show clinical signs of microbial infection, the infections
can have profound effects on research results. Investigators
studying immunological function should be particularly familiar
with the otential effects of microbial agents on their research.
Transmission of contaminants can occur in tumor or tissue
inoculation, from direct transmission or via fomites in the
laboratory. Animals of different health status should be strictly
isolated from one another and all biologic material should be
screened for the presence of viral and other contaminants.
SUMMARY
The concepts of Russell and Burch-refinement, replacement, and
reduction-are generally well accepted in the research community.
Adherence to these concepts includes attempting to minimize the
nonexperimental variables introduced in this chapter. The
maintenance of healthy laboratory animals and the reduction of
nonexperimental variables is the responsibility of the animal
care facility and the investigator working together in an
atmosphere of open communication and cooperation.
REFERENCES
Allert, J.A.; Adams, R.A.; and Baetjer, A.M. l968. Role of
environmental temperature and humidity in susceptibility to
disease. Ach. Environ. Health l6:565-570.
Broderson, J.R., et al. l976. The role of ammonia in
respiratory mycoplasmosis of rats. American Journal of Pathology
85:ll5-l30.
Davis, D.E. l978. Social behavior in a laboratory
environment. pp 44-63 in Laboratory Animal Housing. Proceedings
of a symposium organized by the ILAR Committee on Laboratory
Animal Housing. Washington, DC; National Academy of Sciences.
Guide for the Care and Use of Laboratory Animals, NIH
Publication No. 86-23.
Greenman, D.L.P., et al. l982. Influence of cage shelf level
on retinal atrophy in mice. Lab Animal Science, 32(4):353-356.
Lang, C.M. and Jessell, E.S. l976. Environmental and Genetic
Factors affecting laboratory animals; impact on biomedical
research. Federal Proceedings Vol. 35 No. 5-8, ll23-ll65.
Lindsey, J.R., et al. Physical, chemical and microbial
factors affecting biologic response, pp. 3-43, In: Laboratory
Housing. Proceedings of a symposium organized by the ILAR
Committee on Laboratory Animal Housing. Washington, DC; National
Academy of Sciences.
Pakes, S.P. et al., Factors that complicate animal research,
Laboratory Animal Medicine, Chap. 24. Fox, J.G. (ed.), Academic
Press.
Public Law 99-198. Code of Federal Regulations, Title 9,
Subchapter A, Animal Welfare, 1986.
Russell, W.M.S. and Burch, R.L. The Principles of Humane
Experimental Technique, Methuen and Co., Ltd., London, 1959.
Chapter 4
Principles of Anesthesia and Analgesia
Marilyn J. Brown, D.V.M., M.S.
INTRODUCTION
It is important that all scientists using animals in research
meet their ethical and legal responsibilities to avoid
unnecessary pain and distress to the animal. Studies involving
unavoidable pain and distress must be justified by the
investigator in accordance with Federal regulations and
institutional policies. This chapter will cover some of these
legal responsibilities as well as try to help the investigator
meet these responsibilities through knowledge of the basic
principles of anesthesiology. Included in these principles are
an understanding of some of the basic terms used in the field of
anesthesiology, the types of variables that can affect an
animal's response to an anesthetic agent, the effect of a given
anesthetic protocol on an experiment, some general considerations
and the recognition of pain. Also mentioned in this chapter are
anesthetic monitoring and some fundamentals of anesthetic crisis
management. Controlled drugs and their use are briefly
discussed. This chapter is not meant to be a complete treatise
on the subject of laboratory animal anesthesiology but to give an
introduction to stimulate further reading in areas of specific
interest.
Anesthesiology is not an exact science. Recommendations and
dosages given in textbooks should be taken as guidelines. An
investigator contemplating a procedure requiring anesthesia,
tranquilization or analgesia should not neglect the resource of a
veterinarian who can often provide valuable assistance. In fact
the Animal Welfare Act requires that "in any practice which could
cause pain to animals . . . a doctor of veterinary medicine is
consulted in the planning of such procedures."
There are many variables affecting an animal's response to
anesthesia. Because the absorption and biotransformation of drugs
differs between species, it is nearly impossible to develop a
single anesthetic or analgesic protocol that applies to all
laboratory animals. Morphine can cause profound CNS depression in
the rat and rabbit but can cause tremors and convulsions in mice
and cats. The dosage of xylazine needed to sedate a ruminant is
one-tenth that necessary to sedate a dog. These are but two of
many examples. A common mistake is to extrapolate dosages across
animal species or from man to animals. The strain of animal used
is also a variable to consider. Some rat strains are sensitive to
nitrous oxide. Some breeds of dogs (whippets and greyhounds) are
more sensitive to barbiturates than other breeds. The size and
even the sex of the animal can make a difference in the response
to anesthetics. In rats, females are more sensitive to
barbiturates but in mice, barbiturate narcosis lasts longer in
males. The temperament of the animal can change the way it
responds to a given agent. Some tranquilizers will cause a
vicious dog to become even more difficult to handle.
Fat does not play a key role in the initial absorption of an
anesthetic agent, but it does affect the body weight upon which
the dosage is based. Fat can later serve as a repository for the
agent, thus prolonging recovery. The age of the animal also must
be considered. Since very young animals require frequent
feedings, prolonged recoveries can present a formidable problem.
There are also age-related changes in liver enzyme functions
which affect biotransformation of anesthetic agents. Older
nimals can present an anesthetic challenge due to impaired renal
or hepatic function.
The animal's physical condition can affect its responses. The
presence of pre-existing disease will increase an animal's
anesthetic risk. Respiratory diseases can often be asymptomatic
in the uncompromised animal even though it is endemic in many
rodent populations. Even less obvious is the effect of diet and
environment. Rats fed an inadequate diet are more resistant to
barbiturates, yet fasted mice have an increased barbiturate sleep
time. Abnormal environmental temperatures and humidity cause
stress which can result in a compromised animal and variable
anesthetic responses. High temperatures sensitize rats and
rabbits to anesthesia.
Various factors will influence anesthetic choice. The use of
concurrent drugs changes an animal's response to anesthetic
agents. For example, some antibiotics potentiate barbiturates.
The type of experimental procedure planned may impact on the
anesthetic protocol. In an obstetric procedure, the effects on
the fetus must be considered. When surgery involves the head and
face, there is limited access to the animal so the anesthetic
protocol should be planned to facilitate monitoring under these
circumstances.
LEGAL RESPONSIBILITIES
Minimizing pain and distress in research animals is an ethical
responsibility, produces better scientific results and is the
law. The Public Health Service Policy on Humane Care and Use of
Laboratory Animal states that "Procedures that may cause more
than momentary or slight pain or distress to the animals will be
performed with appropriate sedation, analgesia, or anesthesia
unless the procedure is justified for scientific reasons in
writing by the investigator." The NIH further addresses the
subject of anesthesia in the Guide for the Care and Use of
Laboratory Animals. This document states that the proper use of
anesthetics and analgesics is necessary for humane and scientific
reasons and recommends that the veterinarian provide guidance for
their usage. The Animal Welfare Act (AWA) requires standards for
animal care, treatment, and practices in experimental procedures
to ensure that animal pain and distress are minimized, including
adequate veterinary care with the appropriate use of anesthetic,
analgesic, tranquilizing drugs or euthanasia. It prohibits the
use of paralytics in painful procedures without anesthesia and
states "that the withholding of tranquilizers, anesthesia,
analgesia or euthanasia when scientifically necessary shall
continue for only the necessary period of time." Exceptions to
such standards may be made only when specified by the research
protocol and any such exception shall be detailed and explained
in full in a report filed with the Institutional Animal
Committee. And as previously noted, it further requires that if
practices could cause pain to animals, a doctor of veterinary
medicine be consulted in the planning of such procedures.
TERMINOLOGY
As with all branches of science, there are certain terms one
needs to be familiar with in order to communicate effectively
about anesthesiology.
The following is a list of the most common terms:
Analgesia Insensibility to pain without loss of
consciousness.
General Temporary, controllable and reliable loss of
Anesthesia consciousness induced by intoxication of the
CNS.
Sedation Calm state usually accompanied by
drowsiness.
Tranquilization Calmness without drowsiness or
unconsciousness. Analgesia is usually not
a feature.
Time to Peak Time between initial administration and onset
of the effect maximum expected effect.
Duration of Effect Length of time peak effect can be expected
to last after a single administration of an
anesthetic dose.
Time to Recovery Time between initial administration and the
ability to stand unaided.
EFFECTS OF ANESTHESIA ON RESEARCH
When anesthesia, analgesia, or chemical restraint is used, it
may be advisable to ascertain any distortion of results by
anesthetics through limited trials. Check the literature and
package inserts for the effect of the agent on the systems being
experimentally evaluated. These changes need to be taken into
consideration when evaluating the effect of an experimental
manipulation. Choose the agent which has the least effects on the
systems under investigation. General anesthetics often depress
the cardiovascular and respiratory systems, alter blood gases,
lower metabolism, decrease body temperature, and alter tissue
perfusion. Anesthetics can also produce histopathologic changes.
GENERAL CONSIDERATIONS
Whenever possible, try a new anesthetic protocol in a limited
number of animals before depending on it for surgical or painful
procedures involved in an experiment. This allows determination
of suitability for the anticipated protocol and allows necessary
changes to be made before it effects the data being collected.
It also facilitates familiarization with the anesthetic method to
minimize problems later, when attention is often focused on
surgical procedures or data collection.
Pay particular attention to the health of the animal before
using it in an experiment. A preanesthetic checkup is a good
idea. To minimize anesthetic risks, only use healthy animals and
allow them to acclimate to the facility before an anesthetic
procedure. Consider the general adaptation syndrome: alarm
increases basal metabolic rate which may increase the amount of
anesthetic needed; however, this is often followed by an
exhaustion phase when less anesthetic is required.
Use the minimal degree of CNS depression necessary for the
procedure that is compatible with the animal's welfare. The
degree of depression required for procedures such as radiographs
or blood withdrawal is not the same as that needed for a
thoracotomy or orthopedic procedure. Remember, during painful
procedures, the use of paralytics without anesthesia is
prohibited by law.
Consider if, and to what extent, the anesthetic protocol will
affect the validity of experimental results and how it will react
with other drugs being used. For example, if studying
catecholamine effects, halothane should be avoided since its
combination with catecholamines can cause severe cardiac
dysrhythmias.
Even in the absence of sophisticated equipment, try to have
some basic items available to insure adequate ventilation. This
includes a source of oxygen, the use of endotracheal tubes when
feasible, and aspiration suction to remove excessive oral
secretions, and/or vomitus.
Regard the conservation of heat as an integral part of
anesthetic management. This is particularly important in small
or young animals. A rectal thermometer can help monitor the
animal's body temperature. More sophisticated thermal monitors
are also available. Maintenance of body temperature is enhanced
through the use of external heat sources such as hot water
bottles, thermal blankets and heating pads. Care should be taken
to avoid thermal burns from external heating sources; i.e.,
electric heating pads.
Administer warm, balanced salt solutions by continuous I.V.
drip whenever possible. This is not always possible in very small
animals but is especially important for prolonged procedures or
when significant blood loss is expected. Fluids often come in
bags which are easy to handle and when warmed can double for hot
water bottles.
Pay particular attention to post-anesthetic care. The
anesthetist's responsibility does not end when the animal is
taken off the table. Allow animals to recover in an environment
approaching the normal body temperature of the species. Maintain
intravenous fluid infusions when possible and have an
endotracheal tube in place until the swallowing reflex is
recovered. Be sure the animal is protected from injury, either
self-inflicted or by other animals, during recovery.
Consider the implications for laboratory safety. Scavenging
systems should be used with gaseous agents. Avoid carcinogens
such as urethane and chloroform. Consider flammability when
using ether.
RECOGNITION AND TREATMENT OF PAIN
In the Definition of Terms developed to implement the amended
Animal Welfare Act, a painful procedure is defined as, ". . any
procedure that would reasonably be expected to cause more than
slight and momentary pain or distress in a human being." In both
humans and most animals the total pain experience results from an
interaction between sensory pathways and the affective system,
which provides the motivational and emotional component of pain.
This varies considerably between species and individuals within
a species.
Understanding the degree of pain involved in various
experimental procedures allows a prediction of animal pain or
distress. Physiological responses to pain can include increased
blood pressure and heart rate, pupillary dilation, increased
respiration, and an arousal response on the electroencephalogram.
If baseline values are known for these variables, they can be
monitored for changes.
To detect behavioral signs of pain, one must be familiar with
the animal's normal behavior. Behavioral responses to pain vary
between species, within species, and even within the same animal.
General behaviors to evaluate include: sleeping, feeding,
drinking, locomotion, grooming, exploration, performance in
learning and discrimination tasks, mating behavior, social
interactions, and dominance/subservience responses within the
social system.
Typical behavioral signs of acute pain include:
- protecting the painful area
- vocalizing (especially when handled or moving)
- licking, biting, scratching, or shaking the painful
area
- restlessness
- lack of mobility
- failure to groom
- abnormal postures
- lack of normal interest in surroundings.
Unless there is evidence to the contrary, assume that a
procedure that causes pain in humans will cause pain in animals.
Points to remember are:
- Abdominal surgery appears to be less painful in animals
than humans, probably because most animals do not use their
abdominal muscles for postural support.
- Lumbar and thoracic spine surgery in animals also appears
to be less painful than in man, probably due to man's postural
requirements. However procedures involving the cervical spine
seem to be more uncomfortable in animals.
- In animals, chest surgery involving the sternum appears to
be more painful than surgery using a lateral intercostal
approach.
- Surgery on the eye, ear or surrounding structures seems to
distress most animals. Signs such as head tilt or shaking, or
pawing or rubbing the area may be seen. Perirectal procedures
also seem to produce discomfort. In addition to analgesia,
protection of the affected areas is indicated.
- Surgery of the femur or humerus also seems to be painful
to most animals, which may be due to large muscle mass trauma.
Pain perception can be influenced by drugs and/or
environmental and behavioral factors. Recovery in familiar
surroundings may help to relieve pain and distress.
Acclimatization prior to a procedure may also facilitate
recovery. The environment should be kept stable, minimizing
stimuli that evoke a fearful response in the animal. When
appropriate, interact with the animal through talking or petting.
Always handle the animal in an appropriate manner.
Various analgesics are available to the investigator. These
can be divided into two main categories: the centrally acting
agents such as morphine, butorphanol and buprenorphine; and the
peripherally acting agents such as the anti-inflammatories,
aspirin and phenylbutazone. The short half-lives of many of
these agents may cause a labor-intensive analgesic protocol for
the investigator, but creative delivery systems (such as the
osmotic minipumps and tethering systems) and the development of
new drugs such as buprenorphine with longer half-lives (12 hours)
should facilitate meeting the analgesic needs of most laboratory
animals. When designing an analgesic protocol, the investigator
should consult with a veterinarian who is experienced in
laboratory animal medicine. This will help avoid problems with
species specific responses such as morphine sensitivity in cats
and mice or the unusually short duration of meperidine in the
dog. Interaction of the analgesic with concurrently used drugs
and the effect of the agent on study results (such as the effect
of aspirin on healing or clotting time) must be taken into
consideration when choosing the best agent for a given situation.
Although there is much information available on the use of
various agents in animals, it is not always easily referenced and
may be difficult to find without some guidance.
ANESTHETIC MONITORING
During an anesthetic procedure, the physiologic state of the
animal and the depth of anesthesia should be monitored. This
allows the anesthetist to adjust the depth of anesthesia and to
anticipate impending complications. The degree of jaw tone is an
indication of muscle relaxation. This is easily monitored by
trying to open the animal's mouth -- taking care to avoid the
animal's teeth.
Pulse quality is an indication of cardiovascular function. It
can be checked in several areas but is commonly felt in the
inguinal region. This "hands on" evaluation of the animal also
gives the anesthetist a crude indication of the animal's body
temperature so that hypo- or hyper-thermic states can be
detected. Capillary refill is also an indication of
cardiovascular function. This is checked by pressing firmly on
the mucous membranes of the gums until they blanche and then
releasing the pressure and noting the time it takes the normal
color to return. Full color should return in less than two
seconds. A slow capillary refill time is suggestive of sluggish
blood flow and may be an early indicator of shock. While
checking capillary refill, also note mucous membrane color.
White may indicate shock, while blue may indicate poor
oxygenation. In small rodents, the foot pads or ears offer other
areas to check for color.
Another method for monitoring cardiovascular and respiratory
function is through auscultation of the chest. This takes more
experience and is difficult in small rodents.
Electrocardio-graphic monitors are also available to aid in
anesthetic monitoring.
Keeping written records of your anesthetic monitoring and
administration is important for several reasons. They serve as a
permanent record of the procedure and of any complications and
when they occurred. This can help explain unexpected
experimental data later. Written records also help to visualize
significant trends which could lead to anesthetic complications.
In addition, written records represent the best method to clearly
document compliance with the AWA.
The aim of anesthesia is to prevent the perception of
painful stimuli without undue depression of physiologic
functions. One of the criteria used to monitor the depth of
anesthesia is the animals' response to stimuli or their reflex
responses. Responses vary with the type of anesthetic used, the
species and health status of the animal, and the use of
concurrent drugs, particularly paralytics.
The first reflex lost is usually the righting reflex. This
reflex may be checked by turning the animal over on its back and
watching to see if the animal rolls back over onto its sternum.
Obviously an animal that can right itself is not at a surgical
level of anesthesia!
The next reflex usually lost is the swallowing or laryngeal
reflex. It is the loss of this reflex that allows placement of an
endotracheal tube after induction. Once in place, slight
manipulation of the tube will cause the animal to swallow, if it
is waking up. With some commonly used anesthetics such as the
dissociative, ketamine, the laryngeal reflex may be present even
when a surgical level of anesthesia is obtained.
The palpebral or eyelid reflex is an easy one to monitor. A
light touch to the medial canthus or brush of the eyelashes will
cause eyelid movement if the reflex is present. It may be as
obvious as a blink or just a slight muscle movement. An overly
aggressive touch may cause movement that is not induced by the
animal and can lead to erroneous interpretation.
The reflex most commonly used to determine if the animal is
feeling deep pain is the pedal or paw pinch reflex. The toe is
firmly pinched between the fingers to elicit a withdrawal
response by the animal. A forcep may also be used but care must
be taken not to cause tissue damage. Pinching the ear can also
be used especially in rodents and rabbits. If the animal draws
its head away or shakes its ear, it is still capable of feeling
deep pain and is not ready for any surgical manipulations.
The pupillary reflex can also be monitored but it can be
affected by many things. Common preanesthetic agents often make
the pupil unresponsive to light. A dilated pupil can indicate
either very light anesthesia and the perception of pain or
dangerously deep anesthesia if the pupil is fixed and dilated.
The corneal reflex is usually the last to go and it is usually
not necessary to get to this depth of anesthesia. This reflex is
checked by very gently touching the animal's cornea and watching
for movement of the eyelid.
STAGES AND PLANES OF GENERAL ANESTHESIA
General anesthesia is divided into stages and planes. Stage
one is characterized by analgesia. In stage two, excitement can
be seen. Signs include struggling and erratic movement. It is
preferable to avoid this stage. Stage three is a surgical level
of anesthesia. It is further divided into planes. Plane one is
characterized by a loss of the palpebral reflex. In plane two,
eyeball movement ceases and the animal exhibits deep, regular
respirations. This is usually a good level at which to do
surgery. With plane three comes paralysis of the intercostal
muscles and short, jerky, gasping diaphragmatic efforts.
Artificial ventilation is essential at this plane. Stage four is
one to avoid as it is characterized by total loss of respiratory
movements, cyanosis and cardiac arrest.
SPECIFIC AGENTS
It is not within the scope of this chapter to give a detailed
pharmacologic description of all the anesthetic agents and
regimes used in research animals. However, a brief description
of the advantages and disadvantages of some of the most commonly
used agents will be given. The reader is referred to the list of
references and a veterinarian when help is needed to design an
appropriate anesthetic protocol for a given research project.
Preanesthetics
Preanesthetics are usually given as an anesthetic agent
adjunct to ameliorate some of the deleterious side effects and/or
to decrease the required dose of the primary anesthetic agent.
Atropine or its analogs are commonly given. They depress
secretory activity making them especially useful in animals with
profuse oral secretions such as ruminants and guinea pigs. These
agents also help maintain heart rate by counteracting the vagal
slowing of the heart rate induced by some anesthetic agents and
some surgical procedures. Atropine causes pupillary dilation,
therefore this reflex cannot be used to monitor anesthetic depth
in the atropinized animal.
Other commonly used preanesthetics are tranquilizers and
sedatives. Use of these agents helps provide a stress-free
subject for the induction of anesthesia. Acepromazine produces
good tranquilization, indirectly suppresses the emetic center,
potentiates the analgesic effects of other agents and provides
muscle relaxation. Hypotension can be a serious side effect of
this agent. It is often used in combination with the dissociative
anesthetic agents such as ketamine. Xylazine is a potent
hypnotic, muscle relaxant, and analgesic. Use of this agent can
reduce the necessary barbiturate dose by 50 percent. Like
acepromazine, xylazine is often used in combination with
ketamine. Bradycardia and hypotension can be seen with xylazine.
Premedication with atropine can help prevent cardiac
dysrhythmias. Respiratory rate can be decreased but increased
tidal volume usually maintains normal blood gases. Xylazine can
cause abortion in late pregnancy in ruminants. Diazepam is a
potent tranquilizer which also has muscle relaxant and
anticonvulsant properties. It is useful in combination,
particularly with Innovar-VetR in rodents. Although diazepam can
cause some respiratory depression, it has little effect on
cardiac output or blood pressure. Morphine is a narcotic
analgesic sedative. Anesthetic doses can be decreased as much as
50 percent after morphine administration. Morphine depresses the
central nervous system, particularly the respiratory center, as
well as peristalsis. In dogs, morphine frequently causes emesis.
Morphine is generally contraindicated in the cat and mouse.
General Anesthetic: Injectable
General anesthesia is delivered by two basic methods:
injection and inhalation. It is usually preferable to give
injectable agents by the intravenous route (I.V.) -- given to
effect; however, intraperitoneal (I.P.), subcutaneous (S.C.) or
intramuscular (I.M.) techniques are sometimes necessary or even
preferable. The advantages of injectable anesthetic agents are
ease of administration, low cost and lack of need for
sophisticated equipment. The major disadvantage is that once the
drug is given, it is in the body until it is metabolized or
excreted.
Innovar-Vet(R) is a veterinary drug which combines fentanyl,
a morphine derivative, and droperidol, an alpha adrenergic
blocker. Because it is a combination drug, doses are usually
given in ml/kg rather than mg/kg. It is a potent analgesic. The
cardiac depressant effects can be counteracted with atropine and
the respiratory depressant effects can be reversed with naloxone.
Innovar-Vet(R) is a poor muscle relaxant. It is not recommended
for use in horses, ruminants, or cats.
Ketamine is a commonly used dissociative anesthetic. It is
short acting and produces variable analgesia. It is often
combined with other agents to improve its muscle relaxation and
analgesic properties as well as provide a smoother recovery. It
can be given I.V., S.C., or I.M. It does not cause cardiac
depression and may even stimulate the cardiovascular system;
however mild respiratory depression may be seen. The swallowing
reflex is maintained making intubation under ketamine alone
difficult. The palpebral reflex is lost, so it is necessary to
use ophthalmic ointment to prevent corneal drying.
The most commonly used injectable anesthetic agents are the
barbiturates. There are two classes of barbiturates:
oxybarbiturates of which pentobarbital or nembutal is the most
common; and thiobarbiturates which are much faster acting and
which include thiopental and thiamylal. Barbiturates are
potentiated by acidosis such as that which can be seen with
respiratory depression or diarrhea. Many drugs potentiate the
effect of barbiturates. Glucose or epinephrine cause prolonged
recovery times. Barbiturates are controlled substances as
defined by the Drug Enforcement Agency. Therefore a license is
required for purchase and records must be kept. If possible,
barbiturates should be given to effect which is difficult when
administered I.P. They have an accumulative effect, which means
two subsequent doses combined have a greater effect than the two
doses given alone. Barbiturates are considered poor analgesics.
Respiratory depression can lead to hypercarbia. Cardiovascular
effects include bradycardia, hypotension, myocardial depression,
and increased peripheral vascular resistance. Use of
barbiturates is contraindicated in animals with liver or kidney
disease. Lower doses should be used in young animals. When
small doses must be given, it is often helpful to dilute stock
barbiturate solution. Preanesthetics should be used when
possible to decrease the amount of barbiturate needed.
General Anesthetics: Inhalation
Inhalation anesthesia has the advantages of rapid induction
and recovery. Depth of anesthesia can be rapidly changed.
Typically animals are initially anesthetized with an I.V.
injection of an ultrashort acting barbiturate, or administered
the inhalation agent by mask or by use of an induction chamber.
When using gaseous anesthetic agents particular attention must be
paid to provide an adequate oxygen source and for the removal of
carbon dioxide. This can be done through the use of a properly
maintained gas anesthesia machine. If possible, it is preferable
to intubate the animal for the most efficient delivery system and
to help assure a patent airway. This takes practice, especially
in rodents. If the anesthesia is administered by mask, avoid
placement of the mask over the entire face as these agents are
irritating to the eye. Also avoid direct contact of the liquid
form of the agent with the animal's skin or mucous membranes.
Scavenging systems should be in place to minimize personnel
exposure.
Nitrous oxide is often used in conjunction with an anesthetic
gas due to its potentiating effect. It is always used in
combination with oxygen, usually at a 50:50 or 60:40 ratio. It is
quite safe, since it is neither flammable nor explosive, allows
rapid induction and causes little cardiovascular disturbance. It
is also a very good analgesic. It enters air-filled cavities much
faster than it leaves them which could be a problem with a
pneumothorax or a large gas-filled bowel. Oxygen should be
administered alone for a few minutes at the end of a procedure to
prevent diffusion anoxia.
A commonly used gaseous anesthetic agent is ether. Ether has
a slow induction and recover period. It is highly flammable and
forms explosive mixtures with oxygen and nitrous oxide. It is a
potent CNS depressant and analgesic. It is extremely irritating
to the mucosal lining of the respiratory tract and may induce
laryngospasms, especially in cats and rabbits. Respiratory
secretions are stimulated which can predispose or exacerbate
respiratory infections. The respiratory depression caused is
usually only a problem in guinea pigs and chinchillas. Ether
causes some myocardial depression. Since ether is inexpensive
and can be administered without the use of sophisticated
equipment, it is very popular. To minimize explosive hazards and
personnel exposure, ether should be used under a fume hood.
Three other commonly used inhalation agents are halothane,
isoflurane and methoxyflurane. Halothane is nonflammable and
nonexplosive. It is a good muscle relaxant and adequate
analgesic. It allows rapid, smooth induction and recovery.
Halothane depresses the cardiovascular system and sensitizes the
heart to dysrhythmias. It also depresses the respiratory system
which can lead to acidosis. Halothane requires special
vaporizers and equipment. Isoflurane is also a stable,
nonflammable agent. Induction and recovery are rapid. Arterial
blood pressure is decreased due to lowered peripheral vascular
resistance; however, perfusion is maintained. Other
cardiovascular functions are well maintained, but respiratory
function is depressed. Isoflurance also requires special
vaporizers and equipment. Methoxyflurane is very stable and
because it does not reach high concentrations at room
temperature, it has a good margin of patient safety. It is a
good muscle relaxant and an excellent analgesic. Like the other
inhalation agents, it does cause some respiratory depression and
hypotension can also be a problem. Induction and recovery are
slower than with the other agents which may be an advantage by
keeping the animal quieter immediately post operative as well as
providing longer acting analgesia.
SPECIES-SPECIFIC CONSIDERATIONS
When anesthetizing small rodents, particular care must be
taken to avoid hypothermia. The airway is easily obstructed so be
sure the neck is adequately extended and secretions are aspirated
as necessary. Fasting is not necessary unless gastrointestinal
surgery is planned and even then only a 6-hour fast is necessary.
Water should not be restricted. Loss of the toe pinch reflex
indicates surgical anesthesia in the mouse. In the rat and guinea
pig, the ear pinch is more sensitive. Rodents are difficult to
intubate. If they are intubated, care must be taken to minimize
dead space in the tubing.
Rabbits are probably the most difficult laboratory animal to
anesthetize. Their respiratory center is particularly sensitive
to anesthetics and a lot of individual variation in response
exists. Rabbits should be fasted 6 hours prior to anesthesia.
Water should not be restricted. The rabbit trachea is very
delicate and rabbits are predisposed to pulmonary edema with
prolonged inhalation administration. A normally small lung
capacity combined with enzootic pulmonary disease further
complicates the situation. The best indicator for surgical
anesthesia is the loss of the ear pinch reflex. Intubation in
rabbits is difficult due to lack of visualization of the larynx,
but it can be mastered with practice.
Dogs are usually not difficult to anesthetize. Large, easily
accessible veins make I.V. injection of agents quite easy.
Intubation is not difficult due to the easily visible larynx.
Administration of preanesthetics, particularly in large dogs, may
make induction easier. Dogs should be fasted for 12 hours prior
to nesthetic administration.
Cats are also relatively easy to anesthetize; however, they
are easily stressed when restrained so preanesthetics become even
more important. The larynx is easy to visualize, but
laryngospasms can make intubation difficult. Cats should be
fasted for 12 hours prior to an anesthetic procedure; however if
necessary, xylazine can be given to induce vomiting and serve as
a tranquilizer. As noted previously, narcotics can cause severe
convulsions in cats and should be avoided.
There are several considerations when anesthetizing swine.
The pig heart is smaller in proportion to body size than is the
heart of other domestic animals, which is a disadvantage during
periods of anesthetic stress. Size and temperament can make
restraint difficult and the use of preanesthetics essential.
Pigs are predisposed to ventricular fibrillation and some breeds
exhibit malignant hyperthermia when exposed to halothane. The
anatomy of the larynx and soft palate predispose pigs to
respiratory distress if not intubated; however, this same anatomy
combined with laryngospasms can make intubation difficult. The
ear vein is the most readily accessible for I.V. injections.
Pigs should be fasted 12-18 hours prior to anesthetic
administration.
The temperament and size of ruminants present a challenge to
the anesthetist. Again, preanesthetics are desirable; however,
ruminants are very sensitive to xylazine, so only small doses are
needed. They are also very sensitive to barbiturates. Food should
be withheld for 24-48 hours with water withheld for 6 hours prior
to the procedure. Gastric bloat can be minimized by passing a
stomach tube in the anesthetized animal once it is on the table.
Avoiding prolonged procedures and recoveries will also help
minimize bloat as well as decrease the incidence of pressure
myositis. Thick pleura and extensive pulmonary supportive tissue
necessitate the use of high ventilation pressures. Excessive
salivation is difficult to control even with the generous use of
atropine. The jugular vein is the easiest to use for the I.V.
anesthetic administration.
The use of sedatives and tranquilizers as preanesthetics in
nonhuman primates presents a risk to the handler because the
animal may present a false appearance of sedation in the cage and
become quite active when aroused! Ketamine, given I.M. in a
monkey restrained in a squeeze cage, is the most common form of
preanesthesia. This is often followed with an I.V. injection of
an additional anesthetic agent with maintenance accomplished with
additional injectable agents or inhalation anesthetics. This
procedure minimizes the hazards of bites or scratches to
personnel or escape by the patient. Monkeys should be fasted for
12 hours prior to an anesthetic procedure; however, ketamine
given alone usually does not cause emesis. Monkeys are usually
not difficult to intubate after a little practice.
ANESTHETIC EMERGENCIES
Anesthetic emergencies are usually caused by human error.
This may be due to inappropriate selection of agents or doses,
failure to recognize and treat inadequacies of respiration or
circulation before collapse, neglect in checking equipment or the
use of unhealthy animals.
Respiratory failure is often caused by airway obstruction or
barbiturate overdose. Airway obstruction can occur because of
positioning of the animal, secretions in the trachea, or
misplacement of the endotracheal tube. Barbiturates are
particularly potent respiratory depressants and they must be used
with care and "to effect." Signs of respiratory failure include
gasping, exaggerated chest movements and cyanosis. Gasping
movements can be misinterpreted to be voluntary movements
indicating inadequate anesthesia causing the inexperienced
anesthetist to actually give more anesthetic agent.
When respiratory failure occurs, the first thing to do is
discontinue anesthetic administration. Then check for airway
patency. Artificial ventilation can be performed through the
nostrils or through the endotracheal tube by compressing the
rebreathing bag on the anesthetic machine or the use of a manual
resuscitator bag. An ear syringe can make a good rodent
resuscitator, as it fits right over the nose of larger rodents.
If the failure was caused by a narcotic, reversal agents may be
used. Other drugs such as doxapram can be used to stimulate the
respiratory system.
Causes of circulatory arrest include drugs, hypoxia,
hypercapnia, changes in the vascular volume or bed, deleterious
reflex responses, obstruction of venous return, severe
electrolyte imbalance, and primary cardiac pathology. Careful
maintenance of ventilation is one way to avoid hypoxia and
hypercapnia. Changes
in vascular volume can be minimized through the use of good
hemostasis by the surgeon and adequate I.V. fluid volume
replacement by the anesthetist. Surgeons must be careful when
moving abdominal contents around, not to place too much pressure
on the posterior vena cava and thus impede blood flow return to
the heart. Electrolytes can be monitored and imbalances
corrected during surgery before they get to the life threatening
stage. In some cases the presence of primary heart pathology may
be identified in a routine presurgical physical exam.
Signs of cardiac failure are white or cyanotic mucous
membranes, no pulsation in major arteries, no wound bleeding and
no palpable heart beat.
Treatment of cardiac arrest begins the same way as that for
respiratory arrest and includes discontinuation of anesthetic
administration, checking for a patent airway, and the
administration of oxygen. If possible, also lower the cranial
end of the animal by 30 percent. Closed chest massage can be
done by compressing the thorax by one third to one half its width
or depth at a ratio of 5 compressions to each ventilation. Fluid
replacement should occur as rapidly as possible. Drugs such as
epinephrine, sodium bicarbonate, prednisolone sodium succinate,
calcium chloride and lidocaine can be used but vary with
different situations which may be hard to define without the use
of an electrocardiogram.
Anyone performing frequent anesthetic procedures should have
a well-stocked emergency kit handy with such items as
endotracheal tubes, a manual resuscitator bag, syringes and
needles, and some or all of the drugs mentioned above. It is
helpful to have a card in this kit which list all the dosages for
these drugs to insure proper usage during the rare occasion when
they are needed. Frequent emergencies are an indication of
improper anesthetic or surgical techniques and should be reviewed
with the veterinarian to ascertain a possible cause and implement
a potential solution.
CONTROLLED SUBSTANCES
Many anesthetics, analgesics and tranquilizers are controlled
substances. They are divided into five schedules based upon their
abuse potential. Schedule I drugs are those with a very high
abuse potential for which there is no medical use. Schedule II
drugs also have a high abuse potential but are accepted for
medical use. This schedule includes agents with narcotic,
stimulant or depressant actions such as morphine, codeine,
meperidine, oxymorphone, pentobarbital, cocaine and opium.
Schedule III includes some of the barbituric acid derivatives.
Schedule IV has phenobarbital, chloral hydrate, and diazepam.
Schedule V agents are those with narcotics in limited quantities
such as antitussives and antidiarrheals. This is only a partia 
