The appropriate use of animal models in biomaterials research allows prospective, controlled evaluation of disease processes and candidate therapeutics in a manner that is impossible in human patients or volunteers. Investigators must consider scientific, practical, and humane issues when developing studies that use animals. High-quality research requires close collaboration between veterinary and research professionals to guide model selection and development, minimize animal pain and distress, and to advance the scientific goals of the project. This chapter briefly reviews ethical and regulatory issues, describes available information resources and discusses the design of surgical facilities, animal selection, anesthesia, and analgesia.
Investigators and research institutions have an ethical and legal responsibility to consider animal welfare concerns in research using animals. Russell and Burch expressed the most widely understood ethical principles governing humane design of experiments using animals in 1959, summarized in the concept of the “three Rs:” replacement, reduction, and refinement (Russell and Burch, 1959). The first principle, replacement, states that non-animal models should be used instead of animals to the maximum extent possible. Although definitive safety and efficacy evaluation requires animal models, in vitro and ex vivo biocompatibility and efficacy screening methods are increasingly common in biomaterials research (Fujibayashi et al., 2003; Kirkpatrick et al., 2007; Pariente et al., 2000). These techniques reduce the total number of animals used, and may allow significant savings in time and resources compared to experimental surgical models. The principle of reduction states that investigators should use the minimum number of animals to allow statistically valid inferences to be drawn from the data. In this context, it is as important to avoid using too few animals as too many, because a study that is too small may require repetition. Important considerations in determining the statistical power of an animal experiment include inherent variability of the model, and expected efficacy of the test therapeutic. Consultation with a biostatistician may be useful to estimate required animal numbers. The most important of the “three Rs” is the principle of refinement, which states that investigators should use the least invasive and most modern techniques possible to minimize animal pain and distress (Orlans et al., 1998). Continuous refinement and improvement in animal husbandry and the diagnosis and control of infectious disease has greatly reduced nonexperimental morbidity and mortality in modern research facilities. Surgical models in particular have benefitted from advances in veterinary anesthesia, instrumentation, and monitoring, which have allowed further reduction and refinement of research animal use.
Widespread acceptance of the “three Rs” and continued public scrutiny of biomedical research oblige investigators and institutions to comply with strict regulatory standards governing all aspects of research animal use. In the United States, the United States Department of Agriculture (USDA), Public Health Service (PHS), and the Food and Drug Administration (FDA) are the primary agencies which regulate animal research. Federal animal welfare regulations embody the US Government Principles for the Utilization and Care of Vertebrate Animals Used in Testing, Research, and Training (National Research Council, 2010). The Guide for the Care and Use of Laboratory Animals (Guide) (National Research Council, 2010) elaborates upon the US Government Principles, and provides important reference materials to assist implementation. Federal regulations and the Guide assign primary responsibility for an animal research program to the Institutional Animal Care and Use Committee (IACUC). The institution’s chief executive officer appoints the IACUC, which must minimally include a veterinarian, a scientist, a nonscientist, and an unaffiliated member. One member may fulfill more than one of these requirements. The veterinarian must have appropriate training and experience for the species and models in use. The IACUC has multiple responsibilities. It reviews and approves in advance all animal research protocols to ensure compliance with the US Government Principles. In addition, the IACUC regularly inspects the animal research facility, reviews the animal care and use program, and investigates allegations of animal mistreatment.
The specific focus of animal welfare regulations varies by agency. Well-publicized incidents (Wayman, 1966) of pet theft and their subsequent mistreatment in research laboratories prompted the United States Congress to enact the Animal Welfare Act (AWA) in 1966. The AWA required the Federal government to develop a mechanism to protect animals used in research, for exhibition, and sold as pets in interstate commerce. The AWA does not regulate the use of rats, mice or birds bred for research. In many institutions, the term “regulated species” refers to animals subject to the AWA. The USDA enforces the AWA through the Animal and Plant Health Inspection Service (APHIS). Registered facilities file annual reports of animal usage with USDA, and are subject to unannounced inspections of facilities and records by APHIS inspectors.
The Good Laboratory Practices (GLP) Act (21CFR58.90) requires the United States Food and Drug Administration to regulate preclinical research conducted in direct support of new drugs or medical devices. The primary goal of the GLP regulations is to ensure reproducibility and integrity of data used to support new drug or device applications. The animal care provisions of the GLP are general. Animals must be “free of any disease or condition that might interfere … with the study.” Standard operating practices (SOPs) must govern housing, feeding, and care of animals, and the food, water, and environment must be free of known contaminants.
The United States Public Health Service (PHS), through the National Institutes of Health (NIH), is the largest single sponsor of biomedical research using animals, and has adopted regulations to ensure humane and scientifically valid use of animals. The NIH Office of Laboratory Animal Welfare (OLAW) develops and implements standards for animal care and use at recipient institutions, as formulated in the Public Health Service Policy on Humane Care and Use of Laboratory Animals. Institutions receiving PHS support must file an “Animal Welfare Assurance” with OLAW indicating sufficient institutional resources to provide proper husbandry and veterinary care. Institutions must also indicate how they will approve and monitor research to ensure scientific integrity and prevent inhumane treatment of animals. Assurances are approved for five years, after which a new application must be submitted. The PHS Policy applies to all vertebrate animals, even those not regulated by the AWA (Office for the Protection from Research Risks, 1996).
The Association for the Assessment and Accreditation of Laboratory Animal Care, International (AAALAC) is a nongovernmental independent organization that assists and accredits research institutions seeking to maintain the highest standards of laboratory animal care and use. Approximately 770 research facilities in the United States and in 29 other countries hold AAALAC accreditation. AAALAC policies are determined by the Board of Trustees, representing more than 50 scientific, educational, and professional organizations involved with biomedical research use of animals. AAALAC uses the Guide and applicable governmental guidelines to evaluate the quality of an institutional animal care and use program. The AAALAC Board appoints the Council on Accreditation which reviews, grants or suspends accreditation based on a triennial program review and site visit by at least one Council member and an ad hoc consultant. OLAW accepts AAALAC accreditation as strong evidence that an institution’s animal care and use program is in substantial compliance with the PHS Policy. AAALAC guidelines apply to all live vertebrate animals used in research.
Research using endangered species is generally allowed only for the direct benefit of animals with spontaneous disease. Although rare in biomedical research laboratories, the use of endangered species is regulated by the Endangered Species Act. Use of endangered species is governed by the Convention on International Trade in Endangered Species (CITES), and regulated by the Fish and Wildlife Service of the US Department of the Interior. The provisions of CITES apply to all animals, living or dead. Other agencies which regulate the use of research animals include the Centers for Disease Control (importation of non-human primates and other animals which may harbor zoonotic agents) and the International Air Transport Association (transportation of living animals by air) (National Research Council, 2010). The use of narcotics and barbiturates in experimental anesthesia and analgesia is regulated by the US Drug Enforcement Administration (DEA), which requires users to obtain proper permits, ensure appropriate drug security, and maintain proper records.
Research animals require specialized care that recognizes anatomic and physiologic similarities and differences between species. These differences determine routine husbandry and behavioral needs, and may influence model selection and interpretation of data. Animals vary greatly in their response to anesthetics and other drugs, and anatomical differences often pose significant challenges to endotracheal intubation, surgical approach, and intravenous access.
Veterinarians offer important expertise to investigators using animals. Laboratory animal medicine is a subspecialty of veterinary medicine which involves additional training and acquisition of skills in the diagnosis and treatment of laboratory animal diseases, development of new experimental techniques, and provision of specialized preoperative, intraoperative, and postoperative care. Most large research institutions have at least one staff veterinarian trained and experienced in laboratory animal medicine. Smaller institutions may use consultant veterinarians. Large research programs may also include veterinarians as part of the research team. Other veterinary specialties of potential value in a biomaterials research program include surgery, anesthesia, pathology, and internal medicine. The Academy of Surgical Research (ASR) and the American Association of Laboratory Animal Science (AALAS) sponsor technician training and certification programs in routine and specialized animal care and use. Certified technicians and technologists can provide valuable support and management roles in the experimental surgery laboratory.
Many information resources are available to biomaterials investigators using animals. Several excellent textbooks discuss general laboratory animal medicine (Fox et al., 2002) and basic biology and methodology for rodents (Sharp and LaRegina, 1998), rabbits (Manning et al., 1994; Harkness and Wagner, 1995; Suckow and Douglas, 1997), swine (Bollen et al., 2000; Swindle, 2007) and small ruminants (Borkowski and Allen, 1999). Specialized texts provide detailed reviews of anesthetic techniques (Tranquilli et al., 2007; Fish et al., 2008), infectious disease (National Research Council, 1991; Straw et al., 2006), and drug dosages (Hawk et al., 2005; Plumb, 2005) for experimental animals. Finally, the Internet has excellent resources for laboratory animal users, including reference material, bibliographical databases, and discussion groups on a variety of technical and regulatory topics. The NetVet portal is an excellent entry point to web-based laboratory animal resources (http://netvet.wustl.edu/vet.htm). Other useful websites include OLAW (http://grants.nih.gov/grants/olaw) and the Animal Welfare Information Center (http://awic.nal.usda.gov/).
Quality surgical research requires the use of appropriately designed, equipped, and managed surgical facilities. The Guide requires the use of aseptic technique for all survival surgical procedures on laboratory animals, including rodents. Essential components of aseptic techniques include patient preparation (clipping and disinfection of the surgical site), surgeon preparation (surgical attire, surgical hand scrub, sterile gloves), and the use of sterile instruments. Effective training and staffing levels are necessary to reduce the chance of contamination of instruments and the sterile field (draping, traffic control) (National Research Council, 2010).
Surgical suites represent significant institutional investments that must remain functional for many years. Design and configuration of operating room facilities requires careful planning and consultation with users, veterinarians, laboratory planners, and engineers to meet current scientific and regulatory requirements with appropriate flexibility for possible future changes.
Minimum functional components of the survival surgery suite include dedicated areas for surgery, animal preparation, surgeon’s scrub, postoperative recovery, and surgical support. In the United States, Federal regulations require the use of dedicated surgical suites for major survival surgery on regulated species other than rats, mice, and birds. Surgical suites must allow aseptic surgery, animal preparation, and surgeon scrub to occur in separate dedicated rooms, arranged to facilitate entry and exit of animals and staff while minimizing unnecessary traffic. Animal recovery may often occupy the same room as anesthesia induction and preparation. The Guide permits rodent survival surgery in any procedure space provided necessary conditions for asepsis are present (National Research Council, 2010).
Many research institutions maintain centralized experimental surgery suites to maximize economies of scale and efficient use of skilled support staff. Support spaces in large surgical suites are generally configured to allow two or more operating rooms to run simultaneously from shared animal and surgeon preparation rooms. Smaller specialized facilities may be appropriate to accommodate unique experimental requirements. Principles of research animal operating room design have been described (Ruys, 1991; Hessler and Lehner, 2008; National Research Council, 2010).
Other functional areas commonly required in an experimental surgery suite are: pharmacy, including secure storage of controlled substances; instrument cleaning and sterilization; and record keeping. The use of inhalant anesthesia requires appropriate active scavenging capability. Secure data collection and storage require network connections, and possibly other information technology capabilities. Radiography and other imaging modalities are often necessary in experimental surgical studies, and may be conveniently located within or adjacent to the surgery suite.
Careful preoperative preparation is necessary before invasive surgical procedures. Preoperative preparation includes fasting, premedication, establishment of vascular access, and surgical site preparation. Removal of food and contact bedding is necessary for dogs, swine, and ruminants to prevent regurgitation and aspiration pneumonia. Gastric dilatation may also occur in improperly fasted swine and ruminants. Preoperative fasting is not routine in rodents and rabbits. These species have a high metabolic rate, and are prone to hypoglycemia from prolonged withdrawal of food. Furthermore, they do not vomit and therefore are unlikely to develop aspiration pneumonia. Fasted or not, many species hoard food between the upper molars and the cheek. For this reason the anesthetist should carefully examine the oral cavity during anesthetic induction and remove materials which can be aspirated or pushed into the trachea during endotracheal tube placement. Premedication with analgesic and sedative drugs reduces animal anxiety, facilitates anesthetic induction, and prevents postoperative hyperalgesia and allodynia.
Hypothermia impairs wound healing, and predisposes animals to infection and other postoperative complications. Anesthesia itself is a major risk factor for hypothermia because anesthetic-induced peripheral vasodilation increases conductive heat loss. Many anesthetic and analgesic agents interfere with hypothalamic thermoregulatory systems and the shivering reflex. Cold dry anesthetic gases promote evaporative heat loss from the respiratory tract. Prolonged exposure of the abdomen or thorax during surgery allows heat loss through radiation. Small animals such as rats and mice develop hypothermia more rapidly than larger animals, because they have a large surface area in proportion to body weight. A comprehensive approach to maintenance of normothermia should include minimization of surgical time and exposure of body cavities, and the use of warmed fluids and humidified gases. Care and frequent body temperature monitoring are necessary to prevent hyperthermia, which can rapidly be fatal.
Supplementary heat sources including heated surgical tables and circulating warm air or water blankets are extremely valuable. A recent report evaluated rectal temperature in isoflurane-anesthetized Sprague-Dawley rats and CF-1 mice placed on a circulating warm water blanket or a reusable microwavable warming pad (SnuggleSafe®). Control animals were placed on a stainless steel surgical table with a surface temperature of 21°C–22°C. Surgery was not performed. All control animals became severely hypothermic by the end of the 60 minute test period, and mice lost significantly more heat than rats (−9.9°C versus −4.42°C from baseline). Animals which received supplemental heat support from either source remained essentially normothermic throughout the study (Taylor, 2007).
Preoperative placement of a venous cannula and intraoperative intravenous fluid administration maintains tissue perfusion and electrolyte balance, and allows delivery of emergency drugs when necessary. Appropriate fluids and administration rates for intraoperative maintenance include lactated Ringers solution or normal saline, 5–10 ml/kg/hr. Intraoperative hydration allays dehydration if delayed recovery or physiologic compromise delays spontaneous re-feeding after surgery. Intraperitoneal or subcutaneous fluid administration is a practical alternative to the intravenous route in low risk rats and mice, because intravenous cannulation is difficult in these species.
Minimum monitoring should include core temperature, heart rate, respiratory rate, oxygen saturation, and end-tidal carbon dioxide concentration. Cost-effective devices readily allow determination of these parameters for rabbits and larger animals. Specialized devices are available for reliable assessment of rats and mice. Monitoring of additional parameters such as electrocardiogram, expired anesthetic agent concentration, and arterial blood gases or hematologic or serum biochemical parameters may be indicated for specific studies and will require specialized instrumentation and training.
Anesthesia provides reversible elimination of pain and is essential to all surgical procedures. The ideal anesthetic preserves cardiac output and other physiologic parameters, provides intraoperative analgesia that extends to the postoperative period, and is reversible and safe for the patient, surgical personnel, and environment. Experimental animal surgery almost always occurs under general anesthesia, which additionally renders the animal unconscious and immobile. Local or regional anesthetic techniques are often useful to reduce general anesthetic and postoperative analgesic requirements in major procedures.
Anesthetic agents for experimental surgery may be administered by injection or inhalation. Several considerations should determine the selection of anesthetic agents for experimental surgery. Balanced anesthetic techniques combine two or more sedative, hypnotic, and analgesic agents to reduce adverse effects and increase physiologic stability and analgesia compared to single-agent methods. The Guide requires institutional approval and veterinary participation in the selection of anesthetic agents for experimental surgery (National Research Council, 2010).
Selection of anesthetic technique will vary with the experiment. The anesthetist must maintain oxygenation, perfusion, metabolic balance, and normal core body temperature. Severe or prolonged alterations of these parameters may impair wound healing, predispose to infection or introduce significant scientific variation. Most surgical studies use young adult healthy animals which present minimal anesthetic risk for brief procedures. Unexpected disease may occur in any animal, however, and for this reason all animals should receive appropriate physical examination before anesthetic induction. Extensive procedures or studies with diseased animals pose significant anesthetic challenges and generally require an anesthetist with specialized training and experience.
The American Society of Anesthesiologists (ASA) anesthetic risk assessment categories classify anesthetic risks based on the animal’s age, underlying health status, and other risk factors. Anesthetic, intraoperative and postoperative care and resource requirements can then be determined appropriately. A six-month-old normal Sprague-Dawley rat undergoing implantation of a permanent venous catheter, for example, would be considered ASA category I and require standard anesthetic protocols. By contrast, an 18-month-old Sprague-Dawley rat with advanced experimental myocardial insufficiency following coronary artery ligation would be considered ASA category III or IV, and require correspondingly more intense anesthetic support and monitoring (Tranquilli et al., 2007).
Anesthetics are potent medications which may confound experimental results, either directly or through alteration of normal physiology. Investigators should select agents carefully, and maintain anesthetic and analgesic protocols constant across groups and between experiments to minimize these effects. Pilot studies may be necessary to identify the best agent for specific models, as the following recent report illustrates. Weanling swine were used to evaluate colorimetry, transepidermal water loss, and laser Doppler perfusion imaging to determine the progress of wound healing and efficacy of candidate treatments. Animals were anesthetized with each of three different protocols: ketamine/xylazine (K/X); tiletamine/zolazepam/xylazine (T/X); or tiletamine/zolazepam/xylazine/isoflurane (T/X/I) three times at 24–48 hour intervals, separated by a seven day washout period. The T/X and T/X/I regimens depressed transepidermal water loss less than did K/X. The mechanism was unknown, but possibly associated with differential effects of anesthetic drugs on cutaneous blood flow. The study also identified a progressive reduction in cutaneous blood flow readings during the three day experiment, presumably the result of anesthetic accumulation following multiple doses (Graham et al., 2004).
Injectable anesthesia is popular in experimental surgery. The principal benefits of injectable anesthetic techniques are ease of administration and operator safety. Injectable techniques usually use a combination of agents given by the intramuscular or, for rodents, intraperitoneal routes. Intravenous anesthesia, by continuous or intermittent bolus infusion, may be appropriate in larger animals. Most injectable agents or combinations provide 15–30 minutes of anesthesia. Significant individual, species, and strain variation in the response to injected anesthetics is common. Injectable anesthetics are often used to induce anesthesia before endotracheal intubation and maintenance inhalation anesthesia. Prolonged recovery and physiologic imbalance may follow repeated administration of injectable agents to maintain anesthesia during long procedures. As reversal of injectable agents is often impossible, the operator must avoid administration of an accidental overdose. In addition, the operator must be careful not to breach aseptic technique when administering supplemental anesthetic. Many injectable anesthetic agents have a moderate-to-high abuse potential; therefore, investigators who use these agents in research must register with the United States Drug Enforcement Administration, and maintain proper records and storage facilities.
Inhalation anesthesia, usually with isoflurane or sevoflurane, allows the operator a high degree of control over anesthetic depth, and is the technique of choice for prolonged or invasive surgery. Inhalation anesthesia for major surgery requires a precision vaporizer and endotracheal intubation to allow proper control of anesthetic depth and the airway. Controlled ventilation further facilitates maintenance of normal tissue oxygenation and carbon dioxide balance, and allows rapid increase or decrease of anesthetic depth when necessary. Controlled ventilation is mandatory for intrathoracic procedures, because many laboratory animals lack a complete mediastinum and develop bilateral pneumothorax following thoracotomy. Endotracheal intubation requires special laryngoscopic equipment and training to prevent laryngeal trauma or esophageal placement.
Anesthetic administration by facemask may be appropriate for brief or noninvasive procedures. Prolonged inhalation by mask is undesirable because it does not protect the airway from aspiration in the event of vomiting or allow positive pressure ventilation in an emergency. A tight fitting mask and active scavenging are necessary to maintain anesthetic depth and reduce operator exposure to waste anesthetic agents.
Ketamine and tiletamine are dissociative anesthetics commonly used in experimental surgery because they maintain cardiac output and provide good analgesia with a high margin of safety. Dissociative anesthetics are NMDA (N-Methyl-D-aspartate) antagonists which produce unconsciousness and analgesia through selective disruption of ascending impulses to conscious brain centers, rather than through generalized depression of the central nervous system. Dissociative anesthetics are almost always used in combination with other agents. When used as a sole agent, surgical anesthesia is rarely achievable at safe dosage levels. Other significant disadvantages include unacceptable muscle rigidity, salivation, and seizures, especially in dogs (Lin, 2007; Tranquilli et al., 2007; Meyer and Fish, 2008).
Ketamine solutions are acidic, and perivascular infiltration and large volume intramuscular injection will produce pain and tissue irritation. Telazol® is a commercial mixture of the dissociative agent tiletamine and the benzodiazepine zolazepam. The product is supplied in a sterile vial, which is reconstituted with 5 ml of sterile water. Reconstituted vials contain 50 mg/ml of each agent and may be kept for 48 hours at room temperature or for 14 days in a refrigerator. Telazol® has similar anesthetic efficacy as ketamine/xylazine or ketamine/diazepam. Telazol® may also be combined with xylazine or medetomidine for greater anesthetic depth.
Xylazine (Rompun®), medetomidine (Domitor®) and dexmedetomidine (Dexdomitor®) are common adjuncts to ketamine anesthesia. These agents are alpha-2 adrenergic agonists with potent sedative and analgesic activity. They lack adequate potency to be sole agents for general anesthesia, but may provide adequate sedation for dressing changes, suture removal, and other minor procedures. Coadministration with ketamine generally provides up to 30 minutes of anesthesia with excellent muscle relaxation, smooth recovery, and a moderate degree of postoperative analgesia in many species. These combinations are also useful for anesthetic induction prior to endotracheal intubation and subsequent inhalation anesthesia. Intramuscular administration is preferable, because transient hypertension and cardiac arrythmia can occur after rapid intravenous boluses. Xylazine is the most commonly used alpha-2 agonist in rodents. Medetomidine has increased specificity for the alpha-2 receptor, and increased analgesic potency compared to xylazine. Medetomidine is preferred over xylazine in dogs (Lemke, 2007). Medetomidine is a racemic mixture of the active R and inactive L isomers. Dexmedetomidine is a pure preparation of the R-isomer and produces physiologic and analgesic effects similar to medetomidine at approximately half the dose.
Several side-effects are associated with alpha-2 agonists and are more significant following IV administration. Xylazine commonly induces vomiting in dogs. All alpha-2 agonists significantly depress cardiac output at standard anesthetic doses and may cause severe bradycardia. Cardiac depression is frequently subclinical in young healthy animals. Decompensation may occur in aged animals or those with clinical or experimentally induced illness. The mechanism of alpha-2-agonist associated bradycardia is complex and may require combined anticholinergic (atropine or glycopyrollate) and specific antagonist (yohimbine or atipamezole) therapy.
A distinct advantage of the alpha-2 agonist sedatives is the availability of specific antagonist drugs. Yohimbine effectively reverses xylazine, and atipamezole is the recommended reversal agent for medetomidine (Lemke, 2007). These drugs effectively speed recovery following minor procedures, and may also be used in an anesthetic emergency.
Other adjuncts may also be used to allay animal anxiety and provide muscle relaxation. Acepromazine provides excellent sedation in many species, and protects the myocardium from catecholamine-induced arrhythmias. Disadvantages of acepromazine include hypotension secondary to adrenergic blockade and prolonged recovery. Benzodiazepines such as diazepam, midazolam, and zolazepam provide excellent short-term sedation with little effect on blood pressure (Lemke, 2007).
Propofol is a non-narcotic, injectable sedative and hypnotic agent commonly used for anesthetic induction and for brief procedures in dogs, swine, and small ruminants. Propofol is insoluble in water and is available as an oil-based emulsion in single-dose ampules and multiple-dose vials for intravenous administration. Bacterial growth will occur rapidly in contaminated solutions, and vials should be used or discarded the day they are opened. Anesthetic induction and recovery are smooth. The duration of anesthesia following a single bolus intravenous injection in dogs is approximately 20 minutes. Prolonged anesthesia is possible using intermittent bolus injections or continuous rate infusions. Propofol has little analgesic activity, and should be used in combination with narcotics or alpha-2 agonists for painful procedures. Side-effects of propofol include apnea, bradycardia, and hypotension. The incidence of apnea is related to the total dose and rate of injection, and resolves spontaneously as the drug distributes. Manual ventilation may be necessary if apnea is prolonged. Human patients commonly report transient pain at the injection site. Unlike ketamine and the barbiturates, accidental perivascular propofol injection does not produce tissue damage (Branson, 2007).
The barbiturates are among the oldest anesthetic agents and remain useful for some experimental applications. Barbiturates act through general depression of the central nervous system. There is dose-dependent depression of respiration and cardiac output. Hepatic metabolism terminates the activity of long-acting oxybarbiturates such as sodium pentobarbital. Rapid redistribution into adipose tissue followed by hepatic detoxification characterizes the metabolism of the short-acting thiobarbiturates thiamylal and thiopental.
The principal advantage of sodium pentobarbital for general anesthesia is the ability to rapidly induce deep anesthesia with a single agent. Several disadvantages limit sodium pentobarbital’s utility in prolonged or invasive procedures. Progressive cardiovascular depression occurs with prolonged anesthesia. Despite sleeping times of 5–15 hours, surgical anesthesia is often present only for 30–60 minutes in most species because this agent provides very little analgesia. Stormy recovery with vocalization and an unstable gait is common. Rapid-acting intravenous thiobarbiturates provide about 10 minutes of general anesthesia, and are useful for anesthetic induction. Barbiturate solutions are strongly alkaline and require intravenous or intraperitoneal administration to avoid pain and tissue necrosis (Branson, 2007). The therapeutic index of most barbiturates is very low: most laboratories use these agents primarily for euthanasia.
Specific experimental situations may require the use of nonstandard anesthetics. These agents include chloral hydrate, alpha-chloralose or urethane. Chloral hydrate and alpha-chloralose preserve motor and sensory nerve function, but are poor anesthetics with minimal analgesic effect except at very high dosages. Alpha-chloralose and urethane result in stable cardiovascular performance during prolonged anesthesia. Urethane is mutagenic and carcinogenic in experimental animals, and may be used only for nonsurvival procedures. Urethane may also be hazardous to research staff after prolonged contact. Because of these constraints, most IACUCs require investigators to justify the use of nonstandard agents on scientific grounds (Meyer and Fish, 2008).
Isoflurane and sevoflurane are the agents of choice for clinical and experimental inhalation anesthesia. These agents are poorly soluble in blood and undergo minimal metabolism. As a result, recovery after anesthetic discontinuation is rapid, even in animals with significant hepatic or renal impairment. Dose-dependent cardiac depression may occur during prolonged procedures. A balanced anesthetic technique combining narcotic analgesics and muscle relaxants to reduce the required concentration of isoflurane is suggested for long procedures on animals with cardiac disease. Unlike the obsolete agents halothane and methoxyflurane, isoflurane and sevoflurane do not sensitize the myocardium to catecholamine-induced arrhythmias (Steffey and Mama, 2007; Brunson, 2008). As sole agents, isoflurane and sevoflurane are relatively poor analgesics and should be used in combination with sedatives and narcotics or nonsteroidal anti-inflammatory drugs (NSAIDs) for optimal intraoperative and postoperative pain control. These anesthetics are halogenated drugs which must be scavenged with activated charcoal canisters or by connection to an active exhaust circuit (Steffey and Mama, 2007; Brunson, 2008).
Older inhalation anesthetics occasionally used in experimental surgery include diethyl ether, chloroform, and nitrous oxide. Although ether is inexpensive and readily available from chemical supply houses without prescription, significant disadvantages include flammability and the risk of explosion if peroxides form when ether evaporates to dryness. Mask induction with ether is unpleasant for the patient. High lipid solubility prolongs recovery and analgesia is poor. Repeated exposure to chloroform or methoxyflurane is toxic to the operator. Nitrous oxide provides slight analgesia when administered at high concentrations, but is not an effective anesthetic in animals. The maximum allowable concentration of nitrous oxide is 80% in oxygen, beyond which hypoxemia is likely. As activated charcoal does not absorb nitrous oxide, active exhaust scavengers are required when using nitrous oxide for anesthesia. Nitrous oxide diffuses out of the bloodstream to gas-filled body cavities such as the gastrointestinal tract. This property may cause significant distension of the rumen in sheep and goats. Most institutions strongly discourage the use of these agents without specific scientific justification (Brunson, 2008).
Bell jar administration of volatile anesthetics is no longer acceptable in most institutions. Chloroform, ether, and methoxyflurane have a low vapor pressure and achieve anesthetic concentrations when allowed to evaporate in a closed container, but these agents pose unacceptable risks to animals and operators. Isoflurane and similar modern agents are highly volatile and rapidly attain toxic levels in a nose cone or closed container. Investigators familiar with bell jar administration of obsolete agents commonly experience unacceptable animal mortality when attempting to use isoflurane in the same manner.
Prompt recognition and adequate treatment of postoperative pain are key responsibilities of experimental surgeons. Experimental surgical procedures rarely, if ever, directly benefit the research subject and impose a substantial moral requirement on investigators to prevent or minimize any discomfort related to the experiment. The scientific community has accepted that comparable mechanisms govern the production and response to pain in animals and humans, as the widespread use of animal models in pain research confirms. Animal welfare regulations and accreditation guidelines require the use of sedatives, analgesics or anesthetics for procedures which “may cause more than momentary or slight pain or distress” (Office for the Protection from Research Risks, 1996). Additionally, untreated pain increases catecholamine secretion and causes stress, which introduces experimental variation by impairing wound healing and immune function (Lee-Parritz, 2007). Animal welfare regulations require investigators to justify withholding of analgesics to the IACUC on scientific grounds.
Recognition of postoperative pain is difficult in most laboratory animal species. Anorexia, lethargy, piloerection, and wound hypersensitivity indicate the presence of moderate to severe pain, but absence of these signs does not necessarily prove that the animal is pain-free. Behavioral and physiological factors complicate the diagnosis of less severe pain, which may still require treatment to assure animal well-being. Rodents, for example, are nocturnal animals and are normally less active during the day than at night. Rodents typically nibble small amounts of food at frequent intervals. As a result, pain induced inhibition of activity or feeding is difficult for research staff to recognize. In addition, rodents issue distress calls at ultrasonic frequencies that are inaudible to human beings (Lee-Parritz, 2007). Most animals effectively conceal subtle signs of pain and distress from research staff. Animal husbandry staff may notice subtle signs of pain or distress in research animals that are not apparent to investigators. For these reasons, animal welfare regulations counsel investigators that surgical procedures that would induce pain in a human would produce pain in an experimental animal, unless convincing evidence to the contrary exists (National Research Council, 2010).
Narcotics remain the analgesics of choice for severe pain in most species. Adverse effects of narcotics include dose-dependent sedation, hypoventilation, anorexia, and constipation. These effects are more pronounced with pure agonists such as morphine than with partial agonists, such as butorphanol and buprenorphine. Buprenorphine is widely used in veterinary medicine because it provides excellent analgesia at a convenient dose interval (8–12 hours), and has minimal depressant effects on respiration and cardiac output (Lee-Parritz, 2007; Lamont and Mathews, 2007). Dosage and dose intervals of the narcotic analgesics vary widely between species because of the higher metabolic rate of the smaller species, as well as species-specific responses to the agents. In comparison to larger animals, rodents require very high doses of most narcotics in proportion to their body weight for effective analgesia (Lamont and Mathews, 2007; Lee-Parritz, 2007).
Fentanyl is a highly effective narcotic analgesic. Parenteral fentanyl has a very short half-life, and can be rapidly titrated to effect as a continuous rate infusion in a balanced anesthetic protocol. The fentanyl transdermal patch provides effective analgesia for up to 72 hours as a sole agent or in multimodal analgesic regimens. Several factors may affect transdermal fentanyl absorption, including temperature, skin thickness, and blood supply. Care and consistency in application is necessary to prevent toxicosis or inadequate analgesia. The patch should not be under an occlusive dressing, on a patient’s recumbent side or where it will be in contact with supplementary heat devices used during anesthesia, as these practices may cause significant increase in drug release from the patch. Patches should be placed at least 12 hours preoperatively to ensure adequate blood levels at the time of incision. Poorly applied patches may dislodge and be eaten, causing fatal fentanyl overdose. Facilities which use fentanyl patches should stock naloxone (0.1 mg/kg IV, repeated as necessary) for emergency use in case of accidental patch ingestion (Lamont and Mathews, 2007; Lee-Parritz, 2007).
Nonsteroidal anti-inflammatory drugs (NSAIDs) are most effective against moderate pain of musculoskeletal origin, including incisional pain from ventral abdominal incisions in swine and other large animals. NSAIDs are of particular value in orthopedic procedures. The principal side-effects of many NSAIDs include increased bleeding from reduced platelet activity or gastric irritation secondary to prostaglandin inhibition. Although clinically significant impairment of blood clotting ability is rare, gastric toxicity remains a concern, particularly in dogs (Lamont and Mathews, 2007; Lee-Parritz, 2007).
It is advisable to use multimodal analgesia consisting of local anesthetics, NSAIDs, and narcotics whenever possible. The use of agents from two or more analgesic classes will act at several complementary steps in the pain cascade. This strategy will generally have additive or synergistic activity with reduced potential for drug toxicity. Preoperative analgesic and local anesthetic administration will significantly reduce intraoperative anesthetic requirements, and likely reduce acute and chronic postoperative pain. Local anesthetics are most often administered to the incision or regionally at the time of surgery, but may be of significant value in the postoperative period as well.
The possible effect of analgesic treatment on wound healing is controversial. Most NSAIDs block COX-1 and COX-2 to a variable degree, and COX-2 plays an important role in wound healing. A study of the effect of continuous diclofenac administration for 10 days to rats with experimental dorsal skin incision demonstrated reduced fibroblast numbers in treated versus control rats, but no difference in epidermal thickness or clinical healing (Krischak et al., 2007). In another model, rats undergoing experimental medial collateral ligament transaction demonstrated improved healing at day 14 following 6 days’ treatment with the NSAID piroxicam compared to controls. Animals which received naproxen, rofecoxib, acetaminophen or butorphanol at the same dose schedule demonstrated healing similar to controls (Hanson et al., 2005). A third study evaluated the effect of continuous celecoxib or indomethacin treatment versus controls on bone repair in a rat femur fracture model. Recipients of either NSAID had increased fibrous tissue compared to control at 4 and 8 weeks, but all groups demonstrated equivalent morphological and functional healing by 12 weeks (Brown et al., 2004). A rabbit spinal fusion model demonstrated equal fusion rate and strength at 8 weeks when animals received either ketoprofen or tramadol for 8 days after surgery (Urrutia et al., 2007). These results suggest that NSAIDs may have a small, agent-specific effect on experimental wound healing. The applicability of studies based on extended treatment protocols to the relatively brief treatment periods required for acute pain control requires further study. Narcotic analgesics have not been demonstrated to impair wound healing, and may actually speed recovery in specific settings (Peyman et al., 1994). Consistent administration of standard analgesic drugs across experimental groups, guided by pilot studies when necessary, should allow effective analgesic therapy without compromising the scientific validity of the study.
The following sections provide general recommendations for the selection, care, and experimental use of laboratory animal species commonly used in biomaterials research. When appropriate, separate recommendations will be made for brief or noninvasive procedures. For detailed discussion of anesthesia for specific procedures or disease conditions, the reader is advised to consult the references or a veterinary specialist.
Specific pathogen-free rodents should be used for all surgical procedures to reduce morbidity and mortality from chronic respiratory disease. Infectious agents commonly implicated in chronic respiratory disease of rodents include Mycoplasma pulmonis, Sendai virus, and cilia-associated-respiratory (CAR) bacillus (National Research Council, 1991; Fox et al., 2002). Other infectious agents may alter immune function or impair detoxification of anesthetic or experimental drugs. Several commercial vendors supply common strains of laboratory rodents free from infection with these and other infectious agents. These animals are known as specific pathogen-free (SPF) animals, and should always be used for survival surgical procedures to minimize experimental variability and reduce surgical mortality. The specific panel of excluded agents may vary according to the vendor. The institution should adopt standard operating procedures to prevent introduction of rodent infectious agents, and should regularly survey all holding areas for evidence of infection. Introduction of SPF animals to rooms with enzootic viral infection may result in rapid onset of severe clinical disease. Cedar or pine shavings sometimes used for contact bedding contain aromatic compounds that induce hepatic microsomes and alter hepatic detoxification of anesthetics (Gaertner et al., 2008). To ensure uniform response to anesthetics and experimental drugs, rodents should receive heat-treated wood chip, corncob or cellulose bedding. A conditioning period of at least three days after purchase will ensure that animals have recovered from dehydration and stress associated with shipping. Rodents do not vomit, and preoperative fasting is not recommended. If otherwise indicated, fasting in rodents should be kept to a minimum (2–3 hours) to avoid hypoglycemia and shock (Flecknell et al., 2007).
Most surgical procedures in rodents are brief. Small body size and limited vascular access complicate anesthesia and intraoperative support of rodents. Nevertheless, skilled operators with appropriate instrumentation can accomplish delicate vascular surgery and other procedures in rats and mice with minimal postoperative mortality. Endotracheal intubation and positive pressure ventilation is also possible and requires the use of customized equipment (Gaertner et al., 2008).
Several anesthetic combinations are appropriate for brief, noninvasive procedures in rodents. Tribromoethanol (Avertin®) provides light anesthesia for 10–20 minutes in mice. Prepared solutions must be stored in the dark at 4°C to avoid production of gastric irritant decomposition compounds. The standard anesthetic dose is 0.2 ml/10 gm of a 1.2% solution. This anesthetic is most appropriate for brief, minimally painful procedures in mice, such as retro-orbital blood sampling, embryo transfer, vasectomy, and tail biopsy (Gaertner et al., 2008). Tribromoethanol is contraindicated in rats because peritoneal fibrosis and peritonitis is common following intraperitoneal injection in this species (Reid et al., 1999). Although propofol is intended only for intravenous administration, a recent report described effective surgical anesthesia of 20–30 minutes’ duration and smooth recovery following intraperitoneal administration of propofol (75 mg/kg, mouse; 100 mg/kg rat), medetomidine (1–2 mg/kg, mouse; 0.1 mg/kg rat) and fentanyl (0.15–0.20 mg/kg, mouse; 0.1 mg/kg rat) to CD-1 mice and Wistar rats (Alves et al., 2009, 2010).
Intraperitoneal injection of ketamine (40–100 mg/kg) and xylazine (3–10 mg/kg) provides 20–60 minutes of surgical anesthesia in most rodent species. Muscle relaxation and analgesia are good. Anesthetic duration varies in a dose-dependent manner. If necessary, supplemental administration of ketamine will prolong anesthesia. Ketamine volumes commonly required for rats and mice require division of an IM (intramuscular) dose between two or more sites if the IP (intraperitoneal) route is contraindicated. Yohimbine (1–2 mg/kg IP) will reverse xylazine-associated sedation and speed recovery from anesthesia, but will also likely reverse residual xylazine-induced analgesia. Side-effects of general anesthesia in rodents may include hypercarbia, hypoxemia, and hypotension although these effects are less evident with ketamine and xylazine in comparison with sodium pentobarbital. Medetomidine and dexmedetomidine are safe but short acting in rodent species, and have not replaced xylazine in most models (Gaertner et al., 2008).
Barbiturates such as sodium pentobarbital (30–70 mg/kg IP) are occasionally used in rodent experimental surgery. Disadvantages of these agents include brief periods of effective anesthesia, prolonged recovery, and poor analgesia. In addition to environmental factors discussed earlier, rodents also display marked individual and strain variability in the response to sodium pentobarbital. Pretreatment with buprenorphine prior to incision prolongs the period of effective anesthesia, and lowers the required dose of sodium pentobarbital (Roughan et al., 1999). One report described the use of methohexitone (44 mg/kg of a 6.46 mg/ml solution IP) to achieve 2 minutes of chemical restraint for oral examination in C3H/Neu mice with recovery in 10–15 minutes. The major disadvantage of this technique was a very narrow therapeutic window: 40 mg/kg produced no immobility, whereas 50 mg/kg produced 40% mortality (Dorr and Weber-Frisch, 1999).
Isoflurane provides excellent anesthesia in rats and mice. This agent may be used both for brief restraint and for procedures of up to several hours’ duration. Isoflurane requires the use of a precision vaporizer to maintain consistent anesthetic concentration. Rapid induction of anesthesia occurs following placement of animals into an induction chamber containing 3–4% isoflurane in oxygen. Most animals awake 1–2 minutes after removal from the chamber, which is sufficient time for retro-orbital blood sampling or tail biopsy. To maintain anesthesia for longer periods, the animal’s head and nose may be placed into a customized nose cone connected to a non-rebreathing anesthetic circuit and scavenger. Concentrations of 2–3% isoflurane are commonly used for maintenance. The absence of the pedal withdrawal reflex confirms adequate anesthetic depth. Rodent nose cones are easily fashioned from funnels or disposable syringe barrels. For procedures requiring positive pressure ventilation, endotracheal intubation is easily accomplished. Techniques for endotracheal intubation for rodents have been described (Gaertner et al., 2008).
Buprenorphine is the most widely used analgesic in rodents. Buprenorphine is a mixed mu agonist/antagonist and a kappa receptor agonist with approximately 25–40 times the analgesic potency of morphine. The relative activity at each receptor may be dose-dependent. This agent provides safe and effective analgesia for 6–12 hours after dosing. The current recommended dose for most rodent indications is 0.03–0.05 mg/kg IP or SQ (subcutaneous) (Lee-Parritz, 2007; Curtin et al., 2009).
For procedures under isoflurane or sodium pentobarbital, buprenorphine should be given at least one hour before incision. The anesthetist should be aware of the anesthesia-sparing effects of buprenorphine, and be prepared to reduce the isoflurane vaporizer setting 30–50% as necessary (Brunson, 2008). For procedures conducted under ketamine/xylazine anesthesia, buprenorphine should be given only when the animal has regained sternal recumbency during recovery. Preoperative administration of buprenorphine or other narcotics may cause unpredictable increases in anesthetic depth when using ketamine/xylazine combinations for anesthesia. Local infusion of lidocaine (up to 1 ml/kg of 1.0% lidocaine or 0.25% bupivicaine) before surgical incision will provide additional intraoperative and postoperative analgesia (Lee-Parritz, 2007).
Clinically significant side-effects of buprenorphine include consumption of bedding (pica), and excessive licking or biting of the limbs and cage which may also be directed to the surgical incision. These effects are more common at high doses (0.1–0.3 mg/kg) and can often be managed through temporarily placing animals on a wire grid during the immediate postoperative period or substitution of a synthetic for a natural bedding substrate. A generalized increase in activity and a reduction of ventral grooming may also occur. These effects most likely represent a direct effect of the drug, and do not necessarily indicate the presence or absence of pain (Lee-Parritz, 2007).
The NSAIDs ketoprofen, carprofen, and meloxicam are the next most widely used analgesics. These drugs block the formation of inflammatory mediators associated with surgical injury, and also act centrally to inhibit secondary allodynia. The current recommended dose for meloxicam is 1 mg/kg PO (per os or orally) or SQ in rats, and up to 10 mg/kg in mice. The dose for ketoprofen and carprofen in rats is 5 mg/kg SQ. The dose interval for these agents has not been critically evaluated in rodents, but is 12–24 hours in other species. The NSAIDs are highly effective pre-emptive analgesics when administered one hour or more before surgery, but isoflurane vaporizer settings typically do not require adjustment (Lee-Parritz, 2007).
There are few reported adverse side-effects associated with NSAIDs in rodents. Diffuse intestinal ulceration may occur at high doses, especially in animals with concurrent disease. Most NSAIDs inactivate platelet function to some extent through cyclooxygenase-mediated inactivation of thromboxane. This may slightly prolong the bleeding time, but rarely causes adverse clinical effects in humans even when administered preoperatively (Lee-Parritz, 2007).
Acetaminophen (paracetamol) is a nonsteroidal analgesic which lacks significant anti-inflammatory properties. The drug is readily available in a palatable over-the-counter pediatric syrup formulation (Children’s Tylenol®). Several studies suggest a possible role for this agent in rat analgesia, although specific evaluation for postoperative pain control is lacking. Reported effective dose ranges are 100–300 mg/kg twice a day. These studies suggest that acetaminophen may be effective for mild-to-moderate postoperative pain in rats. Although it is convenient to administer analgesics in drinking water, anorexia in the immediate postoperative period associated with anesthetic recovery and abdominal discomfort may reduce drug intake precisely when pain is maximal. Analgesic activity has not been assessed in mice, for which the LD50 (lethal dose to 50% of the subjects) is close to the doses evaluated for analgesia in rats (Lee-Parritz, 2007).
Infection with the respiratory pathogen Pasteurella multocida is extremely common in conventional rabbits. Colonization of the upper respiratory tract may be clinically silent, but may spread to the lungs, middle ear, and brain in crowded or stressed conditions and produce characteristic disease syndromes (Harkness and Wagner, 1995; Lipman et al., 2008) and compromise long-term surgical studies. Pasteurella-negative rabbit colonies are available and should be used for all surgical protocols. Rabbits do not vomit, and aspiration pneumonia is therefore not a concern. Adult rabbits will tolerate an overnight fast without difficulty, and the resulting reduction in stomach contents may help maintain adequate oxygenation during spontaneous respiration under anesthesia. Rabbits weighing less than 2 kg should not be fasted, as hypoglycemia and metabolic acidosis may be significant (Lipman et al., 2008). Prolonged fasting may cause dehydration and subsequent intestinal motility disorders (Lipman et al., 2008). Traumatic lower-back fracture is common when inexperienced staff handle rabbits. The best way to handle rabbits is to grasp the scruff of the neck with one hand while supporting the rump and hind legs with the other. Sudden onset of flaccid paraplegia in rabbits is almost always the result of lower-back fracture, and warrants immediate euthanasia (Harkness and Wagner, 1995). Rabbits should never be lifted, moved or restrained by the ears.
Combined administration of ketamine (35–50 mg/kg IM) and xylazine (5–10 mg/kg IM) provides excellent anesthesia for a variety of applications in the rabbit. General anesthesia lasts for 30–60 minutes, and provides adequate analgesia and restraint for procedures of moderate intensity. Supplemental use of a narcotic analgesic such as buprenorphine (0.05 mg/kg IM) prolongs anesthesia and improves analgesia. Propofol administered by intravenous bolus or constant rate infusion produces light anesthesia suitable only for intubation or nonpainful procedures such as imaging. Telazol® should not be used for survival procedures in rabbits, because renal tubular damage is a common complication even at standard anesthetic dosages (Lipman et al., 2008).
The intramuscular ketamine/xylazine combination suggested for brief procedures is also an excellent induction agent prior to endotracheal intubation for inhalation anesthesia. Intravenous administration of ketamine and xylazine through the ear vein will achieve rapid induction, but care is required to minimize skin irritation from perivascular infiltration. Mask induction is rarely indicated because apnea, breath holding, bradycardia, and struggling are common when unsedated rabbits are exposed to isoflurane, and because operator exposure to waste anesthetic is difficult to avoid in this setting (Flecknell et al., 1999).
Propofol (10–20 mg/kg IV [intravenous]) is an effective rabbit anesthetic induction agent which produces significantly faster postoperative recovery than ketamine/xylazine in animals maintained on sevoflurane. In a recent report, rabbits anesthetized with propofol and maintained on sevoflurane were extubated 2 ±1 minutes, and achieved sternal recumbency 8 ±0.3 minutes after discontinuation of sevoflurane. Recovery following isoflurane anesthesia should be similarly rapid. By contrast, anesthetic recovery following ketamine/xylazine or ketamine/medetomidine may require up to 120 minutes. Animals should receive preoperative analgesics and be preoxygenated by mask before propofol administration. The initial dose of 10 mg/kg should be administered by hand over 60 seconds (~0.17 mg/kg/s), with additional small increments as required to allow endotracheal intubation. Propofol is not suitable as a sole agent because respiratory arrest may occur, particularly following rapid infusion (>0.25 mg/kg/s) (Allweiler et al., 2010).
Endotracheal intubation of the rabbit may be difficult because of several distinctive anatomic features. The prominent incisor teeth, long oropharynx and limited mobility of the temporomandibular joint hinder direct visualization of the larynx from the front, and require the use of a pediatric laryngoscope with a size 0–1 Wisconsin or size 1 Miller blade. Lidocaine spray on the vocal cords is necessary to prevent further narrowing of the larynx through laryngospasm. Benzocaine spray (Cetacaine®) produces methemoglobinemia in rabbits and should be avoided. The laryngeal opening is often smaller than the diameter of the trachea, requiring the use of a small endotracheal tube (2.5–4 mm). The tongue is short, friable, and difficult to grasp. Supine, prone or lateral positions are all suitable for endotracheal intubation. Hyperextension of the neck will straighten the larynx and facilitate proper tube placement. Blind intubation of the trachea is easily accomplished with practice. The tube is placed in the supraglottic region and advanced toward the larynx in coordination with respiration. Identification of normal breath sounds, visualization of condensate on a dental mirror or capnometry may be used to confirm proper tube placement (Harkness and Wagner, 1995; Lipman et al., 2008). Anesthetic maintenance with isoflurane usually requires a vaporizer setting of 1–4%. Absence of the pinna withdrawal reflex is the best indicator of a surgical anesthetic plane in the rabbit. Many rabbits retain the corneal and pedal withdrawal reflexes even under very deep anesthesia (Lipman et al., 2008; Muir, 2007).
If an anticholinergic agent is required, investigators should be aware that approximately 50% of rabbits produce atropinesterase (AtrE) as a genetically determined trait. Glycopyrrolate resists AtrE, and is therefore recommended over atropine for rabbits. Glycopyrrolate prevents xylazine-associated bradycardia in rabbits (Lipman et al., 2008).
Behavioral signs of postoperative pain in rabbits are vague and inconsistent, and may include lethargy, tooth grinding, and increased activity directed toward the painful area. Anorexia is a common sign of postoperative pain in rabbits. Untreated anorexia can create serious secondary disease, including generalized gastrointestinal stasis, rapid weight loss, and fatal hepatic lipidosis (Lipman et al., 2008).
The best indicator of GI (gastrointestinal) health in the rabbit is the quantity and consistency of fecal pellets. Dehydration and anorexia result in a reduced number of firm, dry pellets. Treatment of anorexic postoperative rabbits should be directed to relief of underlying pain if present and towards restoration of normal gastrointestinal function. A high fiber supplement such as Oxbow Critical Care or blenderized rabbit chow is preferred over low fiber nutritional gels such as Nutrical® to restore normal gut motility. Aggressive analgesia, fluid therapy, and force-feeding may be necessary in some cases.
Narcotic analgesics are preferred for treatment of moderate to severe postoperative pain in rabbits. Buprenorphine (0.01–0.05 mg/kg SQ BID/TID [twice or three times a day]) is the narcotic of choice, because it provides effective analgesia at a convenient dosing interval and is generally well-tolerated. The fentanyl patch (25 μg/hr) is an alternative narcotic analgesic option that provides continuous analgesia up to 72 hours without the requirement for regular injections. Fentanyl patches achieve effective blood levels approximately 12 hours after application, and should be applied the night before surgery. Alternatively, the patch may be applied at the time of surgery, with a single dose of buprenorphine to provide 8–12 hours of bridging analgesia. Clip the fur well and use tissue adhesive (Nexaband® or equivalent) if necessary around the edge of the patch to ensure good skin adhesion. Inadequate fur removal will greatly reduce drug absorption. Use of depilatories such as Neet® for hair removal results in increased drug absorption, and may cause toxicity (Foley et al., 2001).
The NSAIDs are also highly effective in rabbits, particularly for musculoskeletal pain or co-administered with narcotics for multimodal analgesia. Flunixin (1.1 mg/kg SQ q24h), carprofen (4 mg/kg SQ q24h), and meloxicam (0.3 mg/kg PO q 24h) all demonstrate good clinical efficacy in rabbits (Lipman et al., 2008). Oral meloxicam oral suspension (Metacam®, 1.5 mg/ml) has recently undergone pharmacokinetic evaluation in rabbits. Oral administration of 0.3 mg/kg once a day produced blood levels comparable to those shown to be clinically effective in other species for up to 24 hours. Repeated administration of 1.5 mg/kg/day for five days did not result in drug accumulation or drug toxicity. Although rabbits demonstrate more rapid metabolism of meloxicam than other species, delayed gastrointestinal absorption of the drug allows maintenance of effective blood levels throughout the dose period (Turner et al., 2006).
Appropriate vendor selection and conditioning procedures are necessary to identify pre-existing cardiovascular or renal diseases, malnutrition or parasitism that can significantly complicate anesthesia and surgery. Heartworm disease, a result of Dirofilaria immitis infestation, causes eosinophilia and right heart failure. Intestinal parasites, including roundworms (Toxocara canis), hookworms (Ancylostoma caninum), and whipworms (Trichuris vulpis) cause eosinophilia, diarrhea, and general debilitation. All of these parasites are susceptible to common anthelmintics. A calm behavioral profile, reinforced by habituation to the research staff and regular training, is important for animals on long-term studies that call for frequent handling. Purpose-bred dogs have a known pedigree and health history, and often present a more consistent physiological profile than random-source dogs. Random-source dogs are significantly less expensive than purpose-bred dogs, but often have health and behavioral disorders which render them unsuitable for long-term studies (Dysko et al., 2002).
Calm dogs may be trained to cooperate in noninvasive clinical procedures, thereby avoiding the need for sedatives or anesthetics. Short acting anesthetics may be required for more invasive procedures. The preferred technique for short-term injectable anesthesia in dogs is coadministration of a dissociative anesthetic with a sedative or tranquilizer and an anticholinergic drug. All of these drugs are suitable for intravenous administration to achieve rapid induction and recovery, and provide adequate anesthetic depth to allow endotracheal intubation before prolonged procedures. Intravenous ketamine (5–10 mg/kg) and diazepam or midazolam (0.2–0.5 mg/kg) provides 5–15 minutes of light anesthesia suitable for dressing changes, radiography or other minor procedures. Medetomidine (10–40 μg/kg IM) or dexmedetomidine (5–20 μg/kg IM) are also appropriate for this purpose, and may be rapidly reversed with atipamezole (50–200 μg/kg) (Lemke, 2007). Supplementation of medetomidine or dexmedetomidine with ketamine (3 mg/kg) and an opioid such as butorphanol (0.2–0.4 mg/kg) is advisable before major procedures (Bednarski, 2007; Armitage-Chan, 2008).
For longer procedures, intramuscular administration of ketamine (10 mg/kg) and xylazine (0.7–1.0 mg/kg) produces 20–30 minutes of anesthesia. Telazol® (6–8 mg/kg), xylazine (0.7–1 mg/kg) and butorphanol (0.2 mg/kg) produce up to one hour of anesthesia. Anticholinergics (atropine 0.04 mg/kg or glycopyrrolate 5–10 μg/kg) are often useful when using ketamine or xylazine to counteract excessive salivation or bradycardia. Supplemental administration of one-third to half of the original dose will usefully prolong the effective anesthetic period by 30–50%. Ketamine is not acceptable as a sole anesthetic in dogs, because of excessive muscle tone, salivation, and the frequent occurrence of seizures (Bednarski, 2007; Armitage-Chan, 2008).
Slow intravenous bolus injection of propofol (6–8 mg/kg) provides safe and effective anesthesia for approximately 10 minutes followed by complete recovery within approximately 30 minutes. Intermittent boluses of approximately 0.5–2.0 mg/kg or a continuous rate infusion (0.15–0.40 mg/kg/min) are suitable for prolonged procedures. Apnea is a common consequence of rapid IV injection and requires assisted ventilation until the drug wears off. Premedication with acepromazine, medetomidine or morphine will reduce the induction and maintenance doses of propofol by 30–40%. Short-acting barbiturates such as sodium thiopental (8–12 mg/kg IV) or methohexital (4–8 mg/kg IV) provide about 15 minutes of light anesthesia, but have a lower margin of safety than dissociative anesthetics or propofol (Bednarski, 2007; Armitage-Chan, 2008).
Inhalant anesthetics such as isoflurane are most appropriate for prolonged or invasive procedures in the dog. Dogs are prone to vomiting under anesthesia, and proper preoperative fasting and endotracheal intubation are vital to prevent aspiration pneumonia. Premedication with the sedative and analgesic drugs previously discussed will allay anxiety, reduce postoperative pain, and reduce the required concentration of isoflurane for maintenance anesthesia. Rapid anesthetic induction may then be accomplished through slow intravenous administration of thiopental (8–12 mg/kg IV), methohexital (4–8 mg/kg IV) or propofol (2–4 mg/kg IV) to effect. Mask induction is rarely indicated in dogs, and should only be used in premedicated animals to reduce struggling (Bednarski, 2007; Armitage-Chan, 2008).
Endotracheal intubation of dogs is easily accomplished. A variety of endotracheal tube diameters should be available to accommodate individual and breed differences in tracheal diameter. Direct visualization and intubation of the larynx of most dogs is possible without a laryngoscope. A wire stylet and a laryngoscope with an appropriate-sized Miller or Bizarri-Guiffrida blade should be available to accommodate unexpected difficulties. The animal is placed in sternal recumbency, and an assistant holds the mouth open and extends the tongue to expose the larynx. Judicious application of lidocaine spray to the vocal cords will prevent laryngospasm and facilitate intubation in small dogs. After verifying accurate placement, the cuff is inflated and the tube is secured with gauze tied over the maxilla or behind the head (Hartsfield, 2007).
Inhalant anesthesia with isoflurane may be maintained for several hours. The anesthetic concentration should be varied according to the animal’s clinical status. In general, a vaporizer setting of 1–3% will produce adequate anesthesia in dogs. Signs of adequate anesthesia include absent flexor withdrawal or corneal blink reflexes; stable heart rate and respiratory rates in response to surgical stimuli; and moderate to relaxed jaw tone. The eye will generally be central in very light or deep anesthesia and rotate ventromedially at medium anesthetic depth, but this sign is unreliable in animals which receive ketamine for induction (Muir, 2007). Paralytic agents are rarely indicated for experimental surgery in dogs. When these agents are used, the heart rate and blood pressure should be monitored and additional anesthetic administered as necessary to maintain homeostasis. Ketamine and xylazine used for induction will wear off after 30–60 minutes and require an increase in isoflurane concentration or use of supplemental analgesics to prevent sensation. The use of an “MLK” (morphine/lidocaine/ketamine) continuous rate infusion is an appropriate method to achieve balanced anesthesia at very low isoflurane vaporizer settings and maintain adequate perfusion without unnecessary prolongation of anesthetic recovery (Armitage-Chan, 2008). Isoflurane will accumulate in tissues during long procedures and multimodal analgesics often allow significant reduction of the isoflurane vaporizer settings. The vaporizer setting should generally be reduced toward the end of prolonged procedures, to avoid unnecessarily long recovery periods.
Diagnosis of pain in dogs is more reliable than in non-domestic species. Consistent signs of pain include vocalization, drooping of the head and neck (“hang-dog” expression), and site-specific behaviors (sensitivity to palpation, limping, biting or licking the incision). The Glasgow Composite Pain Scale (GCPS) is easily taught and demonstrates good interobserver reliability. A trained observer can evaluate a dog using this scale in approximately 5 minutes. Dogs receive a score of 0 (no pain) to 24 (severe pain), and should receive supplemental analgesia for a score greater than 6 (Hellyer et al., 2007).
Opioid analgesics are preferred for moderate to severe pain in dogs. The recommended dose of buprenorphine in dogs is 0.02 mg/kg IM every 8–12 hours. Fentanyl patches (2–5 μg/kg/h) are useful for postoperative pain longer than 24 hours’ duration, but must be placed at least 12 hours before surgery to achieve effective blood levels. Epidural analgesia provides excellent pain relief with fewer systemic side-effects than parenteral injection. Epidural administration of analgesics can be readily accomplished by trained staff, and provides up to 12–24 hours of analgesia following a single injection. Morphine and oxymorphone are the most commonly used epidural analgesics. The alpha-2 agonists xylazine and medetomidine are also effective by this route, but may produce significant bradycardia and hypotension (Foley et al., 2001).
The NSAIDs are highly effective analgesics in dogs, and may provide effective analgesia up to 24 hours after a single dose. Adverse effects of the NSAIDs include gastritis, which may proceed to ulceration. For this reason, NSAIDs are most appropriate for acute postoperative pain control, and not for treatment of chronic pain conditions in dogs. Recommended NSAIDs for dogs include ketoprofen, meloxicam, and carprofen.
A recent report evaluated pain following routine ovariohysterectomy in dogs receiving buprenorphine (0.02 mg/kg single preoperative dose IM), carprofen (4.0 mg/kg single preoperative dose IM) or combined preoperative buprenorphine/carprofen. All dogs were premedicated with acepromazine, induced with propofol, and maintained on isoflurane. All treatments produced significant pain relief, but 10–20% of the dogs required rescue analgesia for breakthrough pain up to 6 hours after surgery. The carprofen group demonstrated the lowest mean GCPS score at all time points. There was no significant additive analgesic effect following combined buprenorphine/carprofen treatment. Dogs which received buprenorphine or combination analgesia required a lower induction dose of propofol, and experienced a quicker anesthetic recovery than dogs which received only carprofen. All animals were essentially pain-free by 24 hours after surgery (Shih et al., 2008). The duration of analgesic treatment required following orthopedic or major abdominal or thoracic procedures commonly used in biomaterials research is likely to be longer than the 24 hours required following ovariohysterectomy, and generally requires multimodal therapy for optimal control.
Swine are commonly used in experimental surgery because of anatomic and physiologic similarity to humans in many models. Advantages to the use of pigs in the laboratory include low cost, ready availability, and ease of acclimation to the laboratory. Disadvantages include the uncooperative nature of pigs with respect to most clinical procedures, limited number of intravenous access sites, and relative difficulty of endotracheal intubation.
Because animals larger than 100 kg are difficult to handle in the laboratory, selective breeding has produced several types of miniature swine for research. The Gottingen minipigs and the Yucatan micropigs achieve a maximum bodyweight of 35–55 kg at 2 years of age. The Hanford and Yucatan minipigs are somewhat larger, and weigh 70–90 kg at 2 years of age. By contrast, adult crossbred farm pigs weigh 90–110 kg at 6 months of age, and 200–300 kg at 2 years of age (Bollen et al., 2000; Swindle, 2007). Juvenile crossbred farm pigs are less expensive than minipigs or micropigs, and are often used for short-term surgical studies. Juvenile farm pigs gain 2–4 kg/week and require larger pens as they grow, to maintain facility compliance with the Guide (National Research Council, 2010). The choice of research subject should include practical considerations such as length of study and maximum allowable body size, as well as the physiological characteristics of the different breeds (Bollen et al., 2000).
Careful evaluation of the vendor health program is required to reduce experimental morbidity from unrelated clinical conditions. Chronic respiratory disease is common in commercial swine operations. Although subclinical infections are common, transportation, anesthesia, and surgery may activate latent infections and result in excess morbidity and mortality. Causative agents in affected pigs include Mycoplasma hyopneumoniae, Haeomophilus pleuropneumoniae, Bordetella bronchiseptica, and Actinobacillus pleuorpneumoniae (Straw et al., 2006). Many commercial breeders of laboratory minipigs and micropigs maintain specific pathogen-free (SPF) herds that are free from infection with these agents. Rapid infection with respiratory pathogens occurs when SPF pigs are cohoused with conventional swine.
Malignant hyperthermia (MH) is an autosomal dominant trait that causes affected animals to develop marked hyperthermia (rectal temperature >41°C), and extensor muscle rigidity and necrosis after exposure to environmental extremes or halothane or isoflurane anesthesia. The disease has been described only in farm animals bred for rapid growth, and is becoming increasingly rare as commercial breeders identify and cull carrier pigs. Affected pigs are still encountered sporadically in farm pigs used for experimental surgery. The condition has not been described in minipigs or micropigs. MH has not been reported in other commonly used laboratory animal species. Known MH carriers should not be used for experimental surgery. Treatment of animals which develop MH during an experimental procedure requires immediate termination of anesthesia, whole body cooling, and administration of corcticosteroids, sodium bicarbonate, and dantrolene sodium (3–5 mg/kg IV) (Swindle, 2007; Smith et al., 2008).
Food and contact bedding must be removed 6–8 hours prior to general anesthesia (Swindle, 2007). Water may be offered until 2 hours before anesthesia. The presence of food in the stomach frequently results in gastric distension, hypoventilation, and tachycardia during prolonged general anesthesia, requiring prompt decompression through orogastric intubation (Thurmon and Smith, 2007).
Pigs may be easily trained to accept handling for physical examination, blood collection through a vascular access port, and dressing changes. The acclimation period following arrival to the facility before study initiation is an appropriate training time. Uncooperative pigs may require chemical restraint for minor clinical procedures. Slings and hammocks are available which will allow restraint of calm pigs up to 50 kg for up to several hours (Swindle, 2007). The Panepinto sling was specifically designed for the veterinary practice and laboratory environments, and is easy to operate and sanitize. Sedation may be necessary to facilitate initial placement of animals into the sling. Azaperone (4 mg/kg IM) is a useful agent for this purpose. For more invasive procedures, a mixture of Telazol®(4.4–6.6 mg/kg IM), xylazine (2.2 mg/kg IM), and atropine (0.05 mg/kg) provides approximately 30 minutes of anesthesia suitable for minor surgery, followed by smooth recovery. Endotracheal intubation for subsequent maintenance on isoflurane is also possible after Telazol® and xylazine induction (Bollen et al., 2000; Swindle, 2007). Alternative agents such as ketamine, ketamine/xylazine or Telazol® produce light anesthesia insufficient for surgery or intubation, and characterized by rough recovery (Ko et al., 1993, 1995).
Delayed recovery complicated by hindleg weakness may occur following Telazol®/xylazine administration in mature pigs, and is mediated by the zolazepam component of the proprietary Telazol® mixture. For this reason, many clinicians prefer to use a “cocktail” made by adding 2.5 ml of ketamine (100 mg/ml) and 2.5 ml of xylazine (100 mg/ml) to 1 vial of unreconstituted Telazol® powder. The resulting solution contains equivalent dissociative anesthetic potency to standard mixtures, with significantly less zolazepam, and is administered at a dose of 1 ml/35–75 kg IM (Thurmon and Smith, 2007).
Prolonged or invasive procedures are best conducted under isoflurane anesthesia. Endotracheal intubation is warranted for general anesthesia in swine to protect the airway and allow for controlled ventilation when required. Required equipment for endotracheal intubation of swine includes a laryngoscope with a 20–25 cm straight blade and a selection of cuffed tubes of appropriate size (4.5–8 mm). Endotracheal intubation is possible with the pig in dorsal, ventral or lateral recumbency. Two anatomic characteristics can complicate the procedure. First, the soft palate is long and must be displaced dorsally for visualization of the larynx. Second, the laryngeal diverticulum distal to the larynx may “trap” the tip of the endotracheal tube unless the tube is gently twisted as it passes over the epiglottis. After intubation, maintenance anesthesia usually requires an isoflurane vaporizer setting of 1.5%–2.5% in oxygen. The actual concentration of isoflurane will vary according to the anesthetic induction regimen, type of procedure, and concurrent use of other narcotic and sedative agents. Administration of Telazol® and xylazine for anesthetic induction has a substantial isoflurane-sparing effect that may last for the first 30–60 minutes of anesthesia. Adequate surgical anesthesia is indicated by absence of the pedal withdrawal reflex, minimal jaw tone, and stable heart rate and blood pressure (Thurmon and Smith, 2007; Smith et al., 2008).
Diagnosis of pain in swine may be challenging. Pain scoring systems reported by some investigators have not been validated in large-scale studies. Although an indirect measure, depressed appetite is the most consistent indicator of pain in postoperative pigs. In addition, pigs in pain generally exhibit impaired activity and depressed attitude compared to normal pigs.
Narcotics are commonly used postoperative analgesics in swine. Buprenorphine is the most widely used narcotic, because it provides excellent analgesia at a convenient dose interval with few side-effects. The fentanyl patch (5 μg/kg/hr) is effective in swine, and reduces the need to handle animals during the immediate postoperative period, when struggling may increase pain at the incision line or predispose to dehiscence. The 50 μg/hr patch produced therapeutic blood levels and clinically effective pain control for up to 48 hours in Yorkshire-cross white female pigs (26.2 ±2.1 kg) following experimental thoracotomy. Control pigs receiving buprenorphine (0.10 mg/kg) also experienced adequate pain control, but exhibited higher pain scores and required treatment at 4–8 hour intervals. Fentanyl patches should be applied in the dorsal mid-scapular region to prevent accidental dislodgement and ensure consistent absorption (Harvey et al., 2000). Group-housed pigs may remove and consume patches applied to cage mates and develop acute narcotic overdose.
The NSAIDs are highly effective analgesics in swine. Carprofen (2 mg/kg SQ or PO q24h), meloxicam (0.4 mg/kg SQ q24h), and flunixin (1–4 mg/kg SQ or IM q24h) have been used as sole agents or in combination with opioid local anesthetics. Gastric irritation has not been reported following short-term use, but may be a concern in animals which require chronic therapy (Thurmon and Smith, 2007; Smith et al., 2008).
Sheep and goats are desirable research animals, because the adult body weight is comparable to humans and they are hardy, inexpensive, and have a calm disposition. Sheep are particularly useful for reproductive research because investigators may easily obtain cohorts of pregnant animals with known gestational age and hysterotomy, and fetal manipulation is possible with a low postoperative abortion rate. Cardiovascular research also uses sheep and goats because the ratio of the size and weight of the heart and other thoracic organs to body weight is similar to humans (Riebold, 2007).
The quality of sheep and goats used in research often depends on the source. Surgical and transportation stress will reduce resistance to disease. Vendors should avoid mixing multiple sources of animals in one flock to ensure a consistent health profile. Similarly, research institutions should use animals from a single vendor to reduce the possibility of introducing unfamiliar infectious agents to stressed animals. Preventive health measures, including immunizations and anthelmintic treatments, should be provided at least one month before shipment. Recommended immunizations for sheep and goats include Clostridia spp., Pasteurella multocida, P. hemolytica, contagious ecthyma, and parainfluenza III. A conditioning and quarantine period after arrival allows recovery from shipping stress, acclimation to the facility, and reduces transmission of infectious diseases to resident animals (Delano et al., 2002).
Facilities should adopt measures to prevent transmission of zoonoses from sheep and goats. Silent infection with Coxiella burnetti, the causative agent of human Q fever, is common in sheep. Fetal membranes and amniotic fluid from infected animals carry large numbers of hardy organisms. Human infection occurs from direct contact with infected materials or from fomites. Signs of Q fever infection in humans range from subclinical disease to severe flu-like symptoms, pneumonia, endocarditis, and death. Serologic evaluation of sheep is difficult because animals may shed large numbers of organisms in the absence of detectable antibody. Contagious ecthyma (“orf”) is a poxvirus-induced papular disease of sheep and goats. Infected animals frequently have lesions on the mucocutaneous junctions of the head, which can spread to humans by direct contact. Affected animals and humans develop long-lasting immunity and usually recover in 10–14 days. Effective vaccines for sheep and goats are available (National Research Council, 1997).
Careful preoperative preparation of the rumen is required for safe anesthesia of sheep and goats. Adult animals require withdrawal of food and water for at least 12–24 hours before surgery to reduce rumen size and digestive activity. Rumen distension and hypoventilation are common when dorsal or lateral positioning of ruminants is required for anesthesia. Preoperative passage of a 1–2 cm diameter thick walled stomach tube into the rumen allows intraoperative aspiration of gas and fluid if required. Placement of a cuffed endotracheal tube should immediately follow anesthetic induction to prevent aspiration pneumonia. Aspiration pneumonia can be fatal after regurgitation because rumen fluid contains numerous anaerobic bacteria. Infant ruminants (<30 days of age) lack a functional rumen and do not require preoperative fasting (Riebold, 2007).
Sheep and goats are docile and rarely require sedation for blood sampling, dressing changes or other routine procedures. For more invasive procedures, animals may be sedated for 10–20 minutes with a low dose of xylazine (0.02–0.15 mg/kg IV or 0.05–0.3 mg/kg IM). Yohimbine (1 mg/kg IV) or atipamezole (0.02–0.06 mg/kg IV) (Muir, 2007) reverse xylazine-induced sedation when necessary. Research staff should be mindful of the much lower xylazine dose required for ruminants compared to other species. By contrast, the dose of medetomidine is similar in most laboratory animal species (30 μg/kg). Animals in late pregnancy should not receive xylazine to avoid fetal hypo-oxygenation secondary to depressed maternal cardiac output. The benzodiazepines diazepam (0.25 mg/kg IV) and midazolam (1.3 mg/kg IV) produce muscle relaxation and sedation in sheep and goats, with less depression of cardiac output than is seen with xylazine. Flumazenil (1 mg IV) reverses benzodiazepine-induced sedation. Ataxia and excitement are common following benzodiazepine administration (Riebold, 2007).
Isoflurane is the anesthetic of choice for general anesthesia of sheep and goats. Intravenous administration of ketamine (2.75 mg/kg) and diazepam (0.2 mg/kg) or xylazine (0.1 mg/kg) rapidly induces light anesthesia of 10–20 minutes duration suitable for endotracheal intubation. Propofol (4.0–6.0 mg/kg IV to effect) is a suitable alternative for brief procedures or to facilitate intubation and maintenance on isoflurane for longer procedures (Riebold, 2007). In a study comparing propofol to ketamine/xylazine/halothane for subcutaneous biomaterial implantation, propofol recipients exhibited better cardiovascular and respiratory parameters than ketamine/xylazine/halothane recipients. These animals also demonstrated a more rapid and smoother anesthetic recovery (Lin et al., 1997). Thiopental (25 mg/kg IV) is an alternative induction agent; however, significant disadvantages include regurgitation, profuse salivation, and irritation from perivascular infiltration (Dorr and Weber-Frisch, 1999). Profuse salivation follows the administration of most anesthetic drugs in ruminants. Administration of anticholinergic drugs is not recommended in ruminants, because these agents may increase salivary viscosity and impair gastrointestinal motility. Anesthetized ruminants should be positioned with the head down to encourage drainage of saliva away from the airway (Riebold, 2007; Thurmon and Smith, 2007).
Endotracheal intubation requires a laryngoscope with a 20–30 cm blade. Vinyl or silicone cuffed tubes 10–16 mm in diameter are suitable for most sheep and goats. An assistant positions the animal in sternal recumbency, extends the neck, and holds the mouth open. The anesthetist visualizes the epiglottis with the laryngoscope and intubates the trachea during inspiration. Prior application of 2% lidocaine spray to the vocal cords prevents laryngospasm. The use of a stylet may help deflect the tip of the tube into the larynx. A mouth gag is often necessary to prevent damage to the tube by sharp molar teeth. After inflation of the cuff, the tube is secured with gauze to the mandible. Isoflurane vaporizer settings of 0.75–1.0% are often sufficient to maintain general anesthesia in sheep and goats. Absence of chewing motions in response to stimulation indicates an adequate surgical plane. The presence of a centrally positioned eyeball with a dilated pupil and absent palpebral reflex indicates very deep anesthesia (Riebold, 2007).
The NSAIDs are highly effective in small ruminants, and are recommended for first-line treatment unless scientifically contraindicated. Flunixin (1.1–2.2 mg/kg), ketoprofen (3.3 mg/kg), and carprofen (0.7 mg/kg) provide effective analgesia for up to 24 hours after dosing. When used at a dose of 4.0 mg/kg, carprofen provides therapeutic blood levels for at least 72 hours (Thurmon and Smith, 2007). All NSAIDs should be administered parenterally in small ruminants. The intravenous route is preferred for animals with severe acute pain, but the subcutaneous and intramuscular routes have been widely reported.
Opioid analgesics may be indicated as a supplement or as primary analgesics in some cases. The epidural space is easily accessed in small ruminants. Epidural morphine (0.1 mg/kg diluted in sterile saline) administered in the lumbosacral or sacrocaudal spaces provides up to 12 hours of analgesia (Riebold, 2007). The transdermal fentanyl patch (50 μg/hr) produces effective blood levels in sheep and goats for up to 72 hours. The dose rate is lower than otherwise expected for an animal this size, because of significant recirculation from the rumen to the bloodstream, bypassing hepatic detoxification pathways, and animals should be observed for excessive sedation or excitement (Valverde and Doherty, 2008).
A recent report demonstrated the superiority of transdermal fentanyl over parenteral buprenorphine in a multimodal analgesic regimen following experimental tibial osteotomy and locking compression plate placement in mature Polypay-cross ewes. The fentanyl group received transdermal fentanyl patches (2 μg/kg/hr) on the lateral antebrachium 12 hours before surgery. Control animals received buprenorphine (0.01 mg/kg every 6 hours beginning at anesthetic induction). All animals received the NSAID phenylbutazone (2.2 mg/kg IV every 24 hours) for 3 days beginning at anesthetic induction. Blinded observers evaluated the animals for pain every 12 hours for 72 hours, including a final observation 12 hours after patch removal or the last dose of buprenorphine. Fentanyl recipients required a lower dose of diazepam during anesthetic induction, and exhibited lower pain scores than buprenorphine recipients throughout the observation period, although no animals required rescue analgesia. The buprenorphine group exhibited an increased pain score 12 hours after the last dose of analgesic, suggesting possible rebound hyperalgesia (Ahern et al., 2009).
Ruminants are more sensitive to systemic effects of local anesthetics than other species. The mean fatal dose of lidocaine in sheep is 30.8 mg/kg, compared to 80 mg/kg in dogs, and seizures may occur following administration of 10 mg/kg. The maximum suggested dose of lidocaine for sheep and goats is 0.5 ml/kg of a 2% solution. For wide infiltration, 2% lidocaine may be diluted to 1%. Signs of systemic local anesthetic overdose include seizures, respiratory depression, bradycardia, hypotension, and collapse.
Animal models are an invaluable resource in biomaterials research. Surgery is often required in biomaterials models. Animal welfare regulations in most countries require scientists to use animal models only when necessary, and use the minimum number of animals consistent with good experimental design. All necessary steps must be used to prevent and treat pain or distress associated with experimental manipulation.
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