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Weichbrod RH, Thompson GAH, Norton JN, editors. Management of Animal Care and Use Programs in Research, Education, and Testing. 2nd edition. Boca Raton (FL): CRC Press/Taylor & Francis; 2018. doi: 10.1201/9781315152189-31
Management of Animal Care and Use Programs in Research, Education, and Testing. 2nd edition.
Show detailsBackground
Veterinary care is a critical component of an animal care and use program, as the health and well-being of animals used in research, testing, and teaching are essential to humane and reproducible science. Although the veterinarian has primary responsibility for the well-being and clinical care of the animals, veterinary care is a team event, which spans the life of each animal and encompasses all aspects of their care and use.
The regulatory requirement for adequate and timely veterinary care and oversight is delineated in the standards developed by many countries (e.g., United States and Canada) or state unions (e.g., European Union) (European Commission 2007; Zurlo et al. 2009; CIOMS/ICLAS 2012; Bayne et al. 2014). It is the goal of program management to ensure that the provision of veterinary care is not only timely and of high quality, but also sufficiently dynamic to meet the needs of the research, testing, and/or teaching being conducted. A program of adequate veterinary care must be provided regardless of the number of animals used or the size of the research program and sponsoring institution.
Roles and Responsibilities
Although specific responsibilities vary, every individual in the animal care and use program shares the responsibility to ensure the health and well-being of the animals used in support of the program. Clear and timely communication among individuals at all program levels is imperative to ensuring that this duty is met. The specific responsibilities related to veterinary care are as follows.
Institutional Official
The institutional official (IO) bears responsibility for the entire program, which includes indirect responsibility for veterinary care. The official must ensure that the veterinary care program meets the highest quality and most ethical standards as directed by its country, state, and local laws, policies, and regulations. He or she must ensure the availability of resources (e.g., space, personnel, and funds) for both the routine operation of the veterinary care program and unforeseen circumstances (e.g., disease outbreaks and disasters). It is the IO’s responsibility to ensure that the attending veterinarian is trained, qualified in laboratory animal care, knowledgeable and experienced. In addition, IO must make certain that the attending veterinarian, or other official designated to manage the program, has the authority to manage the program and that the veterinary care provided is both accessible and sufficient to meet the requirements of the program (AWR 9 CFR §1.1; CCAC 2008, Item 7; European Parliament and the Council of the European Union 2010). Clear, concise, and timely communication between the IO, attending veterinarian, and institutional animal care and use committee (IACUC) is critical to ensuring the overall program direction and animal well-being.
Institutional Animal Care and Use Committee
The IACUC or its equivalent is responsible for the assessment and oversight of all components of an institute’s program, including the veterinary care program. The committee plays a key role in supporting the attending veterinarian in ensuring the health and well-being of the animals used, in addition to serving as a line of communication throughout the program.
Scientist
The scientist is responsible not only for the use of animals, but also for their well-being. It is highly recommended, and required in some countries (Bayne et al. 2014), that a scientist consult and/or collaborate with the veterinarian in both the design and execution of all projects using animals. It is important that the veterinarian be provided with a clear understanding of how the experimental manipulations may affect an animal’s physical, physiologic, or behavioral appearance or profile. Anticipated changes to an animal’s behavior, physiology, or appearance should be clearly outlined in the investigator’s animal study proposal. Other information that is critical for the veterinary care team to understand includes, but is not limited to, (1) unique phenotypes related to each experimental model, line, strain, or species (e.g., diabetes and neurological behaviors); (2) experimental and humane endpoint criteria; (3) limitations to intervention strategies related to pain, stress, or distress; (4) limitations to medical treatments (e.g., steroids and antibiotics); and (5) limitations to palliative intervention strategies (e.g., special food, bedding, enrichment, and social housing). Because unforeseen complications may evolve throughout the course of a study and life span of an animal, continued communication with the veterinarian and veterinary care team is critical to ensure the well-being of the animals.
The scientist is also responsible for ensuring that all animal procedures and surgeries are conducted by trained, skilled personnel in accordance with his or her approved animal study protocol, which should include all current standards of veterinary care. In addition, the scientist is responsible for the safe use of hazardous agents and creating a safe work environment for his or her research team, including those who care for the animals.
Veterinarian
Regulatory Requirements
In the United States, the Animal Welfare Act is promulgated by the U.S. Department of Agriculture (USDA) as outlined in Animal Welfare Regulations (AWR) published in the Code of Federal Regulations (CFR) (AWAR 2013), Title 9, Animals and Animal Products, Subchapter A, Animal Welfare. The AWR defines the attending veterinarian as follows: “Attending Veterinarian means a person who has graduated from a veterinary school accredited by the American Veterinary Medical Association’s Council on Education, or has completed the American Veterinary Medical Association’s Education Commission for Foreign Veterinary Graduates certification program (https://www.avma.org), or has received equivalent formal education as determined by the Administrator; has received training and/or experience in the care and management of the species being attended; and who has direct or delegated authority for activities involving animals at a facility subject to the jurisdiction of the Secretary” (AWR 9 CFR §1.1). The quality or quantity of the required training or experience has not been specified by the USDA.
Program requirements have also been put in place by the European Union related to the designation of an advisory veterinarian with expertise on laboratory animal medicine for each animal breeder, supplier, and user (European Parliament and the Council of the European Union 2010, Article 25). But here again, like the USDA, the regulations do not specify the nature of the required expertise. The Canadian Council on Animal Care’s policy statement for senior program administrators responsible for animal care and use programs (CCAC 2008, 7.2) states that veterinarians providing clinical services and/or compliance oversight must have the experience and expertise necessary to evaluate the health and welfare of each species used in the context of the work being conducted by the institution.
In most situations, the veterinarian overseeing the care of research animals has completed several years of postdoctoral training under the oversight of an experienced veterinarian. Credentialing by a certifying organization (e.g., the American College of Laboratory Animal Medicine [ACLAM]) is a common way of providing evidence of a veterinarian’s knowledge and experience. Several countries have established colleges that set criteria and standards for obtaining board certification in the laboratory animal medicine specialty (i.e., Europe, Japan, and Korea) (http://www.iaclam.org/about.html). Additional information can be obtained from the International Council for Laboratory Animal Science (ICLAS) (http://iclas.org/) and the International Association of Colleges of Laboratory Animal Medicine (IACLAM) (http://www.iaclam.org/).
Although veterinary practice licensure in the state, country, province, or region may not be required to provide care to laboratory animals, licensure is commonly required to procure various pharmaceuticals and controlled drugs and sign health certificates. Veterinarians and institutions should review their local requirements for licensure and/or accreditation, which often vary by country, state, province, or region.
Veterinary Care Responsibilities
The attending veterinarian has ultimate responsibility for the day-to-day health and well-being of all animals in the program. The range and scope of these responsibilities are outlined in the Guide for the Care and Use of Laboratory Animals (Guide) (NRC 2011), as well as several position (AAALAC 2016; ACLAM 2016) and policy statements (CCAC 2008; European Parliament and the Council of the European Union 2010; USDA 2016). The size of the institutional program and nature of the research being conducted will determine whether a full-time, part-time, or consulting veterinarian is required. When the veterinarian is less than full-time, an explanation of his or her role and a prearranged schedule of when he or she will visit the animal facility should be available in writing (AWR 9 CFR §2.33 a.1). Regardless of the time commitment, the veterinarian’s accountability remains the same. The definition of what is determined to be “adequate veterinary care” may vary between jurisdictions or countries. Adequate veterinary care guidelines commonly define the requirement for veterinary access to the animals and their records, the frequency of veterinary visits and observations, and the provisions for providing timely and appropriate medical care (ACLAM 2016).
A veterinarian is a required member of the IACUC and should have specialized training and/or experience in the field of laboratory animal medicine and science. The veterinarian serves as the subject area expert for issues ranging from animal husbandry to the identification and alleviation of pain and distress. The scientist should take advantage of the veterinarian’s expertise during protocol development and consult with him or her throughout the protocol review and implementation process. Many IACUCs require that the institutional veterinarian be consulted on the proper use of anesthetics, analgesics, and euthanasia agents. At a minimum, the veterinarian should be involved in the review and approval of the final protocol.
The veterinarian works closely with the IACUC to facilitate the postapproval monitoring of approved animal study protocols, as well as the daily monitoring of all animal care and veterinary care program operations. The veterinarian plays a key role in the identification and prevention of occupational health and zoonotic issues. He or she must work closely with program managers and safety specialists in the development of standard operating procedures (SOPs) to mitigate or remove occupational health risk factors. The veterinarian has a role in training the investigative and animal care staff, as well as clinical support personnel. The training should include, but not be limited to, animal procurement, transportation, identification, handling, husbandry, preventive medical care, veterinary care, chemical sedation and anesthesia, sterile and aseptic surgical techniques, analgesia, euthanasia, and recognition of species-specific signs of pain or distress.
The experienced laboratory animal veterinarian is an important member of the management team and should be included in discussions of issues that involve the animal care program and animal holding facilities. Facility construction, caging purchases, investigator-purchased equipment, and so forth, can directly or indirectly impact animal health and well-being or increase disease spread risk.
Animal Facility Manager, Office Manager, and Administrative Staff
Like the veterinarian, the animal facility manager has delegated accountability for daily oversight of the facility’s animal care and use program. The manager works closely with the veterinarian to ensure the health and well-being of all animals. The office manager and administrative staff oversee administrative functions (e.g., procurement and staffing) that are critical to keeping the program operating in an efficient and productive manner. Responsibilities of the facility manager, office manager, and administrative staff that directly impact the health and well-being of all animals include, but are not limited to, (1) facility access; (2) monitoring animal importation, exportation, and procurement; (3) oversight of routine animal health and behavior evaluations; (4) ensuring execution of established facility sentinel programs; (5) ensuring availability of required supplies and pharmaceuticals; and (6) oversight of the management of animal health records. In addition to the above, the facility manager works with facility engineers to ensure the integrity of facility mechanical and support systems (e.g., heating, ventilation, and air-conditioning [HVAC] and environmental control systems).
Veterinary and Laboratory Animal Technician or Technologist
Today, there is a movement to rename the veterinary and laboratory animal technician or technologist as a “veterinary nurse” (Hyde 2016; Kennedy 2016; http://www.bvna.org.uk/publications/veterinary-nursing-journal). The veterinary and laboratory animal technicians and technologists work closely with the veterinarian to provide nursing and diagnostic support to ensure the health and well-being of all animals in the facility or program. In addition to their health care responsibilities, the veterinary support staff commonly executes the facility’s sentinel programs, monitors and orders required supplies and pharmaceuticals, manages animal health records, and may assist scientists with technical aspects of their research. Having additional specific training in the biology, husbandry, health, surgical care, and medical treatment of animals makes the veterinary technician or technologist a real asset. Veterinary technicians commonly hold an associate’s or bachelor’s degree in veterinary technology and may be designated as a “Registered” (RVT), “Certified” (CVT), or “Licensed” (LVT) Veterinary Technician. Individuals receiving their training through the U.S. military receive a 91T or 68T certification in veterinary technology. As technicians gain experience and training in laboratory animal science, they may choose to become certified by the American Association for Laboratory Animal Science (AALAS) as an assistant laboratory animal technician (ALAT), a laboratory animal technician (LAT), or a laboratory animal technologist (LATG) in the United States.
Animal Care Specialist or Husbandry Technician
The animal care specialist or husbandry technician is one of the most important contributors to the veterinary team. They are the “eyes and ears” of the program, being the first line of defense in detecting problems with an animal’s health and well-being. The specialist is trained in laboratory animal husbandry and is the daily caretaker for all animals under their oversight. As a result of their daily observations, animal care specialists gain extensive knowledge of the normal physiology and behavior for each animal and are skilled at ascertaining what is “normal” for the animals under their care. They are trained on the recognition of subtle departures from normal, which may indicate the onset of a problem or disease. Upon identifying a problem, it is the animal care specialist’s responsibility to report the change in the animal’s condition to the technician or veterinarian for further evaluation. The specialists’ contribution is also critical to the execution of many research protocols, as they are often responsible for the provision of protocol-approved special diets, food or fluid restriction, and the administration of study or therapeutic agents (e.g., analgesics).
Veterinary Care: Impact of Facility Design, Medical Support Areas, and Equipment
The requirement for medical facilities and equipment will vary with the species and research to be accommodated in a program. Program size, complexity, and location and dissemination of key program elements also play a role in determining the design of program spaces (Howard and Foucher 2008), the equipment required, the need to duplicate resources, and the amount of storage space required for normal operations and emergencies. As a general rule, rodent and aquatic facilities require less specialized veterinary medical support features than larger species, such as carnivores, nonhuman primates (NHPs), rabbits, and other nonrodent mammals. The requirements for specialized facility design features and specific equipment related to veterinary care may also vary with the country, province, or region in which the work will be conducted.
Several components of facility design (e.g., differential air pressures and surface composition) have a direct impact on a program’s ability to exclude or limit the dissemination of pathogens (Hrapkiewicz et al. 2013). Attention must also be paid to personnel and equipment flow patterns, sanitization schedules, and chemical agents used (Shek et al. 2015). Pathogens do not move on their own, but rather attach themselves to airborne particles (e.g., dust), fomites, and surfaces (Brachman 1996). Therefore, if the movement, accumulation, and spread of dust are controlled, and surfaces are kept clean and sanitized, the transmission of most pathogens can be controlled. Being nonporous, free of cracks and crazing, nonabsorbent, and resistant to chemicals facilitates the ability of facility surfaces to be cleaned and sanitized (Leverage and Roberts 2009).
Since personnel, supplies, and equipment can also serve as fomites (NRC 1991; Becker et al. 2007; Clifford and Watson 2008; Watson 2013) for the transport of potential pathogens throughout the facility, these factors should also be taken into consideration when designing a facility. Whenever possible, the design should facilitate the movement of personnel and equipment from “clean” or low-risk areas (e.g., clean cage wash, food and bedding storage areas, and pathogen-free animal holding areas) to “dirty” or areas of higher risk (e.g., dirty cage wash, and isolation and quarantine areas) (Hessler 1991) rather than the reverse. In addition, attention should be given to the personal protective equipment (PPE) required in various areas of the facility to allow for the presence of gowning areas, as well as areas for the storage and disposal of PPE.
It is common to have procedure rooms or laboratories that are used to support both research and veterinary and technical procedures. Appropriate measures and SOPs must be in place to minimize the transmission of pathogens between animals while they are in these areas and exposed to common equipment. This includes, but is not limited to, surfaces where animals are used, restraint devices, scales, anesthetic machines, and equipment for behavioral studies. Policies should be developed to clearly define when animals must be handled in biosafety cabinets (BSCs) or other equipment designed to prevent the dissemination of pathogens or biological agents.
Experimental Testing and Procedure Rooms
It is important to determine if the two-way movement of animals will be permitted between the housing facility and research laboratory. If the movement of animals is restricted, the design of the animal facility should include space for the experimental manipulation of the animals. However, from a veterinary health perspective, testing and procedure rooms constitute a risk factor for the spread of pathogens from people to animals and between research animals (Koszdin and DiGiacomo 2002; Hrapkiewicz et al. 2013; Shek et al. 2015). Here again, attention to the selection of equipment, surfaces, and design of the area to facilitate cleaning and sanitation will help prevent transmission of pathogens. Closed cabinets and storage drawers help to prevent the collection of dust on porous, hard-to-sanitize supplies and materials. Consideration should be given to the presence of running water, a refrigerator, a freezer, a fireproof safety cabinet, a BSC (e.g., BSL-2 and BSL-3), a chemical fume hood, and CO2 euthanasia chambers within the procedure area. If anesthesia will be conducted in the area, the need for oxygen and anesthetic gas scavenging (e.g., downdraft tables, snorkel scavenging systems, and passive vacuum systems) should be evaluated. If necropsies or whole-animal perfusions will be conducted, adequate ventilation systems must be present. The installation of a commercially available flushing downdraft necropsy table and systems for the collection of excess fixative to prevent contamination of public water and sewer systems should also be evaluated. Where animals will be handled on open countertops, room air pressure is commonly neutral or negative to the outside corridor. All personnel using the rooms must be educated on disease spread and their responsibility in maintaining clean and sanitary working conditions.
Medical Treatment Facilities
Ideally, an area for the treatment of sick animals should be included in the design of the facility. The size of the program, species being used, and type of research being conducted will play a role in the size and design of the treatment area. A dedicated treatment and holding area, separate from common procedure and holding rooms, may be warranted for the treatment of animals with potentially contagious conditions. Treatment areas should be easily cleaned and sanitized as outlined above for experimental procedure areas. Treatment areas should contain the equipment and supplies required to treat animals without contaminating other areas of the facility or leaving an animal unmonitored to collect needed supplies.
Diagnostic Facilities
Imaging Facilities
Like many veterinary practices around the country, it is becoming common for laboratory animal facilities to have in-house medical imaging capabilities. The requirement for and extent of in-house imaging is determined by the size of the program, the species being used, and the kinds of research being conducted. Where in-house facilities are not possible or limited, provisions should be made to obtain the needed support from outside programs.
Today, in-house imaging capabilities may include radiography, ultrasound, positron emission tomography/computed tomography (PET/CT), x-ray reconstruction of moving morphology (XROMM), magnetic resonance imaging (MRI), fluorescence imaging, and nuclear medicine. Because the imaging equipment and facility can function as a fomite to disseminate pathogens, SOPs must take into account the ease and thoroughness of routine sanitation, as well as the flow patterns of both animals and personnel. Specialized programs must be in place for training personnel who perform or attend imaging procedures, and depending on the potential exposure risk, personnel exposure monitoring may be required. Personnel protective devices such as lead aprons and gloves should be stored in a manner that maintains the integrity and functionality of the material. Personnel protective garments should be routinely evaluated to ensure their functionality. The process consists of visually and manually inspecting the garments for wrinkles, cracks, crazes, or other deteriorations, followed by testing the items using radiography. There are commercial companies that will certify the functionality of lead aprons and gloves. The purchase of preventive maintenance agreements for imaging equipment is often beneficial to keep equipment in good repair and ensure safe operation.
Depending on the research being conducted and animal models used, imaging facilities may also require additional procedural areas for animal preparation and clinical support. This is of particular importance in clinical emergencies or where invasive procedures are required for the introduction of tracers or dyes.
Clinical Pathology Facilities
Diagnostics (e.g., clinical chemistry, parasitology, bacteriology, and serology) can be provided in-house or through outside programs. In many situations, a combination of the two approaches is the best solution. In-house diagnostics offer fast results, but care must be exercised to confirm that the test results are valid for the species being tested. Reagents and equipment must be maintained in accordance with the manufacturers’ instructions. An appropriate quality assurance program must also be in place to certify the validity of the test results. Personnel must be trained in sample handling, equipment operation, maintenance, and identification and resolution of problems. At a minimum, the clinical pathology area should be equipped with running water, a refrigerator, a freezer, and sufficient space for sample handling, equipment operation, and storage of supplies. Inclusion of a fireproof cabinet, BSC, chemical fume hood, and CO2 euthanasia chambers within the area may be useful.
Necropsy Facilities
A dedicated necropsy area is advantageous for the containment of potential pathogens or known agents requiring a higher biosafety level. Containment and sanitization are key functional components of this area. As with most areas within an animal facility, it is critical that surfaces should be smooth, impervious to water, and free of cracks, crazed areas, and ledges. Specially designed flushing necropsy tables address all these requirements. Many necropsy tables are designed to also serve in downdraft mode with directional airflow to facilitate the containment of airborne particles. If the table will be used for perfusion of tissues with formalin or other fixatives, the table can be modified to collect the unused fixative as chemical waste. Necropsy rooms must be equipped with running water, soap, and paper towel dispensers. Necropsy equipment and instruments should be dedicated for that area. SOPs should be developed that outline the required PPE, traffic flow patterns, and decontamination procedures. Consideration should be given for the disposal of medical or pathological waste, as well as the dedicated refrigeration and storage of unneeded tissue and cadavers prior to disposal. Because infectious agents may be the cause of an animal’s death, it is critical that the necropsy area be isolated and the personnel traffic patterns controlled to prevent cross-contamination. Unless separated from the main animal facility, the room air pressure must be negative to the outside corridor.
Pharmacy Facilities
The size and complexity of the pharmacy will depend on the size of the program, species being utilized, and research being conducted. Whether the pharmacy is a lockable cabinet or drawer in a procedure or treatment area or a dedicated room, the requirements are the same. The pharmacy must provide an environment that protects the integrity of the pharmaceuticals and allows for their security. Drugs that are governmentally regulated (i.e., controlled substances) must be stored in a securely locked, substantially constructed cabinet. These substances must be kept in a secured location that is accessible only to a minimum number of authorized individuals. Larger quantities of controlled substances may require storage in a safe. Smaller quantities of controlled substances can be stored in procedure and treatment areas, in a double-lock and double-door narcotic cabinet or other secured lockbox, for which the keys or combinations are available to only authorized personnel. Some factors that must be considered when evaluating the storage of controlled substances include (1) the regulations applicable to the storage location, (2) the number of individuals requiring access, (3) the required location of the drugs, (4) the presence of alarm systems, (5) the quantity to be stored, and (6) the past history of thefts or diversions (U.S. Department of Justice 2016).
Surgical Facilities
The facilities required to conduct rodent and aquatic survival surgery are less extensive than those that are required for other animals. For most rodents and aquatics, surgery may be performed in a laboratory or facility procedure room as long as the area being used is dedicated at the time for that purpose and appropriately managed to minimize contamination from other activities within the room during the surgery. The chosen area should be free of clutter, easily sanitized, and in a low-traffic and low-noise area of the room.
The size of the program, species to be used, and nature of the procedures being conducted will all play a role in determining the size and complexity of the surgical facility required for other species. For most species other than rodents and aquatics, a defined facility is required. At the minimum, three distinctive functional rooms or areas are needed: (1) an animal preparation area, (2) a surgeon preparation area, and (3) a dedicated surgery area (NRC 2011). Other functional areas that are commonly present include a dedicated recovery room, storage space, and instrument preparation areas. The surgical suite should provide easy access to emergency response equipment (e.g., defibrillator) and supplies.
Surgical suites are often maintained under positive air pressure to help minimize the movement of potentially contaminated airborne particles into the dedicated surgery room (NRC 2011; Perkins and Lipman 2014). While nonsurgical use is discouraged, if a surgical facility is used for other purposes, thorough decontamination is required prior to reuse for surgery (NRC 2011; Perkins and Lipman 2014).
Postoperative Recovery and Intensive Care Facilities
A dedicated area for postoperative recovery is not routinely required for most rodent and aquatic species, although these species still require close monitoring and support after surgical procedures. A dedicated area in which the animal can be closely monitored and supported throughout the postoperative period is required for other animals (NRC 2011). In many facilities, this area also supports animals that require intensive care. Intensive care areas are commonly equipped with specialized caging that is designed to provide a supportive environment for the animal and, in some cases, also prevent self-injury during the postoperative recovery period. Intensive care caging often allows for the provision of oxygen, fluid, medical, and thermal support to the recovering or convalescent animal. The area should be designed to support the electrical requirements of the various monitors, infusion pumps, and other equipment. The area is commonly stocked with supplies, pharmaceuticals, and equipment needed for treating medical emergencies.
Isolation and Quarantine Facilities
Although isolation and quarantine facilities are covered in detail in Chapter 30, it is important to understand their importance to the veterinary care program. Animals are quarantined when they are known to carry or may potentially carry a contagious organism that could adversely impact the health of other animals (Carty 2008). Animals are isolated for a variety of veterinary and research reasons. For example, pathogen-free animals are routinely isolated to verify or protect their “clean” health status. This is common practice with immunodeficient animals, such as nude and SCID mice. The amount of isolation and quarantine space required by a program will depend on the program’s need to obtain animals from sources that may not meet the pathogen-free status established for the facility (Rehg and Toth 1988; Roberts and Andrews 2008) or the research being conducted. It is critical that appropriate SOPs be developed that define biocontainment practices, procedures, personnel traffic patterns, and PPE to prevent contamination and cross-contamination of critical areas within a facility. Some programs may require personnel to enter isolation and quarantine areas last or take a shower before entering or leaving the area. Facilities of this nature will require the proximity of a locker room and shower. Anterooms provide a convenient area to don or remove required PPE when entering or leaving the area. Personnel working in isolation and quarantine areas must be highly trained in disease transmission and containment to prevent cross-contamination. Isolation and quarantine facilities should have tightly controlled access and often require dedicated supplies, equipment, and autoclave support. The autoclave should be of an appropriate size and be in close proximity to the isolation or quarantine area. Some facilities are designed for a double-door, “pass-through” autoclave to process material into or out of the area. In general, the room air pressure of isolation and quarantine areas is kept negative to the outside corridor when housing animals that potentially harbor pathogens (NIH/ORF 2008; Huerkamp and Pullium 2009; Lipman et al. 2015). When isolating animals to protect their clean health status (immunodeficient animals, etc.), the room air pressure is often maintained positive to the outside corridor (Kowalski et al. 2002; Lipman et al. 2015).
Veterinary Programs
Animal Health Status
Establishment of the Required Animal Health Status
Using healthy animals is a foremost consideration in conducting sound research and generating quality data. Health status requirements will vary depending on the nature of the planned research (Baker 2003; Desrosiers 1997) and must be determined before the animals are acquired. In addition, it also must be determined if the program has the ability to maintain the health status once the animals are received. Advances in facility design features, health monitoring techniques, and specialized shipping containers allow vendors to meet a wide range of research demands. Institutionally managed breeding programs may also contribute to achieving and maintaining the appropriate animal health status. The researcher must make sure the animal procurement staff understands any nonstandard health requirements essential for the research (e.g., need for immunocompromised animals in tumor studies, and Helicobacter species–free mice in gastrointestinal [GI] research). It is important that research staff discuss any health requirements with the vivarium’s management and veterinary staff prior to the acquisition of the animals. They can help the investigator determine if the vivarium has the capability to maintain the desired animal health status, as well as meet the other requirements of the research.
Research Requirements
There is no universal menu of research requirements. Each research project requires consideration of parameters unique to the given research. For instance, using immunocompromised animals may be appropriate for tumor studies but be quite inappropriate for infectious disease studies where a robust immune response is required. Clear communication between the researcher and the veterinary staff is paramount and should be initiated by the former. In the ideal world, every aspect of a research project should be understood by all parties involved. At a minimum, the veterinary staff needs to be familiar with the species to be used and understand the aspects of the research project that may potentially impact the health and well-being of the animals involved. Such aspects can vary from simple parameters, such as gender, weight, age, and coat color, to more complex aspects, such as strain and specific body condition (e.g., lean vs. obese). Unique features, such as physical and physiological fitness, size and functional capacity of select organs (e.g., liver and prostate in males), ability of the host immune system to fight off challenges (e.g., microorganism infection and foreign organ or tissue tolerance or rejection), spontaneous mutations, and propensity for developing endocrine imbalances (e.g., diabetes), may also be important factors to take into account.
Species Considerations
Rodent Colonies
Establishment of the appropriate health status for a rodent colony or facility is dependent on many factors. First is the potential impact of animal health status on the research to be conducted. Some studies have strict requirements for germ-free animals, whereas other studies are less susceptible to the effects of pathogens. In studies requiring pathogen-free animals, further questions must be asked to determine which organisms may adversely impact the research. The term specific pathogen-free (SPF) is commonly used in laboratory animal science to describe animals free of a defined specific list of pathogens. The term SPF itself does not indicate that an animal is free of particular pathogens of interest to the receiving facility or researcher. Each vendor or facility supplying animals must be asked which pathogens they exclude from their animals. It is the receiving program’s responsibility to ensure that the vendor’s list of organisms they test for includes the pathogens of importance to their facility and research. In addition, the vendor or supplying facility should be asked how the current health status of their animals is determined and how the status is ensured throughout their production program, as well as during shipment of the animals to the institution.
The second factor that must be considered is what health status is optimal for the general health and well-being of the receiving colony or facility. Ideally, most program managers and veterinarians would like their animals to be free of all viral, bacterial, and parasitic pathogens, but in reality, there is a direct relationship between the colony’s desired health status and the cost of doing business within a facility. Generally, the “cleaner” the desired colony health status, the higher the cost of maintaining the colony becomes. The higher cost results from several factors, including the need for (1) specialized caging and equipment, (2) increased levels of PPE, and (3) more intensified health surveillance programs. There is also a cost to the research, in that clean facilities often must not allow animals to return to the home facility after being tested or studied in another facility or laboratory. In addition, maintaining a clean facility may limit the ability to import animals from other collaborating research groups or vendors. Therefore, careful consideration must be given to the pathogen exclusion list developed for the colony or facility (Mähler et al. 2014; Shek et al. 2015). Programs must consider not only the health impact on the colony and the research, but also the final cost to maintain the desired health status (Kowalski et al. 2002).
Maintaining animals of a specific health status can require specialized barriers designed to exclude pathogens (Nicklas et al. 2015). Therefore, a third factor to consider is the available infrastructure and personnel to support the desired health status (e.g., flexible film isolators, microisolator caging, BSCs, and autoclaves), as well as the availability of funds to maintain the program.
Nonhuman Primates (New and Old World)
At a minimum, NHPs need to pass a physical examination and undergo a period of quarantine to determine the presence of zoonotic agents. A 31-day quarantine is required at the port of entry into the United States (42 CFR §71.53) to safeguard against the introduction of tuberculosis-infected animals. During the quarantine, a thorough physical examination and clinical workup is conducted by a qualified veterinarian and the animal is administered a series of three intradermal tuberculosis tests. Depending on the future destination of the animal, additional tests and requirements may include combinations of the following:
- Dental exam
- Thoracic and abdominal radiology
- Hematology and serum biochemistry
- Internal and external parasitology
- Bacterial cultures: Fecal (Shigella sp., Salmonella sp., Campylobacter sp., and Yersinia sp.), nasal, oropharyngeal, tracheobronchial
- Virology: Macaque profiles”“Macacine herpesvirus 1 (McHV-1; formerly herpes B), measles, retroviruses (Tardif et al. 2012); baboon SA8; and Herpesvirus tamarinus (marmosets)
- Urinalysis
- Vaccination status: Measles, rabies, and tetanus
- Neurological assessment: Hand and eye coordination and motor functions
- Psychological assessment: Basic (e.g., shy vs. aggressive, willingness to work with humans, advanced capacity for and ease of learning, memory, and suitability for complex research tasks)
- Tests focused on specific criteria required for a particular field of research: Specific phenotypes (e.g., obese, diabetic, or blind), congenital abnormalities (e.g., dwarfism, scoliosis, or polydactyly), or special conditions (e.g., amputees or geriatric)
- Genetic profiles
When available, information on the origin of the animal (e.g., wild or captive bred, country of origin), may be valuable. Additional information that may prove valuable is the animals’ past social housing, medical, surgical, or research history.
There are several well-established NHP vendors offering a variety of species, ages, and sizes of monkeys with a specified health status, for example macaques free of retroviruses and McHV-1. While some institutions have adopted policies requiring their researchers to work exclusively with zoonosis-free animals, other institutions have created carefully crafted procedures for working successfully with populations of macaques known to be infected with zoonotic organisms (Mansfield 2005) or that convert to a positive status during a study. Programs must understand that even SPF animals, believed to be free of McHV-1, have been known to seroconvert and shed the virus (Weigler 1992). The only definitive, but not practical, method to ensure an animal does not carry the virus is postmortem testing. Therefore, starting with serologically negative animals can be helpful to prevent human exposures, but appropriate SOPs, including the use of PPE (see Chapter 14), must still be implemented to work safely with these animals.
Carnivores
Most U.S. institutions use purpose-bred (Class A) dogs. A USDA Class A license is issued to dealers who sell animals that were bred and raised at their facility. A Class B license is issued to dealers who buy and sell warm-blooded animals that were not born and raised on their property. At the U.S. Congress’s request, the National Academy of Sciences (NAS) assessed whether there is a scientific need for National Institutes of Health (NIH) grant recipients to purchase dogs and cats from random source (Class B) dealers. In May 2009, the National Research Council (NRC) released its report “Scientific and Humane Issues in the Use of Random Source Dogs and Cats in Research.” (NRC 2009b). The report concluded that there was not. Moreover, in response to congressional concern, the NIH has since advised its grant recipients that it is phasing out the practice (NOT-OD-11-055). Effective October 1, 2014, the NIH no longer funds research on dogs procured from pounds, breeders, and other so-called random sources (http://www.news.sciencemag.org/news/2014/10/nih-ends-funding-experiments-using-random-source-dogs).
Class A dogs, in addition to being clinically healthy, are typically inoculated with core vaccines (e.g., canine hepatitis/adenovirus type 2, canine distemper, rabies, and canine parvovirus virus) and noncore vaccines for canine parainfluenza virus (which some protocols still consider core), Bordetella bronchiseptica, borreliosis, and leptospirosis (Welborn et al. 2011). Class A dogs are bred in closed colonies, which are tested for external and internal parasites and checked for apparent physical and physiological abnormalities (Nemzek et al. 2015).
Use of Class A and B cats is scrutinized using the same criteria as dogs. Class A cats, like dogs, derive from operations geared toward generating non-pet, but socialized, healthy animals. Core vaccines include feline panleukopenia, feline rhinotracheitis, feline calicivirus, and rabies virus (AAFP 2013). Noncore vaccines include feline leukemia virus, feline infectious peritonitis virus, Bordetella, and Chlamydia. Such animals are also free of external and internal parasites and apparent physical and physiological abnormalities. Giardia vaccination is optional.
The domestic or European ferret (Mustella putorius furo) has unique applications for its use in research. Ferrets should only be procured from reputable research vendors with colonies of known health status. Unless restricted by research, it is recommended that ferrets be fully immunized against rabies and canine distemper virus (AFA 2006).
Lagomorphs
Conventional health status rabbits are prone to secondary bacterial infections caused mainly by Pasteurella multocida, which can prematurely terminate research projects (Deeb et al. 1990). There are a number of commercial vendors generating rabbits that are SPF for Pasteurella. These animals are significantly healthier than their conventional counterparts, allowing for successful completion of longer-term research projects. When housing Pasteurella-free animals in the same facility with animals known to harbor the bacterium, care must be taken to establish appropriate traffic patterns and PPE requirements to ensure that the Pasteurella-free animals are not inadvertently contaminated with the agent. Investigator requirements will determine the rabbit breed to be selected. For example, both albino and pigmented breeds are available, and some ocular work requires the use of pigmented eyes.
Agricultural Animals
Agricultural species include pigs and small ruminants. These species require special enclosures because of their size and rapid growth, and their care may be labor-intensive (FASS 2010). Pigs are social, large animals that are very popular in teaching labs and are robust enough to be used in chronic and invasive projects. Both conventional and SPF animals are available, and the demands of contemporary research have created a plethora of genetically selected breeds (mini- and micropigs), as well as transgenic hybrids (Swindle et al. 2012).
Small ruminants (sheep and goats) have an expanding use in the research setting, although they may harbor some zoonotic concerns, such as Orf (parapoxvirus) and Q fever (Coxiella burnetii) (Underwood et al. 2015). Animals free of Q fever are now commercially available. For more details, see Chapter 23.
Amphibians
Amphibians have long been utilized in scientific research and education as models for a variety of developmental and physiological processes, largely due to their unique ability to undergo metamorphosis and to regenerate limbs in some species. Their embryos have been used to evaluate the effects of toxins, mutagens, and teratogens. They have short generation times and genetic constructs desirable in transgenic and knockout technology, and also in genetic and genomic research. They are useful due to their sensitivity to climatic and habitat changes and environmental contamination. Amphibians begin life as aquatic larva and, through the process of metamorphosis, emerge as terrestrial adults. While many species of amphibians do not strictly adhere to this developmental pattern, they remain the only vertebrate class with such a unique adaptation (Zug et al. 2001; O’Rourke 2007). Xenopus laevis is the most commonly used species in the laboratory due to their ease in embryo manipulation and applicability to early pregnancy testing and developmental studies. There are well-established U.S. vendors (e.g., NASCO [https://www.enasco.com/page/xen_care]) that provide commercially raised animals with a known health status.
With the exception of African clawed frogs (Xenopus spp.) (https://www.enasco.com/page/xen_care and http://www.xenopus.com), amphibians are not available from research purpose-bred commercial breeding colonies. Bullfrogs (Rana catesbeiana) and other amphibian species may be purchased from pet suppliers, or must be collected from the wild by the investigator, or obtained from other researchers, zoos, or agencies, such as the U.S. Fish and Wildlife Service. It is essential to replicate their natural environments in order to breed them in a research setting. Unlike domesticated mammals, amphibians are wild animals and the husbandry methods used must take this into account (Browne et al. 2007). The welfare of the animals must have the highest priority in the design of animal rooms, tanks, and tank furnishings.
Fish
The most commonly used fish in research is zebrafish (Danio rerio). It is an attractive alternative to mammalian species for the following reasons: (1) it is less sentient than mammals, (2) it has ease of reproduction and fast-growing and developing offspring, (3) it can tolerate higher doses of chemical mutagenesis than rodents, (4) transparent embryos and larvae permit noninvasive imaging strategies when following genetic manipulation or pharmacological treatment, (5) it offers insights into human disease due to functional homology with mammals, and (6) it is easy to replicate the natural environment of zebrafish, which reduces stress and its impact on experiments. There are a number of websites with information on sales, tutorials, and genetics (e.g., http://www.zfic.org/linksindex.html). Zebrafish are available from established vendors (e.g., Zebrafish International Resource Center of the University of Oregon). Zebrafish embryos are commonly washed with hypochlorite solutions prior to their introduction into an animal facility to help remove unwanted pathogens.
Birds
A large number of avian species are used in laboratory animal science. The range spans many domestic species (e.g., chickens, ducks, geese, pigeons, doves, zebra finches, Japanese quail, and parakeets) and nondomestic species (e.g., crows, sparrows, and hawks) (Fair et al. 2010). Many species of birds are obtainable from commercial sources, whereas wild birds are often used solely in field studies. Avian models have advanced our understanding of developmental biology, aging, immunology, endocrinology, and genetics, as well as other aspects of medicine and science. Many avian species are highly social and should be socially housed whenever possible (Hawkins et al. 2003). When birds must be brought from the wild and maintained in the laboratory animal facility, they should ideally be housed separately from other species and, if possible, provided with conditions that approximate their natural habitat (Hawkins 2001). Along with temperature and humidity, consideration should be given to the type of food, perches, and cover provided, as well as the manner in which water is offered (NRC 1977). Both domestic and wild birds can be carriers of zoonotic diseases (e.g., chlamydiosis, cryptococcosis, histoplasmosis, psittacosis, and salmonellosis), as well as several infectious and parasitic agents that can adversely impact other birds (e.g., Newcastle disease, Marek’s disease, and coccidiosis) (Baer et al. 2015; Patterson and Fee 2015; Taylor et al. 2016).
Other Considerations
The desired health status of acquired animals may vary depending on the short- and long-term goals of each project. For instance, the animals needed for an acute project may require no or only a basic physical examination and short quarantine period, whereas those for lengthy projects, in most cases, require a thorough clinical evaluation and evidence of robust health during a period of quarantine. Similarly, the demand for clinical robustness will be much higher for invasive than noninvasive studies.
Acquiring Animals from Commercial Sources
The quality and health status of animals procured from vendors can vary greatly depending on the size and experience of the vendor. A program’s first priority should be to maintain its own biosecurity, protecting against the introduction of unwanted pathogens. Unfortunately, under most circumstances, it is neither possible nor cost-effective to test or rederive every animal entering a facility to ensure its health status, or to breed all animals in-house. Therefore, a relationship must be established with reputable vendors to provide healthy, pathogen-defined animals.
An ideal start to procuring any animal is to find out which vendors can meet the program’s requirements by talking to the animal facility veterinarian, manager, or other investigators with animals in the same facility. Additional sources of information include the materials and methods sections of published research similar to that being conducted in your program, laboratory animal association publications, and websites such as the AALAS Laboratory Animal Science Buyers Guide (http://laboratoryanimalsciencebuyersguide.com/) or the LabAnimal® Europe and Asia Pacific Buyers Guides (http://www.labanimaleurope.eu/buyers_guide/), as well as regional repositories. Many large vendors post their current health status online via their websites. Once potential vendors have been identified, inviting them to present a seminar on their company’s products and services, including their health surveillance program and quality assurance guarantees, can be helpful. In some situations, conducting a site visit to the vendor’s production facility may also be beneficial.
When establishing a long-term commitment with a vendor, developing a contract detailing the relationship between both parties should be considered. A contract can be used to detail a program’s requirement for availability of specific products, custom-designed surveillance panels, shipping specifications, product returns, reimbursements, and replacements. In addition, a contract can be used to detail the vendor’s responsibility to notify the program in a timely manner of any change in the health status of animals in its facility.
Acquiring Animals from Noncommercial Sources
With the advent of transgenic technology, it has become commonplace to acquire animals from noncommercial sources, such as universities or other research institutions. Noncommercial sources can vary widely in their health status and surveillance programs (Pritchett-Corning et al. 2009). When procuring animals from noncommercial sources, an assessment must be made of the risk they pose to the procuring program and how the program can mitigate that risk. It is helpful to ascertain the current and past health status history of each source. At a minimum, source colony information should include an overview of their husbandry program, use of PPE, disease barriers (e.g., use of microisolator caging and BSCs), colony health surveillance program, policies for the introduction of new animals, and colony pathogen history. It is not uncommon to request that the shipping facility provide a year’s worth of relevant health surveillance testing data prior to approval of the animals for shipping. Regardless of the shipping colonies’ current health status or history, some very clean facilities may require that all mice and rats coming from noncommercial sources be rederived into the facility using embryo rederivation techniques. Alternatively, other programs have a strict quarantine and testing policy to screen animals being received from noncommercial sources. Animals obtained from the wild should be quarantined (CDC, African Rodent Importation Ban, http://www.cdc.gov/poxvirus/monkeypox/african-ban.html) and thoroughly tested for zoonotic diseases, as well as detrimental organisms known to infect closely related species (42 CFR §71.56; Karesh 2005; Hutson et al. 2015).
Animal Transportation and Stress (Shipping Containers, Temperature, Food and Water, Conveyance, and Personnel)
Specific parameters that must be met during transportation to ensure the well-being of a species have been set by many countries and the International Air Transport Association (IATA) (Live Animal Regulations, http://www.iata.org/publications/store/pages/live-animals-regulation.aspx). Using properly equipped, clean vehicles in conjunction with trained and experienced personnel throughout the transportation process is essential. Appropriately sized shipping containers, control of environmental factors such as ambient temperature, provision of food and water, in-transit safety precautions, and minimization of stressful events are all factors that must be considered when transporting animals in order to preserve their health and well-being (NRC 2006). Animals traveling long distances may experience fear, anxiety, and metabolic imbalances and require time to recover once they are received (Syversen et al. 2008). It is recommended that research institutions adopt policies to protect animals from being used immediately upon receipt, without an acclimation period. The lack of an acclimation period may confound the data or make many procedures (e.g., anesthesia) unsafe for the animal. For most species, a 48- to 72-hour minimum acclimation period following transportation is considered good practice (NRC 2006). However, recent evidence indicates that up to 2 weeks may be needed for some species to recover from the stress of transportation (Ochi et al. 2016).
Animal Receipt, Examination, and Acclimation
Newly arrived animals should be received by trained and experienced personnel in a designated area that is protected from extreme heat or cold, sufficiently illuminated, clean, uncluttered, dry, and quiet. Animals appearing sick or injured at the time of receipt, as well as animals arriving in damaged shipping containers, should be isolated for veterinary assessment prior to entering the colony. Everyone involved in the animal receipt process should be notified in a timely fashion and be able to devote adequate time and effort to the task of safe and expedient placement of the new animals in their new enclosures. Ideally, cage cards and/or animal records should be prepared ahead of time. The enclosures should be supplied with water promptly, and when possible, it should be established that new animals are drinking before they are placed on an automatic watering system. Feeding animals immediately after receipt is not as critical and should be carefully considered. Food consumption should be controlled and monitored to avoid GI problems, such as emesis and diarrhea, or other health issues. Animals should be observed for a period of time and allowed to adjust to their new environment, especially if they appear anxious or aggressive, or are vocalizing.
Some species (e.g., cats, rabbits, and marmosets) and individual animals are sensitive to abrupt changes in their diet and may refuse to eat upon receipt, or display signs of GI problems, such as diarrhea or bloat. These situations may require knowledge of the originating facility’s diet prior to the arrival of the animals and a careful program of transition from one diet to another.
Monitoring the Colony Health Status
Once the health status of a colony has been established, routine monitoring is essential to preserve that status. A comprehensive monitoring plan may include a variety of measures, such as daily health checks, routine physical exams, testing, health surveillance programs, and environmental monitoring. Postoperative observations and assessment of unanticipated or abnormal research-related effects and outcomes can also provide important indicators that a colony’s health status has been compromised. Ideally, an initial response plan to deviations from the chosen health status should be in place before a problem occurs.
Daily Health Checks
Daily health checks are a critical way to quickly detect changes in the colony health status. These observations are mandated in the U.S. AWR (9 CFR §2.40 b.3) and the Guide (NRC 2011), as well as some research funding entities in the United States and Europe. The number and timing of health checks will depend on the nature of the facility, species, and research being conducted. For example, infectious disease research animals may require multiple checks each day at specific intervals, whereas breeding colony animals may require less frequent observations with more flexible timing. In some situations, it may be advantageous to observe animals when they are most active, immediately prior to, during, and after they are fed.
Daily health checks are typically performed by individuals designated by the attending veterinarian who are trained and qualified to recognize the normal behavior and appearance of the animals under their care and identify abnormalities. Because the animal care specialists observe these animals each day, they are uniquely suited to quickly recognize any subtle changes in an animal’s posture, activity level, behavior, physical appearance, respiratory patterns, bodily functions, and use of its environment, including enrichment items (i.e., untouched favorite treats or decreased nest quality), regardless of the species under observation (i.e., birds, fish, amphibians, small mammals, or NHPs). Abnormalities should be documented and promptly reported to a veterinarian. It is also beneficial to provide staff with scoring sheets, potential symptoms, phenotypic differences, and pictures of common ailments seen in the research setting.
Sufficient time must be allotted for the care staff to conduct a thorough visual health check of each animal. This should be done a minimum of once a day, and preferably twice daily. For example, cages or fish tanks containing complex enrichment strategies or testing apparatus can complicate the visual assessment of the animals. In these situations, programs may require each cage to be removed from the rack daily to ensure visual assessment of all animals, or in the case of aquatics, the tank to be visible from various angles. Another key time to assess animals is during cage or tank changing. While changing cages or tanks, care staff should be attentive to both physical and behavioral signs exhibited by animals, regardless of the species, that may indicate the need for further assessment by a veterinary technician or technologist or veterinarian. Anomalous findings include, but are not limited to, changes in body conformation or physical appearance, guarding of a body part, abnormal respiration, and abnormal behavior (e.g., aggression or hiding).
When animals are housed in stacked cages, it may be helpful to equip care staff with flashlights to facilitate visualization of the animals housed in the lower cages. The use of binoculars can help visualize and assess group-housed animals in both outdoor and indoor facilities.
Postprocedural Observations and Assessment of Protocol-Related Effects
Postoperative observations are not only a way to determine if analgesia and surgical techniques are adequate; they also provide a means of assessing colony health. Changes in health status can be manifested through postoperative or postprocedural complications. For example, opportunistic pathogens may only become evident after an animal experiences a stressor such as surgery (Carty 2008). Close monitoring for abnormalities in respiratory, enteric, integumental, and other systems after surgery can alert staff to changes in the colony health status. In a similar fashion, protocol-related procedures and results may also point to problems in colony health (Gulani et al. 2016). However, one must distinguish whether these protocol-related effects are simply due to study procedures or genotypes, or are due to an unwanted pathogen in the colony. For example, weight loss could be a sequela of the experiment or a new pathogen in the colony. Abnormal performance in rodent behavioral tests could indicate a response to research interventions or new subclinical disease. Adverse anesthetic responses may be related to the protocol but could also be due to subclinical respiratory disease. When complications are observed after a surgical or other experimental procedure, the veterinarian and scientist should work together to determine whether these complications are due to changes in colony health status or to the research procedures involved.
Routine Physical Exams
Findings from recurring complete physical examinations and testing can indicate changes in colony health. Regular exams involve monitoring for abnormalities that may not be evident at visual cage- or kennel-side observations. Examples of these abnormalities may include decreased body condition, masses, enlarged lymph nodes, and periodontal disease, which could indicate the presence of tuberculosis, Pasteurella abscesses, shigellosis, or even McHV, in a NHP colony. Routine physicals for larger species should be performed no less than annually to determine viral, bacterial, parasitic, hematological, dental, and overall health status. The nature of the facility, the required health status, and local or national regulations will ultimately determine what tests are performed and how frequently. For smaller species (e.g., mice, rats, and fish), individual routine physical examinations are not regularly performed unless dictated by the research being conducted.
Testing and Health Surveillance Programs
The Federation of European Laboratory Animal Science Association (FELASA) has issued useful guidelines (Weber et al. 1999; Rehbinder et al. 1998, 2000; Voipio et al. 2008; Mähler et al. 2014) for monitoring the health of both large and small animals used in research. For large animals, the recommendations generally involve direct testing of research animals. Because the size of rodent colonies and modern husbandry techniques typically preclude individual examination and testing, colony surveillance testing is an efficient way to screen for many pathogens. If sentinel animals are used and tested for seroconversion, care must be taken to ensure that the animals are immunocompetent, relatively young, and capable of mounting immune responses to pathogens (Besselsen et al. 2000; Mähler et al. 2014). Sentinel animals are housed within the animal colony and, in most cases, receive dirty bedding from other colony cohorts. In cases where pathogens are not reliably transferred by dirty bedding (Artwohl et al. 1994; Cundiff et al. 1995; Compton et al. 2004; Perdue et al. 2008; Henderson et al. 2013), sentinels may be cohabitated with research animals. Sentinel animals are typically exposed to the colony cohorts for 6–8 weeks to allow for exposure, infection, and seroconversion and then tested by a variety of methods. FELASA recommends sentinel testing a minimum of every 3 months. Testing can include serology; polymerase chain reaction (PCR) testing of fecal material, hair and anal swabs, or intestinal contents; microscopic fecal and tape test examinations; bacterial cultures of a multitude of samples; histology; and gross necropsy. As a supplement to sentinel testing, research animals (typically those no longer needed for study) can be tested by the same methods to detect those organisms that may not be transferred via dirty bedding or that the sentinels may have cleared postexposure (Clarke and Perdue 2004). Alternatively, with investigator permission, survival testing that uses blood, fecal, and swab samples can be performed. Biological samples such as bone marrow and cells may also be tested before use in animals to detect unwanted pathogens in the originating colony.
Environmental Monitoring
Testing the animals’ macro- and microenvironments is an alternative or supplementary way to monitor colony health. Testing methods should be sensitive, specific, and able to detect a wide range of pathogens. Microbiological testing can be accomplished by RODAC™ (Replicate Organism Detection and Counting) or ATP-based systems (Ednie et al. 1998). RODAC is inexpensive, uses quick sampling methods, and offers quantifiable data. However, it only identifies viable aerobic bacteria and fungi that can be cultured and will not identify parasites or viral pathogens. This testing also requires several days of incubation before results are available. ATP-based systems use bioluminescence to indicate levels of ATP in live or dead organic material. These systems are especially useful in detecting and quantifying organic material on many substrates (walls, floors, and doors) and assessing the efficacy of sanitation practices (Turner et al. 2010). Unfortunately, these systems do not specify what organic material and pathogens are present and whether they are alive or dead. In addition, some disinfectants may alter the results (Turner et al. 2010). These systems also require that facilities establish their own standards for what values are acceptable and what values require action. Setting action cutoff values can be challenging and may require correlating ATP testing results with RODAC results.
Use of PCR testing to examine housing units, racking systems, supply and exhaust vents, behavioral equipment, and other items in the environment is becoming more popular as a way to monitor colony health, and incorporating its use into a surveillance program can reduce or eliminate the number of sentinel animals used (Henderson and Clifford 2013; Jensen et al. 2013; Compton and Macy 2015; Manuel et al. 2016). Food, water, and bedding can be sampled for pathogens as well (Rice et al. 2013). PCR testing is sensitive, relatively inexpensive when compared with purchasing, housing, and caring for sentinel animals; and available for a large number of organisms. Its exclusive use for health monitoring eliminates introducing sentinel animals into closed colonies. In some facilities, PCR testing has replaced rodent sentinel testing or is used in conjunction with a reduced sentinel testing program. However, care must be taken to ensure that PCR testing plans are appropriate for the type of caging system and filters used in the program. Some caging systems have been demonstrated to isolate the organisms to the animal’s cage, preventing contamination of and pathogen detection in the rack plenums and ducts (Henderson and Clifford 2013; Henderson et al. 2013). Therefore, a PCR testing program must also take into account how the pathogen spreads and persists in the environment.
Potential problems with PCR testing are (1) the interpretation of negative results and (2) the potential for false-positive results. Failure to collect the sample from the contaminated area of the equipment can lead to false-negative results, whereas false positives can be the result of nucleic acids remaining on previously contaminated equipment or in the environment (Leblanc et al. 2014). Ensuring that contaminated equipment and supplies are free of all traces of lingering nucleic acids can also be problematic.
Monitoring temperature, relative humidity, and air pressures for each housing room is important and required by some regulations (NRC 2011). This monitoring is critical to health status, as temperature and humidity can affect the growth of pathogens, and differential air pressures can affect the integrity of the separation of barrier rooms, surgical suites, rooms in which biohazardous agents are being studied, or known dirty areas from other colony rooms. Monitoring can be accomplished at the room level or by building automated environmental control systems that alarm when measurements fall outside set parameters. In some cases, both types of monitoring are performed to allow for redundancy in the event of a system malfunction. The NIH Office of Laboratory Animal Welfare (OLAW) strongly encourages institutions to use electronic technology for environmental monitoring, and the Guide (NRC 2011) advises the use of systems with automated alarms.
Response Plan to Health Status Deviations
Appropriate contingency plans should be prepared in advance and be ready to implement when unwanted pathogens are detected in a colony. Advance preparations can include cryopreservation of important rodent lines, maintaining separation of animal rooms through strict room entry order, adjustment of air movement from clean to less clean areas, the use of PPE, and keeping complete records of animal movements. Plans for health maintenance and monitoring, as previously discussed in this chapter, will also allow for early detection of a pathogen and reduce the likelihood of an outbreak spreading. The contingency plan should also include the steps to be taken when the containment barrier is breached, for example, when a rodent is found outside of its cage or is accidentally dropped on the floor.
When a positive result is obtained, one must consider (1) the nature of the pathogen, (2) how the pathogen is shed, (3) how the pathogen is spread, (4) the sensitivity and specificity of the pathogen tests, (5) prevalence of the agent, (6) how seroconversion may complicate test results, (7) how persistent the pathogen is in the environment, and (8) how treatment may affect research results. Responses should be aimed at limiting pathogen spread and eradicating the pathogen from the colony (Reuter et al. 2011). The first, and most important, step is to isolate the potentially affected animals and conduct tests to confirm the previous results. Next, notify the appropriate institutional personnel and report the disease and quarantine as required by local and national regulations. Other important management practices to consider include changing room entry and access procedures, increasing the level of sanitation and PPE, implementing special handling requirements for animals and enclosures, closing the colony to imports and exports, and ceasing breeding in some cases. If the pathogen is zoonotic, staff will need to be educated on the risks to their health. It is critical to post signage on room or enclosure doors that very clearly describes all of this information and the new required management practices.
Responses to outbreaks are dependent on the pathogen and may involve treatment, testing and culling, caesarean section, embryo transfer, and cessation of breeding. When treatment is utilized, one may need to give consideration to the current research being conducted in the affected area. Consideration should also be given as to whether research animals will naturally clear the infection or serve as reservoirs. Using diagrams and schematics to track testing and test results can be extremely valuable in managing outbreaks.
The time, cost, and energy required to stop an outbreak will vary based on the pathogen and type of animals affected. Testing and other necessary responses, such as euthanasia of animals, can tremendously impact the responding staff, both physically and emotionally. Boosting morale and being sensitive to personnel fatigue are critical to successfully ending an outbreak. In some cases, research staff may be resistant to new management practices during an outbreak, but it is important to inform them of the enormous time, cost, and loss of research data and animals that can occur if outbreaks are not addressed in an appropriate and timely manner. If there is the potential that animals from multiple research groups may be impacted, it is useful to meet with all groups involved to explain the situation and plan for resolution.
Clinical Care of Animals
Diagnosis and Treatment of Ill Animals
When physical or behavioral abnormalities are observed in research animals, it is important to determine whether the animals are experiencing a natural or experimentally induced illness and whether intervention with treatment, monitoring, or euthanasia is appropriate. Diagnostic and treatment approaches and humane and experimental endpoints must be considered when addressing the observed illness. All these considerations must be done in the context of balancing animal welfare with research objectives.
Natural or Experimentally Induced Illness?
When routine daily observations detect physical or behavioral abnormalities in a research animal or group of animals, prompt reporting to the veterinarian is critical. Unexpected findings should also be reported to the scientist, who can provide specifics about the nature and timeline of the research. The veterinarian and scientist should work together to determine whether the illness observed is caused by a natural process or the research. Ideally, issues resulting from the research (e.g., test compounds, surgical procedures, deleterious phenotypes, and infectious agents) should have been described by the scientist in the protocol and understood by the veterinarian and animal care staff prior to study initiation.
If the abnormality is related to an unanticipated phenotype, the investigator, in consultation with the veterinarian, should determine the importance of the animal and if further characterization is justified. This may include routine diagnostics, necropsy and histology, initiating a genotyping or phenotyping study (Crawley 2000), or seeking assistance from other researchers who have experience with the model. If further characterization is justified, the IACUC should be apprised of the situation and the actions being taken to further define the potential new model.
Veterinary Diagnostic Procedures
Veterinary diagnostic procedures include physical examination, serum biochemistry panels, complete blood counts, serology, bacterial cultures, urinalysis, cytology, fecal exams, PCR testing of various samples, radiography, ultrasound, MRI and CT scans, biopsies, endoscopy, exploratory surgery, histology, and necropsy. These tests are available for rodents, just as they are for larger species, so one should not overlook their value in assessing small animal cases. Many of these tests are available through commercial services, but they can be readily performed within the facility with the proper equipment, personnel training, budgets, and space. To the greatest extent possible, the diagnostic test should be chosen with consideration of the research goals. For example, sedation for radiographs would be contraindicated just before an animal is scheduled to undergo behavioral testing at a critical research time point. The veterinarian and scientist should work together to devise a diagnostic plan that meets the needs of the animal and the study.
Treatment and Monitoring Approaches
When a diagnosis is made, a plan for treatment, monitoring, or euthanasia will be established by the veterinarian and should be made in consultation with the researcher. The decision to treat, observe only, or euthanize is complex and is discussed in the next section “To Treat, Monitor, or Euthanize?” (page 751). If treatment is chosen, there are several approaches to consider. Medications can be provided in the drinking water, regular feed, or treats, or through injections, continuous intravenous delivery systems, topical applications, or transdermal patches. Supportive care can be in the form of supplemental heat, fluids, or oxygen; additional bedding or nesting material; high-calorie and high-value foods or treats; and bandaging lesions. The treatment approach should consider whether an individual or multiple animals need to be treated, what resources are available for treatment, the nature of the sick animals, and the research objectives. For example, if mice in several cages must be treated, labor costs for providing medicated water or feed to multiple cages will be less than injecting each individual mouse needing treatment. This is an important factor to consider when staffing is limited. Providing medications through feed or water will also result in less stress to the animal than injections. If the nature of an animal is such that it refuses to take medications through water, food, or treats, treatment may be limited to oral gavage or injectable routes. Research objectives may affect the treatment plan as well. For example, if only two of five mice in a cage require treatment, and treatment may interfere with the study, the two sick mice can be treated separately from healthy cage mates or removed from the study. Also, if the research requires a strict and/or measured diet, oral medications in feed and treats may pose problems to the study and necessitate an alternative dosing route.
If the needs of the animal and the research are such that only monitoring can be performed, scoring systems may be useful in determining when treatment or euthanasia will be initiated (Langford et al. 2010; Bekkevold et al. 2013; Nunamaker et al. 2013). Scoring systems provide an objective way to determine when endpoints have been met and/or interventions are necessary. A standard scoring system lists abnormal signs that can be observed and then asks the observer to state whether a specific abnormal finding is present or absent (Figure 31.1). Scoring systems may also involve rating abnormalities using a numerical scale or based on qualitative observations. For example, arthritis in mice can be scored using millimeters of paw swelling or using subjective numbers to indicate increasing degrees of lameness, swelling, and erythema. When a single number, tally of numbers, or a certain qualitative criterion reaches a predetermined score, interventions with treatment, removal from the study, or euthanasia will occur. Scoring sheets can also list humane and experimental endpoints, provide documentation of supportive care, and provide a means to evaluate complications and general health of the animals on the study. Scoring sheets should be created prior to study initiation and be specific to the study and the species. The scoring criteria should be adapted as the study progresses to incorporate unforeseen developments. Scoring sheets should be available and readily retrievable for all staff involved in the care of the animals being scored.
Figure 31.1
EAE score sheet. (Adapted from Division of Veterinary Resources, National Institutes of Health.)
Two sample scoring and monitoring sheets are provided in Figures 31.1 and 31.2. Figure 31.1 includes the experimental autoimmune encephalomyelitis (EAE) scoring system that rates the degree of the disorder on a graded scale from 0 (no abnormality) to 5 (quadriplegia or premoribund state). Figure 31.2 includes an induced arthritis scoring system that rates the degree of abnormality of each paw from 1 (no erythema or swelling) to 4 (erythema and severe swelling encompassing the foot and digits). The scores for each paw are then added for a total arthritis score. Both sheets prompt the observer to indicate several things: (1) whether a mouse is bright, alert, and responsive (BAR) or quiet, alert, and responsive (QAR); (2) the score for a given mouse according to the scoring system; (3) the body condition score (Burkholder et al. 2012); (4) the pain score (Burkholder et al. 2012); and (5) whether supportive care treatments, such as food supplements and longer sipper tubes, are present in the cage. The sheets also require observer initials, provide space for any important additional observations, and provide other important instructions (e.g., notify the veterinarian immediately if a mouse reaches an EAE score of 4).
Figure 31.2
Arthritis score sheet. (Adapted from Division of Veterinary Resources, National Institutes of Health.)
To Treat, Monitor, or Euthanize?
The decision to treat, monitor only, or euthanize is complex and necessitates good communication between veterinarians and scientists. A complete understanding by both parties of the research objectives, the species involved, the severity and prognosis of the illness, and required IACUC-established humane and experimental endpoints, along with sound professional veterinary and scientific judgment, will facilitate the decision. In addition, the scientist should describe in the protocol any limitations on the veterinary interventions that can be provided to the animals due to adverse effects on the research being conducted (e.g., use of steroids). Monitoring may be chosen when treatment would interfere with the study and the clinical condition is mild and has a favorable prognosis. If the condition is more significant and the prognosis less favorable, it may be best to treat or euthanize the animal before data is lost due to unexpected death. Euthanasia and final sample collection may be the optimal choice if treatment negates the ability to obtain scientifically useful data from the animal. Ideally, preestablished humane endpoints and intervention contraindications outlined in the study design are detailed enough to direct decisions about treatment, monitoring, or euthanasia. While the veterinarian and researcher should collaborate on decisions about whether to treat, monitor, or euthanize, the final decision in critical cases is that of the attending veterinarian.
Humane and Experimental Endpoints
Humane and experimental endpoints are vital to managing animal welfare while concurrently meeting research objectives (Demers et al. 2006). Experimental endpoints are those that relate to the timing of the study and are typically dictated solely by the science behind the research. An example of an experimental endpoint would be to terminally collect tissues from mice 4 weeks after administration of a test drug. Humane endpoints are those that determine when animals will be relieved of pain or distress by being taken off study, treated, or euthanized. These humane endpoints minimize pain and distress to the greatest extent possible without compromising the goals of the study. Ideally, humane endpoints are devised in such a way that the study ends before an animal experiences pain or distress, and also at a time when valuable research data can still be collected. Humane endpoints can be qualitative (unkempt coat, poor response to stimuli, hunched posture, or dyspnea) or quantitative (low body temperature, weight loss, decreased food or water consumption, or predetermined biomarker values) (Toth 1997). Because humane endpoints balance the needs of the animals as well as the research, they can be challenging to establish. Some useful references regarding endpoints include those created by the Organisation for Economic Co-operation and Development (2000), the Canadian Council on Animal Care (1998), and the National Research Council (2000). Establishing humane endpoints requires collaboration between veterinarians, scientists, and compliance bodies, such as the IACUC (Stokes 2000; NRC 2011, pp. 27). This must be done prior to initiating a study, although humane endpoints may need to be adjusted during the course of a study.
If the expected outcomes are unknown and humane endpoints cannot adequately be determined prior to study initiation, a pilot study should be performed in a small number of animals, ideally under the veterinarian’s supervision. Pilot studies can determine the earliest predictive signs of adverse study effects and/or impending death. Information collected from pilot studies will allow the development of endpoints that avoid causing more pain or distress than necessary to the animals, while meeting the research objectives. Pilot studies can also be a useful tool for training staff to accurately understand, anticipate, and detect the protocol-defined humane endpoints. Although animals should always be monitored for the presence of unanticipated adverse effects, this is especially true when conducting pilot studies. When observed, adverse effects should be reported to the veterinarian, investigator, and IACUC.
Once humane endpoints have been established, the lines of authority and communication regarding animal well-being must be clearly defined. It is critical to determine who will be responsible for observations, the timing and frequency of observations, and how to properly document and follow up on findings. Close monitoring of the animals will be necessary to ensure that the endpoints are correctly implemented and no further refinements are needed. Animals that are being monitored for humane endpoints need to be clearly identified. Personnel performing the monitoring must be trained to recognize pain and distress in the species and understand the research procedures and expected outcomes. Observations may need to be more frequent than usual (e.g., every 4–6 hours), rather than once each day, and should be timed to have the maximum value (when animals are most likely to reach a humane endpoint). As often as possible, studies should be scheduled so that this timing of critical observations occurs during normal work hours. Animals should be observed first at a distance with minimal disruption, followed by assessing their responsiveness to stimuli and direct handling. Data for an endpoint assessment can be obtained by telemetry, activity monitors, video recordings, or other methods that are not disruptive to the animals.
After humane endpoints have been determined and put into practice, there may be instances where it is difficult to determine whether an animal has reached a defined humane endpoint. In this case, the veterinarian and researcher should work collaboratively to come to a consensus. If an agreement cannot be reached, the attending veterinarian has the final decision as to the animal’s disposition (9 CFR §2.33 a.2).
Animal Welfare Considerations
It can be challenging to balance the welfare of research animals with study objectives. This is especially true when making the decision to treat, monitor, or euthanize and when defining and adhering to endpoints. If the research or animal care staff have any animal welfare concerns during a study, there should be multiple mechanisms in place to report their concerns without fear of reprisal. A safe reporting mechanism allows veterinarians, scientists, managers, and compliance bodies to discuss and address issues as they arise. Addressing issues early helps the laboratory animal medicine and scientific community to uphold the three Rs (replacement, reduction, and refinement) (Russell and Burch 1959) of animal research and meet regulatory requirements. Discussion and resolution of issues can result in refined endpoints, reduced pain and distress, use of fewer and/or healthier animals, and better-quality science.
Normal versus Abnormal Behavior
The only way to be able to recognize “abnormal” behavior is to devote time to learning normal behavior patterns to use as a point of reference (Arnold et al. 2011; Rock et al. 2014). Animal behavior is a set of functions that reflect the animal’s response to all environmental factors and challenges. These behaviors include food and shelter and mate seeking, as well as nest-making activities and performing complex tasks, such as operating equipment and using tools by NHPs. Every individual working with animals should spend time studying a species’ normal behavior in different situations”“resting, feeding, interacting with conspecifics and humans, posturing, and responding to easy and difficult challenges. A careful student of behavior will notice that most behaviors displayed in nonstressful situations follow certain patterns. Learning these patterns is very useful for interpreting animal health and well-being. Knowledge of behaviors displayed in response to stressful situations is also very useful for providing insights into an animal’s mental state, which can help with choosing the best animal for a needed task (e.g., an aggressive or anxious animal may not be the best candidate for a task that requires a confident and calm animal).
Pain versus Distress
Pain results from potential or actual tissue damage. Pain can be considered a potent source of stress, that is, a stressor in and of itself, that can lead to distress and maladaptive behaviors (NRC 2009a). Distress is an aversive state in which an animal is unable to adapt completely to stressors and the resulting stress, leading to maladaptive behaviors, such as abnormal feeding, absence or diminution of postprandial grooming, inappropriate social interaction with conspecifics or handlers, and inefficient reproduction (NRC 2009a).
Recognition and Assessment of Pain and Distress
The nervous system processes the sensory features of tissue-damaging stimuli, such as damage quality, intensity, location, and duration. What is perceived results in behavioral and physiologic responses that are under the influence of emotional, motivational, and cognitive processes (NRC 2009a). Stress and distress are complex syndromes that are difficult to define and even harder to interpret and recognize (Wright et al. 1985). If an animal is in pain, it might also be in distress, and vice versa.
Score Sheets and Training
Biomedical research puts much emphasis on using objective parameters to avoid misinterpretation of the data. A set of data collected by one researcher can be interpreted differently by another researcher who is not as familiar with the data, science, or subject animals. If the animal is performing behavioral tasks that are critical in the research project, it is paramount that the animal’s training be done in a comprehensive and logical fashion that is transparent for all research team members and leaves no room for speculations. The training should be designed in such a way that it incorporates the individual traits of the animal.
As discussed earlier in this chapter, score sheets and scoring scales can be useful tools when evaluating an animal’s well-being in a nonsubjective manner, as well as when training individuals to recognize abnormalities. Our ability to recognize an animal experiencing pain or distress and assess its severity is the first step toward prevention and treatment of the problem. Recent work has demonstrated that the facial expressions of mice, rats, and rabbits can provide a rapid and reliable means of assessing the presence and severity of pain (https://www.nc3rs.org.uk/grimacescales). “Grimace scales” that equate facial changes related to narrowing of the animal’s eyes and position of the whiskers and ears have been demonstrated to be reliable indicators of both the presence of pain and its intensity.
Species-Specific Signs of Pain or Distress
The identification of pain and distress in animals is a challenging task (NRC 2009a). Because verbal feedback is not an option, veterinarians and researchers must rely on their experience, understanding, and ability to detect species-specific signs of pain or distress. Critical to the assessment of pain or distress is the ability to distinguish between normal and abnormal animal behavior. This is especially true when dealing with a species that often exhibits pain and distress with only subtle behavioral changes. Animals must be monitored by trained individuals throughout a study for pain and distress as appropriate for the species, conditions, and procedures. Therefore, it is critical that the individuals assessing an animal be trained in the species’ specific signs of pain and distress, as well as the potential outcomes of the research manipulations. As discussed earlier in this chapter, pain and distress scoring is one method to convert subjective animal observations into an objective system, which some have found to be helpful in assessing animal behavior.
Rodents
Ailing rats and mice most commonly show decreased motor and nest-building activity, piloerection, and an ungroomed appearance, but may exhibit other signs as outlined in Table 31.1 (Kohn et al. 2007). Although decreased food and water consumption is commonly associated with pain or distress in other species, these parameters are challenging to monitor in rodents. Also, changes in body weight are not sensitive enough to be used as an indicator of acute pain or distress (NRC 2009a).
Table 31.1
Potential Signs Associated with Pain or Distress in Rats, Mice, and Rabbits.
Cavies
Guinea pigs in pain usually remain quiet (as opposed to stampeding and squealing when frightened but pain-free) (Kohn et al. 2007; Shomer et al. 2015). They also display behaviors similar to those of other rodents, such as rats and mice.
Rabbits
Rabbits in pain can appear apprehensive, anxious, dull, or inactive and assume a hunched appearance, attempt to hide, and squeal or cry. When experiencing acute pain, they may show aggressive behavior with increased activity, excessive scratching and licking, and exaggerated reactions to handling. With abdominal or muscular pain, they sometimes grind their teeth and salivate excessively, in addition to displaying increased respiratory rate and inappetence. Rabbits in distress might cannibalize their young and tend to be more susceptible to the “tonic immobility reflex” (aka immobility reflex, animal hypnosis, tonic immobility, playing possum, mesmerism, and dead faint). This reflexive behavior is considered to be a mechanism of defense against predators, as it renders the animal less sensitive to pain. It abolishes voluntary motor activity. Spinal reflexes are suppressed, but not abolished. In rabbits, the most susceptible species, fine muscle tremors can occur initially or be induced by stimulation of the patellar tendon reflex. Fully immobilized rabbits exhibit pronounced catalepsy with reduced muscle tone. Additional signs of possible pain or distress are outlined in Table 31.1 (Kohn et al. 2007).
Cats
A general lack of well-being is an important indication of pain in cats (Robertson 2005; Epstein et al. 2015). They may be quiet and show an apprehensive facial expression, with the forehead appearing creased. They might not have an appetite and may cry, yowl, growl, or hiss if approached or made to move. The cat’s posture becomes stiff and abnormal, and it tends to hide or separate itself from other cats. If the pain is located in the head or ears, the cat might tilt its head toward the affected side. When the pain is only thoracic, the head, neck, and body might be extended, while a cat with generalized pain in both the thorax and abdomen might be crouched or hunched. A cat with abdominal or back pain might stand or lie on its side with its back arched or walk with a stilted gait. Pain in one limb is usually manifested by limping or holding up of the affected limb, with no attempt to use it. Incessant licking is sometimes associated with localized pain or psychological distress. Cats in severe or chronic pain look ungroomed and behave markedly different from normal (NRC 2009a).
Dogs
Dogs in pain generally appear less alert and quieter than normal and may demonstrate stiff body movements and an unwillingness to move. A dog in severe pain might lie completely still while watchful, or adopt an abnormal posture to minimize its discomfort. With less severe pain, dogs can appear restless and more alert, but have a loss of appetite, shivering, and increased respiration with panting. Spontaneous barking is unlikely. Instead, they are more likely to whimper or howl, especially if unattended, and might growl without apparent provocation. When handled, dogs can bite, scratch, or guard painful regions and be abnormally apprehensive or aggressive (NRC 2009a; Epstein et al. 2015).
Ferrets
Behavioral changes are often the best indication of a ferret’s well-being. These changes can include the display of unusual aggression toward other animals or their handlers, as well as hiding. Ferrets in pain often become lethargic and stop bodily grooming, resulting in a disheveled look. Ferrets rarely vocalize when they are in pain or distress, and specific signs are largely dependent on the individual animal. Animals in pain or distress can display a loss of appetite, stop drinking, and experience weight loss (Hillyer and Queensberry 1997). Additional potential signs of pain or distress may include (1) drooped or laid-back ears; (2) closed or squinty eyes; (3) clicking, grating, or grinding of the teeth; (4) drooling, salivating, or slobbering; (5) hesitant, immobile, or slow movements; (6) guarded, noncurled, or tense movements; (7) trembling; (8) crying or whimpering; and (9) guarding, avoiding, or withdrawing from touch.
Nonhuman Primates (New and Old World)
NHPs show remarkably little reaction to surgical procedures or injury (NRC 1998). Some animals can hide pain and may look well until they are gravely ill or in severe pain. A NHP that appears sick should get immediate assessment and possible intervention. A NHP in pain has a general appearance of misery and dejection. It might huddle in a crouched posture with its arms across its chest and its head forward with a grimace and glassy eyes. Acute abdominal pain can be shown by facial contortions, clenching of teeth, restlessness, and shaking accompanied by grunts and moans, with food and water usually refused. A monkey in pain can also attract altered attention from its cage mates, ranging from a lack of social grooming to attack (NRC 2009a).
Sheep and Goats
Sheep and goats may appear dull and depressed, hold their heads low, and show little interest in their surroundings or eating. On handling, they might react violently or adopt a rigid posture designed to immobilize the painful region, combined with grunting and grinding of teeth. Localized pain may result in persistent licking and kicking at the offending area and, when the pain is severe, vocalizing. Sheep can tolerate severe injury without overt signs of pain or distress, although abnormal changes in posture and a reluctance to move are often apparent. Goats are more likely to vocalize in response to pain. After castration or tail docking, lambs may show signs of pain by standing and lying repeatedly, wagging their tails, occasionally bleating, and displaying neck extension, dorsal lip curling, kicking, rolling, and hyperventilation (NRC 2009a; Underwood et al. 2015).
Swine
Swine normally squeal and attempt to escape when handled, and pain can accentuate these reactions. When in pain, swine often show changes in social behavior, gait, and posture and, when bedding material is present, will refrain from making a bed. Adults typically become more aggressive. Squealing is characteristic when painful areas are palpated. When in severe pain, swine are often unwilling to move and might hide in bedding (NRC 2009a; Underwood et al. 2014). As swine generally have good appetites, any individuals who go off feed should be assessed for potential signs of pain or illness.
Fish
Fish respond to pain differently than mammals. They exhibit a pronounced initial response to injuries or to contact with irritants, but their response to chronic stimuli might be small or absent. Fish with severe wounds, which would cause immobility in a mammal, often appear to behave normally and even resume feeding. Fish react to noxious stimuli, such as puncture with a hypodermic needle, with strong muscular movements. When exposed to a noxious environment, such as an acidic solution, they show abnormal swimming behavior and attempt to jump out of the water, their coloring becomes darker, and their opercular movements become more rapid. Such effects indicate some, perhaps considerable, distress, but it is not possible to describe the distress unequivocally as pain induced (NRC 2009a; Sneddon 2009; Smith 2014).
Birds
Birds in pain can show escape reactions, vocalization, and excessive movement (Machin 2005). Small species struggle less and emit fewer distress calls than large species. Head movements increase in extent and frequency, and there can be an increase in heart and respiratory rates. Prolonged pain results in loss of appetite, inactivity, and a drooping, miserable appearance, with the eyes held partially closed, the wings held flat against the body, and the neck retracted. When a sick bird is handled, its escape reaction is often replaced by tonic immobility (see the “Rabbits” section above). Birds with limb pain avoid use of the affected limb and “guard” it from extension (NRC 2009a). Like swine, birds generally have good appetites, and any changes from the expected consumption of food should be investigated immediately.
Amphibians
Acute pain in amphibians can be characterized by flinching and muscle contractions. There might be aversive movements away from the unpleasant stimulus and attempts to bite. More chronic and persistent pain might be associated with anorexia, lethargy, and weight loss, although a direct and specific association of any of these signs with pain may be sometimes challenging (NRC 2009a).
Category E Studies: Unrelieved Pain or Distress
The Guide states that the IACUC should consider the use of appropriate sedation, analgesia, and anesthesia for animals (NRC 2011) expected to experience pain or distress during research manipulations. The AWR (paragraph §2.36 b.7) specifically require painful procedures, for which such drugs cannot be used, to be placed in Column E of the Animal and Plant Health Inspection Service/Animal Care (APHIS/AC) annual report (AWR). Placement of study animals in the Column E category must be scientifically justified and can only be approved after careful consideration by the IACUC. Committee review must include a careful examination of the requested animal numbers with emphasis on an investigator-provided extensive search for alternatives. A smaller pilot study to harvest preliminary data may be stipulated and include oversight by a veterinarian.
Prevention and Treatment of Pain
Prevention and relief of pain and distress in animals is humane and essential in biomedical research. The IACUC has the responsibility for ensuring that all animals under their oversight are used humanely and in accordance with a number of federal regulations and policies (NRC 2011; AWR 9 CFR §2.31 d.1.i). The principal investigator (PI) and the IACUC must fully understand their legal requirements to establish both appropriate endpoints and expectable methodologies for relieving pain and distress. The individuals responsible for monitoring the animals for pain and distress must be identified and trained.
The obligation to reduce pain and distress starts with the submission, review, and approval of an animal study proposal, but does not end there. Animals must be continually monitored for pain, distress, illness, morbidity, and/or mortality during the course of a research study. If unexpected pain or distress is observed, and is more than an isolated incident, the PI must submit an amendment delineating the unexpected problem and stating his or her proposed resolution to the issue. Alternatively, as previously stated, the PI could justify the need for unrelieved pain or distress in the amendment or, in the case of regulated species, submit a Category E justification (USDA Animal Care Policy 11, https://www.aphis.usda.gov/animal_welfare/downloads/Animal%20Care%20Policy%20Manual.pdf).
Whenever more than transient pain or distress is anticipated, measures should be taken to minimize or prevent the development of pain and/or distress (Mench and Blatchford 2014). Following any intervention strategy, the animals must be closely monitored to ensure the effectiveness of the measures taken and determine if or when additional treatment will be necessary. The extent and frequency of monitoring will depend on the level of postsurgical/procedural pain and/or distress anticipated and the chosen intervention strategy.
Intervention strategies for the management of pain and distress may include the use of both pharmacological and nonpharmacological approaches. Nonpharmacological strategies may include, but are not limited to, (1) modified housing and husbandry practices, (2) dietary modifications and supplements, (3) surgical approaches, (4) desensitization and acclimation strategies, (5) acupuncture, and (6) euthanasia under the right circumstances (e.g., https://www.primatevets.org/Content/files/Public/education/NHP_Endpoint_Guidelines.pdf). The chosen strategy will vary with the species, the procedures being performed, the duration of action needed, the route of administration preferred, the degree and type of analgesia required, and the research being conducted (Table 31.2) (ACLAM 2006; http://oacu.od.nih.gov/ARAC/documents/Pain_and_Distress.pdf).
Table 31.2
Postprocedural Pain Potential.
When administering pharmaceuticals, it is important to understand the pharmacokinetics and duration of action of the drugs used to alleviate pain or distress. The pharmacokinetics of an agent must be taken into consideration when designing a strategy to monitor the effectiveness of the drug intervention. The strategy should include a plan to carefully monitor the animal during the period when the effectiveness of the agent is starting to wane, to determine if additional treatment is required. Resources (Fish et al. 2008; NRC 2009a) and formularies (Hawk et al. 2005; Carpenter 2012) are available today that provide extensive information on the recognition and alleviation of pain and distress in laboratory animals.
The documentation of monitoring of the animal for pain and distress is important. The identification of cages containing animals where a potentially painful or distressful procedure has been performed can prove helpful in drawing special attention to the animal during the caretaker’s daily health check. A “special observation” cage card works well for this. Cages containing animals requiring more intensive monitoring should also be appropriately identified and their monitoring and/or treatments documented at either the room, cage, or animal level (e.g., room log, cage card, or medical record). This is in addition to the investigator’s notations in his or her laboratory notebook. It is important that the documentation be available to all personnel monitoring the cage or animal (e.g., IACUC, veterinarians, and animal care staff).
Anesthetics, Analgesics, Tranquilizers, and Sedatives
Anesthesia is a reversible process for the purpose of producing a convenient, safe, and inexpensive means of restraint so that clinical or experimental procedures may be conducted with a minimum of pain, discomfort, distress, or toxic side effects to the patient or subject. The goal of anesthesia is to preserve normal physiologic function without confounding the experimental protocol under consideration. The Animal Welfare Act (9 CFR 2143 a.3) requires that anesthetic, analgesic, and tranquilizing drugs be utilized in accordance with currently accepted veterinary practice and produce in the individual subject a level of anesthesia, analgesia, or tranquilization appropriate for the design of the experiment. The use of anesthetics, analgesics, and tranquilizers in laboratory animals is necessary for humane and scientific reasons. The choice and use of the most appropriate drugs are matters for the attending veterinarian and professional veterinary judgment. The veterinarian must provide research personnel with guidelines, training, and advice concerning the choice and use of these drugs. If a procedure is likely to cause more than momentary pain or discomfort to the subject animal, but the use of anesthetics, analgesics, or tranquilizing agents would defeat the purpose of the procedure, a written justification of a Category E listing for unrelieved pain or distress must be submitted and approved by the IACUC.
The following are common terms related to the use of anesthetics, sedation, and tranquilization:
- Anesthesia: A state of controllable, reversible insensibility in which sensory perceptions and motor responses are both markedly depressed. A total loss of sensation in the whole body or one of its parts occurs.
- General anesthesia: Loss of consciousness in addition to loss of sensation. Ideally, general anesthesia includes hypnosis or narcosis, hyporeflexia, and analgesia.
- Balanced anesthesia: Surgical anesthesia produced by a combination of two or more drugs or anesthetic techniques each contributing its own pharmacological effects. Balanced anesthesia is characterized by unconsciousness, analgesia, and muscular relaxation.
- Dissociative anesthesia: A central nervous system (CNS) state characterized by catalepsy, profound peripheral analgesia, and altered consciousness produced by the cyclohexamine drugs (e.g., ketamine).
- Tranquilization (neurolepsis or ataraxia): A state of tranquilization and calmness in which the subject is relaxed, awake, and unconcerned with its surroundings. Tranquilizers act by suppression of the hypothalamus and reticular activating system.
- Sedation: A mild degree of CNS depression in which the subject is awake but calm. The subject can be aroused. Action is by a dose-dependent depression of the cerebral cortex.
- Hypnosis: Artificially induced sleep or a trance resembling sleep from which the subject can be aroused by stimuli.
- Narcosis: Drug-induced sedation in which the subject is oblivious to pain, with or without hypnosis.
- Analgesia: Loss of sensitivity to pain.
- Neuroleptanalgesia: Hypnosis and analgesia produced by a combination of a neuroleptic drug and an analgesic drug.
- Catalepsy: A state in which there is a malleable rigidity of limbs, which, if the limbs are placed in various positions, is maintained for a time. The subject is generally unresponsive to audio, visual, or minor pain stimuli.
It is beyond the scope of this chapter to discuss the wide range of agents available to the veterinarian for anesthesia. Many excellent references are available on the selection and use of anesthetics, sedatives, and tranquilizers in laboratory animal medicine and general veterinary practice (Fish et al. 2008; Gaynor and Muir 2014; Grimm et al. 2015).
The selection of the most appropriate agents and techniques are dependent on
- Duration of and type of operation or procedure
- Species, breed, age, and relative size of the subject
- Physical status of the subject and concurrent medication or treatments
- Disposition or demeanor of the animal
- Presence of concurrent pain or distress
- Type of facilities and help available, including level of experience and training
- Personal knowledge, experience, and familiarity with available agents and equipment
In general, all anesthetics cause a dose-dependent depression of consciousness, as well as a depression of normal physiological homeostasis (e.g., heart rate, respiration, blood pressure, and thermoregulatory centers) (Brunson 2008; Meyer and Fish 2008). In balanced anesthesia, various drugs are administered together that synergize each other, yielding the desired outcome at lower doses than would be possible with the administration of the single drugs alone. At lower doses, most drugs are safer and demonstrate fewer adverse effects.
All general anesthetics induce predictable stages of dose-dependent anesthesia: (1) voluntary movement or excitement, (2) involuntary excitement, (3) surgical anesthesia (Planes 1–4), and (4) medullary paralysis or death. The goal of anesthetic induction is to take the patient through the first two stages as quickly as possible. This is often facilitated by the intravenous administration of the drugs. When intravenous administration is not possible, the drugs are commonly administered intramuscularly (IM) in larger species and either subcutaneously (SQ) or intraperitoneally (IP) in smaller species. Non-intramuscular injection commonly leads to a slower absorption of the drug into the circulatory system and to prolonged voluntary and involuntary excitement phases. Care must be taken to anticipate the response of the animal and prevent injury to the animal and to protect personnel from being scratched, kicked, or bitten.
Sedatives, anxiolytics, and neuromuscular blocking agents do not provide analgesia, and anesthetics that induce unconsciousness at lower doses may not produce a pain-free state. It must be understood that unconsciousness does not always equate with a state of analgesia. In addition, it is critical to understand that neuromuscular blocking agents that produce a state of total paralysis, including respiratory paralysis, are not anesthetics and do not induce unconsciousness. Therefore, neuromuscular blocking agents must never be used in an animal that is not deeply anesthetized. When monitoring an animal that is both anesthetized and paralyzed, it is critical to, at a minimum, monitor both the animal’s heart rate and blood pressure to ensure an adequate depth of anesthesia. A rise in heart rate with a stable or increasing blood pressure can indicate the animal’s perception of pain or distress and inadequate anesthesia. However, a rise in heart rate coupled with decreasing blood pressure can mean the animal is too deeply anesthetized. It has also been recommended when using neuromuscular blocking agents that the plane of anesthesia be periodically lightened until a slight increase in heart rate is observed and then returned to the deeper plane to ensure that adequate anesthesia is present.
While careful monitoring of an animal’s reflexes provides a good indication that a plane of surgical anesthesia has been reached, concurrent monitoring of the animal’s heart rate and blood pressure is required to determine if the animal is pain-free. As stated above, if an animal’s heart rate is increasing and its blood pressure is normal or increasing, the animal is displaying a physiological response to pain. Costly equipment is not required for monitoring; manually taking a peripheral pulse and a mucous membrane capillary refill time will suffice in most situations.
In most species, reaching a surgical plane of anesthesia is accompanied by loss of a toe pinch withdrawal response; surgical anesthesia may also be accompanied by disappearance of head shaking in response to pinching of an ear (Smith and Dannerman 2008; Gaynor and Muir 2014; Grimm et al. 2015). The pattern of respiration, slow regular deep breaths, has been demonstrated to be a good indicator of the level of surgical anesthesia.
Care must be taken throughout the anesthesia and recovery periods (Hampshire and Davis 2008) to provide the animal with appropriate monitoring and nursing support. This should include the provision of a quiet environment, a warm ambient temperature, fluid support, and additional thermal support as required. Physiological monitoring is critical, but the nature of the monitoring requirement can vary by species, length and type of procedure, available equipment, and funding. The availability of precision vaporizers, pulse oximeters, and respiratory, cardiac, and blood pressure monitors increases the safety of anesthetic use.
Whenever possible, care should be taken to prevent the development of pain, rather than focusing on the treatment of established pain. Tissue injury and other noxious stimuli cause the release of chemicals that activate sensory neurons (i.e., nociceptors). Once activated, the nociceptors send electrical impulses to the brain that are interpreted as pain. Left unchecked, the chemicals set off a cascading inflammatory reaction within the tissues around the nociceptors that become abnormally sensitive. Use of preemptive analgesia, the administration of preoperative and intraoperative medication to block pain pathways and induce analgesia (Beilin et al. 2003), can diminish or prevent the development of pain.
In many situations, pain is easier to prevent than to treat. Preemptive approaches focus on inhibiting changes in the peripheral and central nervous system that contribute to heightened postprocedural pain. By taking preemptive measures to prevent the sensitization or “windup” of pain receptors and peripheral or central pain pathways, we minimize the development of pain and make it easier to manage. Benefits of preemptive analgesia include a more stable patient throughout surgery, a smoother postoperative recovery period, and a reduction in postoperative pain and distress (Gaynor and Muir 2014). The preemptive use of opioids, nonsteroidal anti-inflammatory agents, and local anesthetics forms the foundation of most preemptive interventions. In addition, agents such as α2 adrenergic agonists (e.g., xylazine and medetomidine), N-methyl-D-aspartate (NMDA) antagonists (e.g., ketamine), and corticosteroids are also used.
The same pain prevention drugs listed above are used to treat pain once it has developed. The drugs chosen will depend on the species, age, nature, severity of the pain, and possibly the research objectives. Combinations of analgesics are often used both preemptively and to treat pain. This strategy is called balanced or multimodal analgesia. It involves the simultaneous administration of two or more drugs classes that have additive or synergistic analgesic effects. An added benefit of using two or more drugs with additive or synergistic effects is that the dosages of each agent can often be reduced, leading to fewer adverse effects and lower incidence of drug tolerance.
Use of Pharmaceutical-Grade versus Non-Pharmaceutical-Grade Compounds
A pharmaceutical-grade compound/substance (PGC) is any active or inactive drug, biologic, or reagent for which a chemical purity standard has been established by a recognized national or regional pharmacopeia (e.g., the U.S. Pharmacopeia [USP], British Pharmacopeia [BP], European Pharmacopoeia [EP], or Japanese Pharmacopeia [JP]). These standards are used by manufacturers to help ensure the products are of the appropriate chemical purity and quality, in the appropriate vehicle solution or compound, stable, safe, and efficacious (AAALAC 2015). In addition, the Food and Drug Administration (FDA) maintains a database listing of FDA-approved commercial formulations for both FDA-approved human drugs (U.S. FDA 2016b) and veterinary drugs (U.S. FDA 2016a).
In the United States, it is a requirement that compounds used for the clinical treatment of animals or to prevent, reduce, or eliminate animal pain or distress be PGCs whenever possible (USDA Policy 3”“Veterinary Care, March 25, 2011). In addition, when compounds are used to accomplish the scientific aims of the study, PGCs are preferred if available and suitable. These issues pertain to all components, both active and inactive, contained in the preparation to be administered. Therefore, the vehicle used to facilitate administration of a compound is as important of a consideration as the active compound in the preparation. The NIH OLAW suggests that the IACUC, in making its evaluation, should consider factors such as grade, purity, sterility, acid–base balance, pyrogenicity, osmolality, stability, site and route of administration, compatibility of components, unwanted effects and adverse reactions, storage, and pharmacokinetics (http://grants.nih.gov/grants/olaw/faqs.htm#662). The Guide (NRC 2011) further states that the use of non-PGCs in laboratory animals must be described and justified in the animal use protocol and/or be covered by an IACUC policy developed for their use and approved by the IACUC. There are many useful institutional guidelines available that can be used as templates for the development of an individualized program (NIH 2013).
Medical Records
Medical records document animal care and use, and they are an essential part of any laboratory animal care program (USDA Animal Care Policy 3; Field et al. 2007). This is true for all laboratory animal species, and the following recommendations apply to both USDA-regulated and nonregulated species. Some legislation and guidelines require the use of medical records. The U.S. AWR (9 CFR §2.35) require maintenance of some records for at least 3 years and study-related records for 3 years after study completion. European Union member states are required to maintain some records for 3 years after the death of an animal and other records for 5 years (European Parliament and the Council of the European Union 2010). The institution and attending veterinarian should determine the method by which records are maintained, and professional judgment and performance standards should be used in creating facility standards for medical records. Veterinarians should be responsible for the oversight of records, although records should be readily retrievable and available for all staff involved in animal care. Records should also be reviewed during IACUC facility inspections.
Medical record entries should be dated, legibly written, and clearly state who recorded the entry. Entries are made by those administering treatments, performing procedures or observations, evaluating test and exam results, and uploading or attaching files. Entries should be complete enough to reconstruct the care and use of the animal and allow for clear communication between research and care staff. Recommendations of what to include in records are provided in Figure 31.3. Individual records are the most common, but group records and/or entries may be utilized when performing the same procedure on a group of animals.
Figure 31.3
Recommended items to include in laboratory animal medical records.
Medical records may be on paper or in electronic systems. Electronic records are more legible, automatically include entry dates and personnel, and may be capable of quickly generating reports for easily assessing study complications, mortality rates, and colony health. They also provide an efficient mechanism for including information in large numbers of records at the same time. They can also be utilized to easily schedule treatments, testing, and research procedures. However, electronic systems are dependent on having a reliable and secure server, easy access for updating at the point of use, and payment of licensing fees, and therefore may be cost-prohibitive and unavailable without server or computer access.
Summary
The timely provision of quality veterinary care is a critical component of all laboratory animal care and use programs. Quality veterinary care not only ensures the health, well-being, and welfare of the animals, but also improves the quality, quantity, and reproducibility of the research. Because a program is only as strong as the individuals staffing it, the strength of a veterinary care program is determined by the training and experience of the attending veterinarian or designated program official and his or her staff.
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