ESSENTIALS FOR ANIMAL RESEARCH A PRIMER FOR RESEARCH PERSONNELProvided by the Animal Welfare
Information Center
|
Second Edition
By
B. T. Bennett
M. J. Brown and
J. C. Schofield
United States Department of Agriculture
National Agricultural Library
Beltsville, Maryland
Revised October 1994
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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 ever-changing 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 second edition of this manual has been updated to reflect recent changes in the regulations, the report of the 1993 AVMA Panel on Euthanasia and the expanded resources and services of the Animal Welfare Information Center.
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 and Ms. Lisa Halliday and Ms. Doris Thomas for their assistance in preparing the second edition. 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.
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 Associa- tion for Accreditation of Laboratory Animal Care (AAALAC)
2. Requirements of Individual Users
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 46 Veterinary Medical Officers working under one of the four REAC Sector Supervisors. The Sector offices are located in Fort Worth, Texas, Tampa, Florida, Annapolis, Maryland, 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 strengthened the standards for providing laboratory animal care, increased enforcement of the Act, provided for collection and dissemination of information to reduce unintended duplication of experiments using animals and mandated training for those who handle animals.
The 1985 amendment to the AWA also included 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 has 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 required 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 designated this committee as the Institutional Animal Care and Use Committee (IACUC) and charged 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 euthanatized 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 procedures and that the work being proposed does not unnecessarily duplicate previous experiments. To provide assurance of the former 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 mainte- nance and experimentation.
2. Research or testing methods that mini- mize 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 ani- mal care and treatment should be re- ported.
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.
The most recent amendment to the AWA (PL 101-624) was passed in 1990 and was entitled the Pet Protection Act. The regulations developed to implement this amendment define the minimal holding period for animals in pounds and shelters that are sold to dealers, and establish record keeping requirements for dealers who obtain dogs or cats from these sources.
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 September 1991, the instructions for completing 398 can be found in two locations within the application package. On page 13 the research investigator's responsibilities for assuring compliance with the PHS Policy are clearly addressed. Detailed instructions for completing Section 6 of the Research Plan which describes the use of Vertebrate Animals can be found on page 23.
The institution is responsible for maintaining all the necessary records to document compliance with the PHS Policy and for filing annual reports developed by the IACUC 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.
American Association for 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 39 professional organizations. An 18-member Board-appointed Council on Accreditation along with four scientific/technical panelist make 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 program inclusive of selected animal usage areas. Should deficiencies be identified in a previously accredited program, the institution is either granted a definied 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.
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.
Application for Public Health Service Grant, PHS, 398. Revised September, 1991. 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, 1978, Part II, pp. 59986-60026.
Public Law 99-198. Code of Federal Regulations, Title 9, subchapter A, Animal Welfare 1989.
Townes, J. Federal Regulations an Overview, Lab Animal, July-August 1980; 9:4 l6-22.
Chapter 2
Alternative Methodologies
B. Taylor Bennett, D.V.M., Ph.D.
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 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 & 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.
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 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.
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 euthanatized 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 animals, 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 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 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 microliter 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 euthanatize 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.
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.
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 & 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, & 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 & 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. 1976. Animal Models of Thrombosis and Hemorrhagic Diseases, NIH Publication No. 76-982.
Chapter 3
Animal Care and Use: A Nonexperimental Variable
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.
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 macroenvironment. 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; however, 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 microenviron- mental 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 rabbits 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 effect of shipping animals can be a significant physiological stress. Studies have documented the 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 nonexperi- mental 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 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.
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 potential 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.
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.
Allert, J.A.; Adams, R.A.; and Baetjer, A.M. l968. Role of environmental temperature and humidity in susceptibility to disease. Ach. Environ. Health 16:565-570.
Broderson, J.R., et al. 1976. The role of ammonia in respiratory mycoplasmosis of rats. American Journal of Pathology 85:115-130.
Davis, D.E. 1978. 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. 1982. Influence of cage shelf level on retinal atrophy in mice. Lab Animal Science, 32(4):353-356.
Lang, C.M. and Jessell, E.S. 1976. Environmental and Genetic Factors affecting laboratory animals; impact on biomedical research. Federal Proceedings Vol. 35 No. 5-8, 1123-1165.
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 & Co., Ltd., London, 1959.
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 animals 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 uncompromi-sed animal even though they are 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.
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.
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 Anesthesia - Temporary, controllable and reliable loss of 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 Effect - Time between initial administration and onset of the 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.
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.
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.
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.
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-VetR 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-VetR 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., I.M or I.P. 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, such as thiopental, which is much faster acting. 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 eyes. 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 postoperative 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 anesthetic 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 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