Anatomic studies are used to evaluate the nervous system by examining the cellular organization or chemical composition of specific brain regions or by examining how different brain regions are related by afferent or efferent connections. These studies most commonly involve either the use of tracer substances to label and visualize neural pathways or the use of lesion techniques to destroy a discrete area of brain cells and examine the course of degenerating fibers. Electrolytic and radio frequency techniques, as well as those using neurotoxins, can be used to make brain lesions. Stereotaxic approaches are often used to make more focal lesions or lesions in deeper brain structures. When tracers are used, they are injected into the nervous tissue, where they are incorporated into neuronal cell bodies and/or processes and then transported anterograde or retrograde. Transport of tracers and degeneration of fiber pathways generally occur over a period of several days after injection or a lesion; therefore, an animal must be allowed to survive for a short period before being sacrificed for study of its nervous system. The possibility of using labeled substances, such as manganese, in combination with brain imaging to trace anatomic connections is also developing (Saleem et al., 2002); the same animal can be examined repeatedly with this technology, so it reduces the number of animals needed for a particular study (see “Imaging Studies,” below).
Various factors will determine whether and to what extent the IACUC and the investigator need to exercise flexibility in interpreting and implementing the recommendations of the Guide. Those factors include the invasiveness of the
procedures, the surgical setting, whether multiple injections are necessary, and the characteristics of the injected substance. This brief discussion is derived from page 12 of the NIH workshop report, Preparation and Maintenance of Higher Mammals During Neuroscience Experiments (NIH, 1991).
Grading the Invasiveness of the Procedure(s)
The invasiveness of the procedure required to inject a tracer will establish whether it constitutes a major survival surgery, which should be performed in a facility intended for that purpose. Thus, while an injection into the eye is a relatively minor surgical procedure similar to a biopsy, an injection into a central brain structure, which usually requires performing both a craniotomy and a durotomy, is usually considered a major surgical procedure (NIH, 1991).
For further discussion of major versus minor surgery, see “Surgery and Procedures” in Chapter 3.
Modified Surgical Settings
Anatomic studies are often performed in modified surgical settings (for a discussion of the characteristics of a modified surgical setting, see “Surgery and Procedures” in Chapter 3). The reason for allowing an exception to the Guide’s recommendations about performing major survival surgery in a dedicated surgical facility is that it may not be possible to sterilize the necessary experimental equipment (amplifiers, oscilloscopes, audio monitors, micropipette pullers, pressure microinjection devices, and micromanipulators) or move it into the dedicated surgical facility (NIH, 1991). Other factors that may influence an IACUC’s decision to approve a modified surgical setting are (NIH, 1991):
the use of a radioactive tracer substance;
the need to manufacture, fill, and position into the brain several injection micropipettes during the course of a single procedure;
the relatively short duration of the post-surgical survival period (2–4 days);
the suitability of the proposed laboratory area for aseptic surgery;
the infrequency of the procedure (less than once per month, on average);
verification of the absence of post-surgical infection or other complications in a series of animals from a pilot project.
The scientific requirements of certain experiments may require subjecting a single animal to multiple injection procedures. As a result, when the injection site is a central brain structure, it will be necessary to subject an animal to multiple major survival surgeries. Experiments of this sort include those in which two or more tracers are to be injected and it is known that they require
markedly different survival periods for transport to occur. Another example is experiments in which injections will be made at different ages in the same animal in order to label the arrangement of connections at different stages in the development of a neural pathway (NIH, 1991).
Each of those situations would require an IACUC exception to guidelines and regulations on multiple major survival surgeries. In both examples, the surgeries are “related components of a research project” (NRC, 1996), and IACUCs may choose to grant approval for these multiple major survival surgeries.
Standard aseptic technique is used to expose the surface of the occipital cerebral cortex. At this point the animal often is redraped and the surgeon breaks sterility in order to fill and insert the injection micropipettes into the micromanipulator, position the pipettes into the brain, and adjust and activate the pressure injection device. Whether or not the micropipettes are able to be sterilized prior to surgery will depend on several factors, including the material out of which the pipette is made and the substance being injected. . . . Throughout this period, the surgeon has no direct contact with the wound site or the surgical field. When the injections have been made, the surgeon re-gloves (and re-gowns, if necessary), the top level of drapes is removed, and the wound is closed using standard aseptic technique (NIH, 1991).
The topic of micropipette sterility is similar to the discussion of implant sterility in the succeeding section (“Asepsis and the Introduction of Devices or Implants into Neural Tissue”).
Characteristics of the Injected Substances
Another important consideration is the chemical properties of the substance to be injected. For instance, many tracers are sensitive to temperature and cannot be heat sterilized prior to injection. Others may exhibit high levels of tissue toxicity and can cause a marked local inflammatory response at the injection site. Some neural tracers are radioactive and proposed procedures for their use must be reviewed for compliance with institutional policy on radiation safety (NIH, 1991).
Neurophysiology Experiments in Awake, Behaving Animals
Neurophysiology experiments in awake, behaving animals have fundamentally shaped our understanding of the processing of information throughout the brain because they provide the most direct information about neural signals.
Studies linking neurophysiology and brain imaging are useful for clarifying the meaning of the brain activation seen during functional-imaging experiments. The behavioral repertoires of many mammals resemble those of humans, and data generated using awake, behaving animals can have considerable relevance when extrapolated to humans. Awake, behaving animals make it possible to study the “higher” functions of the brain by involving the active participation of the animal. Trained animals not only can serve as subjects in experiments on motor control, but also can be sophisticated participants in psychophysical studies of the processes involved in perception and memory (NIH, 1991). In addition, the results of brain-recording experiments have probably had more influence on the fields cognate to neuroscience than any other kind of experiment; their influence is evident in fields as disparate as behavioral psychology, image-processing, and computer design.
Experiments on an awake, behaving animal generally involve a long initial phase during which the animal is trained to perform a task. Once the animal is trained, experimental sessions are held several days a week for months and sometimes years, during which specific physiologic variables are measured. For example, a nonhuman primate may be trained to look at a particular object on cue. Then, during subsequent experimental sessions, the researcher measures the neural activity associated with the eye movement. Monitoring a physiologic process, whether it is the experimental variable (the electrical activity in the brain) or the acquisition of the behavioral task (the eye movement), frequently necessitates the implantation of various devices. In the previous example, eye coils may be implanted to monitor eye position and microelectrodes may be placed in the brain to measure neural activity.
Neurophysiology experiments on an awake, behaving animal frequently require that the animal be confined to a restricted working space for many reasons, such as to enable precise positioning of recording or stimulating electrodes into the correct region, to stabilize the spine or head for neurophysiologic recording, to maintain the animal’s posture in relation to the behavioral task, to maintain the orientation of the animal relative to a sensory stimulus, to restrict the range of reach of an animal to prevent it from accidentally damaging implanted equipment that is exposed during recording sessions (e.g., electrodes or lead wires), or to prevent movement that would induce errors or variation in the experimental data (Lemon, 1984c). Restraint of an awake, behaving animal often involves transferring the animal to a special apparatus, such as a restraining chair or an operant chamber. There is a long tradition of studying the neurophysiology and behavior of rodents in various kinds of mazes (including water mazes), running wheels, or open-field areas (Porsolt et al., 1993). Depending on the experiment, the apparatus into which an animal is placed may or may not be inside a larger chamber that is designed to attenuate extraneous visual or auditory stimuli during the experimental session (Ator, 1991). Whatever specialized chamber is used, the animal remains in it for the duration of the experimental session.
Some types of neurophysiology experiments require that a probe be placed in the brain only during the actual experimental sessions, either to stimulate the brain (stimulating electrode), to record electrical activity (recording electrode), or to sample the fluid in the interstitial space (microdialysis probe). Owing to their fragility, or the need to reposition them to sample from a different area of the brain, these probes are removed at the end of each recording session. The use of these types of devices requires the implantation of chronically indwelling hardware called guide cannulae or chambers. When these are implanted, a piece of the skull is removed (craniotomy) and the hardware is placed over the hole and attached to the skull. The hardware is hollow, allowing free access to the brain, and is filled with a sterile solution (typically saline) and capped with a sterile cover to prevent introduction of microorganisms. When the investigator needs to place a probe to begin an experimental session, the cover to the guide cannula or chamber is removed and the probe is introduced into the brain.
Major surgery to implant hardware devices for head restraint, data collection, and stimulation can be accomplished with standard aseptic surgical techniques and typically can be performed in a facility dedicated to aseptic surgery (see Gardiner and Toth, 1999, and Lemon, 1984b, for discussions of surgical issues related to cranial implants). When implanting guide cannulae or chambers, the size of the craniotomy should be large enough to allow access to the structure being studied without unnecessarily exposing neural tissues (Lemon, 1984a). If an implanted device is necessary during the training of the animal, the animal should be conditioned to the training environment prior to any surgery. In this way, animals that will not accept training can be removed from the study before they are subjected to an unnecessary surgery.
Neurophysiology Experiments in Anesthetized Animals
Cases where recordings are made while an animal is anesthetized raise critical questions regarding anesthesia, maintenance of physiologic status, and monitoring of the animal’s condition. The choice of anesthetic must satisfy the need of the experimenter to perturb neuronal status as little as possible while ensuring that the animal remains free of pain and distress. Maintaining an anesthetized (and sometimes immobilized) animal in appropriate physiologic condition is a considerable technical challenge (see “Prolonged Nonsurvival Studies” in Chapter 5). Monitoring both the anesthesia and the animal’s general condition requires careful attention to a number of measures. Although animals in some studies are used in repeated experiments (with intervening recovery periods), in other cases they are maintained under anesthesia for long periods of time for nonsurvival studies (see Chapter 5). If there will be repeated sessions of prolonged anesthesia, special attention should be paid to maintaining the animal’s normal physiologic status between anesthetic sessions.
Animal Care and Use Concerns Associated with Neurophysiology Experiments in Awake, Behaving Animals
Many of the animal care and use concerns associated with recording in awake, behaving animals were discussed in Chapter 2 and 3. They include the use of food and fluid regulation, monitoring animals for signs of pain or distress, and the general use of restraint. Additional animal welfare issues specific to or frequently encountered in neurophysiology experiments include head restraint systems, chairing nonhuman primates, multiple survival surgeries, modified surgical settings, asepsis during introduction of probes into the brain, monitoring the site surrounding implanted devices or hardware for signs of infection, dealing with rejected or failed implants, maintaining chambers free of infection, and periodic durotomy.
Restraint During Neurophysiology Experiments in Awake, Behaving Animals
Most experiments that involve the monitoring of neural activity require some limitation of the animal’s working space and/or freedom of body movement (NIH, 1991). Many forms of restraint are acceptable as long as the particular procedures for accomplishing and monitoring restraint are well justified and consistent with the Guide, and the period of restraint is as short as possible. Animals that are restrained must be monitored closely to ensure that the restraint method permits reasonable postural adjustment, does not interfere with respiration and does not cause skin abrasions or bruising. If ulceration or bruising develops, the animal should be removed from the study until the injured area is fully healed, and adjustments should be made to correct the source of the problem.
In some neurophysiology studies, the restraint is the independent variable in an experiment (for example, to study the physiological responses believed to be affected by unfamiliar restraint). However, in most cases, the restraint is not a variable in the experiment, and a training phase is carried out to habituate the animal to the restraint before the experiment begins. Because animals in behavioral experiments are handled frequently (often 5 or even 7 days a week), they usually become habituated to the head restraint, tether, or chairing quickly. The best evidence of behavioral adaptation to restraint is voluntary movement into the device (NIH, 2002) and performance of the behavioral task once there.
In experiments in which animals are tethered to treadmills or other devices used to study locomotor behaviors, care should be taken to ensure that they cannot become trapped in the apparatus (see “ Exercise” in Chapter 8). That may require intensive and continuous monitoring of animals during training and recording sessions. As mentioned before, appropriate and thorough habituation to the apparatus before experiments begin can substantially reduce the risk of entrapment and the distress that could arise with the use of the device.
In some types of neurophysiology experiments, the animal’s activity may be restrained with a tether. For example, in experiments involving intravenous drug self-injection or intragastric drug delivery (Lukas et al., 1982) the animal may have a chronic indwelling intravenous or intragastric catheter that (Lukas and Moreton, 1979; Meisch and Lemaire, 1993) exits from a site on the back (typical in monkeys) or the top of the head (typical in rats and cats), and is threaded through a protective tether that is connected to a swivel. The tubing emerges from the swivel and is connected to a pump, which is used to deliver the drug. Monkeys that have been fitted with chronic indwelling catheters often wear specially designed vests, shirts, or harnesses to protect the catheter exit site. Habituation of an animal to a harness-tether arrangement is best carried out well in advance of the planned date of implantation of the catheter. Inspection of the animal during the habituation process allows the experimenter to determine whether the vest or tether fits well and permits adjustments to prevent discomfort.
Head-restraint systems minimize the movement of the head during neurophysiology experiments without causing discomfort if the animal is properly conditioned (NIH, 2002). Hardware, generically called a head-holder, is implanted chronically on the animal’s skull. Three different styles of head-holders are generally used: implantable, halo, and headpiece (Lemon, 1984a). Small screws or bolts and dental acrylic or bone cement anchors the head-holder to the skull. Then, during a training or experimental session, the head-holder is attached to a freestanding platform to immobilize the head. Besides minimizing movement, these systems provide a structural element to which to anchor connectors from other surgically implanted monitoring devices, such as eye coil wires, chronically implanted recording electrodes, or indwelling cannulae for delivery of pharmacological agents. They also can provide a superstructure through which microelectrodes are introduced into the brain for the recording of neural activity. Animals should be properly conditioned to the restraint to eliminate any discomfort or stress that might be associated with it (see “Physical Restraint” in Chapter 3).
Chairing Nonhuman Primates
Macaques and squirrel monkeys can be trained to move voluntarily from the home cage into a restraint chair (Ator, 1991). Commonly, nonhuman primates wear either a collar with a small metal ring attached or a collar with a slot to which the pole directly attaches. The monkeys acclimate to having a chain clipped to their collar; the chain is then pulled through a ring at the top of a metal pole.
Squirrel monkeys usually grasp the pole and ride to the chair on it, while larger monkeys, such as adult macaques, learn to walk to the chair. By holding one end of the pole snugly at the collar and pulling the chain down to the other end, the experimenter can control the monkey’s movements and proximity and thus be protected from the possibility of a bite in the process of training and transfer. Larger monkeys can be trained to move from the home cage into a smaller shuttle device that can be wheeled to the experimental chamber. Treats may be used during the various steps of training the monkey to cooperate in the transfer process and sitting in a chair. While the amount of time that an animal is chaired can be gradually extended during the training process, the animal should not live in the chair. In instances where long-term (greater than 12 hours) restraint is required, the nonhuman primate must be provided the opportunity daily for unrestrained activity for at least one continuous hour during the period of restraint, unless continuous restraint is justified for scientific reasons and approved by the IACUC (AWR 3.81 (d)).
Multiple Survival Surgeries
In many experiments using awake, behaving animals, the implantation and maintenance of recording devices, head restraint devices, and stimulation devices necessitates multiple major survival surgeries. The use of multiple surgeries in these experiments, including surgeries to repair implants, is permitted by the Guide because they are related components of a research project, they will conserve scarce animal resources, or they are needed for clinical reasons.
The need for multiple major surgeries may arise for several reasons. In some cases, it arises for clinical reasons. For example, a head restraint device and a chamber may need to be implanted in close proximity. Implanting them during one long surgery could undermine the structural integrity of the skull, create an extremely large wound, and increase the risk of infection. Multiple major surgeries may also be necessary due to limitations of the experimental devices used. For instance, eye coils typically will function reliably for a limited period of time after implantation. If a prolonged training period with head restraint is necessary before experimental sessions can begin to monitor eye position using eye coils, implanting the head restraint device and the eye coils during one surgery prior to the start of training could mean the eye coils will not function reliably during the subsequent experimental sessions. In this case, performing a second major survival surgery is necessary and justified to ensure that the eye coils will function reliably during the experimental session.
Performing multiple major surgeries may also be the best surgical approach if doing so allows a major surgery to be performed in an aseptic surgical suite, rather than in a modified surgical setting. Often, probes must be positioned precisely in the brain of an awake, behaving animal by means of a head restraint device or recording chamber that is implanted on the skull in stereotaxic coordi-
nates. However, it may be impossible to move all the equipment necessary both to implant the device or chamber on the skull stereotaxically and to monitor the output of the neurophysiological probes into a suite dedicated to aseptic surgery. Rather than performing a single long surgery to implant the head restraint device and then position the implanted probes in a modified surgical setting, performing multiple surgeries may be preferable. In this way, the head restraint device can be implanted in the aseptic surgical suite and the animal can recover from the surgery and heal. Then a second smaller surgery to place the electrodes could be performed in the modified surgical setting. This minimizes the potential for infections and subjects the animal to two short surgeries rather than one prolonged surgery.
Multiple major surgeries may also be required to maintain the viability of implanted devices. Though all percutaneous implanted devices are designed so that the skin can heal around them and the devices can be used without causing the animal pain or distress, it may be necessary to replace electrodes or eye coils that no longer function or to replace implanted hardware that has failed or been rejected.
In all of these cases, PIs, veterinarians, and IACUCs must work together to balance the animal’s well-being and the scientific goals of the experiment. Consideration of such factors as the use of scarce or conserved species and the disposition of individual animals (especially the case with higher mammals such as nonhuman primates) will influence the decision of how many survival surgeries are acceptable.
There is no need to treat all procedures for the clinical management of an implanted animal as major survival surgeries that must be performed in a facility dedicated to aseptic surgery. Procedures, such as treating surgical wounds as they heal, cleaning and maintaining implanted devices, and removing the granulation tissue that typically forms over the dura mater inside chronically implanted recording chambers (Lemon, 1984a; Toth and Gardiner, 1999), are commonly performed under light anesthesia in a laboratory setting, using aseptic techniques within a local sterile field (NIH, 1991). Classifying those procedures as major or minor surgeries according to regulatory guidelines is not straightforward (see “Surgery and Procedures” in Chapter 3), demanding that professional judgment, guided by outcome or performance-based consideration, be employed. Certainly, the long tenure of these animals in the research setting and the many hours devoted to their training militates in favor of exercising maximum precautions to avoid infection. However, the majority of these procedures are brief and innocuous, with minimal risk of infection, and the investigator, veterinarian, and IACUC should use professional judgment to balance the well-being of the animal with the practicality of performing the procedure in a facility dedicated to aseptic surgery.
Modified Surgical Settings
Sometimes, it is necessary to implant recording or stimulating devices using neurophysiologic responses to identify the correct location in the brain. This
surgery should be performed in a facility dedicated to aseptic surgery whenever possible. However, if the procedure requires specialized equipment that cannot be sterilized or moved into a dedicated surgical facility, then all or a portion of the surgery may be performed in an approved modified laboratory setting. In these cases, the surgery sometimes can be performed in two steps. The first step— which often entails the implantation of the hardware for head restraint and opening of the skull—is performed in a dedicated aseptic surgical facility. A temporary cap is placed over the opening of the skull. On the following day, the animal is taken to the laboratory and restrained with the hardware that has been implanted for head restraint, and the temporary cap is removed. A microelectrode, micropipette, or microdialysis probe may then be implanted into the brain, maintaining asepsis in the area immediately around the site and using specialized equipment to position the device accurately. If the two-step approach is not feasible and if the laboratory can be sanitized and prepared to allow aseptic technique, the entire procedure may be performed as a single survival procedure in the laboratory (for more discussion of this subject see “Surgery and Procedures” in Chapter 3).
Animal Care and Use Concerns Associated with Introduction of Probes into Neural Tissue
Questions about sterility arise when considering the implantation of probes, such as microelectrode, micropipette, and microdialysis devices, into neural tissue. Most implanted probes can be sterilized, but this may not always be the case for sensitive or delicate probes such as microelectrodes or micropipettes, as there is no consensus on whether they can be sterilized without degrading their performance. Many laboratories do not sterilize microelectrodes and micropipettes because of their fragility, and this practice does not seem to introduce infections into the brain. Currently, there is no published, systematic evidence that the use of micropipettes or microelectrodes that have not undergone rigorous sterilization before implantation has a deleterious outcome on experiments, on the brain, or on animal health. This could be because the materials and fabrication methods used to produce microelectrodes and micropipettes may result in their being relatively free of microorganisms without additional intervention. With this in mind, any material that will be inserted into or implanted in the brain should always be handled and stored with care to protect against contaminants. The above notwithstanding, whenever possible, probes should be sterilized or alternatively disinfected before they are inserted into neural tissue.
The success with which a probe can be sterilized or disinfected immediately before its use will depend upon several factors, including the materials out of which it is made. Existing options for sterilization include heat or gas methods, soaking in bactericidal solutions, and irradiation with ultraviolet light (Lemon, 1984a). In many cases, the materials used to manufacture probes may not withstand those
rigorous sterilization procedures. In such situations, a method of disinfection should be used if possible, such as soaking in povidone iodine, chlorohexadine, or aqueous alcohols and then rinsing with sterile saline prior to insertion. If none of those options preserves the viability of the probe, attention to maintaining its cleanliness during handling and storage becomes even more important.
Investigators, veterinarians, and IACUCs should monitor for deleterious effects caused by nonsterile probes by developing performance-based standards for the histopathologic analysis of postmortem tissue specimens. As new methods become available to sterilize microelectrodes and micropipettes without compromising their utility, such as vaporized hydrogen peroxide, they should be implemented.
In some types of neurophysiologic experiments, probes such as microelectrodes, micropipettes, or microdialysis probes are introduced into the brain through guide cannulae or chambers at the beginning of the daily experimental session and removed at the end of the session. These types of probes are usually introduced without anesthesia, and their introduction typically does not require they be performed in a dedicated surgical facility, though aseptic technique when handling and inserting the probes is necessary to prevent infection. The brain itself lacks sensory endings, so the passage of these probes gives rise to no sensation. The dura mater does contain nociceptive fibers, primarily adjacent to large blood vessels (e.g. the middle meningeal artery) (Baker et al., 1999; Wolff, 1963); however, the insertion of probes through the dura mater usually evokes no reaction from an animal. On occasion though, an indication of momentary or minor discomfort may be noted. The U.S Government Principles state that “procedures that may cause more than (emphasis added) momentary or slight pain or distress should be performed with appropriate sedation, analgesia, or anesthesia.” Accordingly, in most instances, the placement of probes in awake, behaving animals may be performed safely and humanely without sedation, analgesia, or anesthesia.
Potential adverse consequences of insertion of probes into neural tissue are infection or brain injury as a result of cerebral edema or hemorrhage. The likelihood of those deleterious effects is affected by the frequency of probe insertion, the location of the probe insertion site, the depth of penetration, the physical characteristics of the probe, the expected duration of experimental sessions, and the course of the experiment for each animal. Training laboratory personnel in identifying adverse reactions and fostering a team approach that includes veterinarians and husbandry staff will help to ensure the well-being of animals used in these types of studies.
Monitoring the Site Surrounding an Implanted Device
Sites surrounding implanted devices or hardware, such as chambers, head-restraint devices, eye coils, nerve cuffs, electromyography (EMG) electrodes,
etc., should be examined regularly for signs of irritation, infection, or device damage. Specifically, investigators and animal-care staff should watch for signs of inflammation or infection of the eye coils, along the subcutaneous length of eye coil leads, and near the sites where wires, chambers or other hardware devices are externalized. Similarly, the attachments for nerve cuffs around nerves or of EMG electrodes onto muscles should be closely monitored for signs of inflammation or infection. Leads from these types of implants often are externalized to connectors that are attached to the skin or bone. These connectors should be positioned so that they are not easily manipulated or broken by the animal. Implant protection may also necessitate the use of connector hoods or fitted jackets for the animals to protect the externalized wires or connectors. Like eye coil leads, the wires from other devices should be examined throughout their subcutaneous lengths and at the skin margins for any signs of inflammation or infection.
Unambiguous experimental endpoints should be established before any devices or hardware are implanted. These endpoints should indicate when devices or hardware should be removed because of failure, infection, or inflammation. Successful reimplantation after implant failure may be possible in some circumstances. Therefore, the necessary conditions for reimplantation of previously used or replacement hardware should be described in the animal-use protocol and approved by the IACUC. Anticipating the potential consequences of implant failure before its occurrence is crucial for the viability of the study and animal well-being. A team approach involving veterinary staff, caretakers, neuroscientists, and technicians is critical to the long-term success of experiments that use animals with chronic implants.
Occupational Health and Safety
It is prudent to reiterate that risks, such as exposure to B virus, are associated with working with awake, behaving nonhuman primates (see “Experimental Hazards” in Chapter 2). Investigators, their laboratory personnel, veterinarians, and veterinary-care support staff should all be aware of the resources that provide information about appropriate precautions in these types of experimental settings. Investigators should make certain that their research personnel are fully trained in the proper handling, husbandry, and maintenance of nonhuman primates and, if necessary, in the disposal of devices and other materials that have been in contact with their tissues or fluids. To minimize the risk of personnel exposure to biologic agents or puncture, used probes should be disposed of in approved biological hazard sharps containers.
Developments in imaging technologies have led to groundbreaking advances in our understanding of neural and physiologic functions in normal and diseased humans and animals by offering a view of the living brain at work (Hoehn et al., 2001). The technologies are generally less invasive than other investigative scientific methods and offer an opportunity to address questions of structure and function without significant consequences for the research subjects (Balaban and Hampshire, 2001).
Several imaging techniques are used in animals. They include positron-emission tomography (PET), single-photon emission computed tomography (SPECT), magnetic resonance imaging (MRI) and functional MRI (fMRI), nuclear magnetic resonance imaging or spectroscopy (NMR), near-infrared spectroscopy, ultrasonography, computed tomography (CT) and optical imaging (Balaban and Hampshire, 2001; Hoehn et al., 2001; Rolfe, 2000). Some of the techniques, such as PET and SPECT, enable measurement of blood flow, oxygen and glucose metabolism, receptor density, or drug concentrations in regions of the living brain (Mathias, 1996). Others, such as MRI and NMR, provide imaging of superficial and deep brain structures with a high degree of anatomic detail. High-field MRI, SPECT, and PET techniques can also be used to provide in vivo longitudinal evaluation of receptor binding and gene expression following gene therapy (Auricchio et al., 2003; Kasper et al., 2002).
Each of those techniques allows researchers to test hypotheses about the functions of different regions of the brain on the basis of functional composition or physiologic activity. The hypotheses can often be explored further with human subjects performing specific tasks during PET, SPECT, or fMRI. However, many of the technologies provide even better resolution when used in small mammals, providing more information about physiologic function than can be obtained with human subjects (Balaban and Hampshire, 2001). Animal models enable variables associated with specific diseases to be manipulated and controlled to a degree that is not possible with human patients. Furthermore, individual animals can be evaluated repeatedly during the course of a disease or can serve as their own control instead of sacrificing large groups of animals at different time points, and thereby reducing the number of animals used (Hoehn et al., 2001).
Animal Preparation and Maintenance During Imaging Studies
Imaging generally requires anesthesia so that the animal remains motionless throughout the duration of image collection. The exception is ultrasonographic images, which can be collected from a restrained nonanesthetized animal, pro-
vided that the process does not create substantial stress in the animal. Conditioning animals to the type of handling associated with the scans obviates anesthesia. In fact, PET and fMRI scanning has been conducted on conscious monkeys that have been trained to sit in a chair (Stefanacci et al., 1998; Tsukada et al., 2000). However, it can take much time and effort to train the animals (Tsukada et al., 2000).
Generally, animals are sedated or anesthetized and then intubated either before or after transportation to the imaging facility. Because imaging facilities are rarely close to the vivarium, the methods by which animals will be transported to and from the imaging site must be considered when animal-use protocols involving these techniques are being prepared. Special attention must be given to the unusual occupational health and safety risks associated with transportation, including exposure of the transportation route or the imaging facilities to animal tissues or fluids; training and supervision of research and imaging personnel; and development of procedures for dealing with emergencies that arise during imaging or transport (such as bites and scratches). Furthermore, the personnel and methods used to monitor the animals and to administer appropriate care to ensure their well-being during imaging should be identified. Often, animals are imaged after normal business hours using facilities primarily dedicated to humans (such as at hospitals). The imaging facility professional staff may not be onsite after business hours to assist if there is a problem with the equipment, so identifying a member of the professional staff to contact in the event of an emergency may be necessary.
Special Considerations of Animal Maintenance in the Imaging Environment
Some of the features of imaging machines that make them powerful tools create an environment that may be inhospitable to routine maintenance of anesthesia and monitoring of animals. For instance, the strong magnetic field associated with an MRI machine may damage ferromagnetic components in monitoring devices or traditional ventilators (Chatham and Blackband, 2001; Kanal et al., 2002) and indeed may actually attract ferromagnetic devices or standard surgical equipment to the magnetic-field coil. This can result in injury to personnel assisting in scanning or to experimental animals, and may damage the monitoring device and scanner (Chatham and Blackband, 2001). Before an animal is imaged with any device that creates a strong magnetic field, the research staff must ascertain that the animal does not have any ferrous implants. A variety of implants, made of nonferrous materials, are available and are suitable for use with imaging equipment.
Monitoring equipment that is compatible with the imaging equipment is available at imaging facilities and may be appropriate for monitoring animals. MRI-compatible physiologic monitoring capacity includes heart and respiratory
rate, pulse oximetry, and temperature. Identifying the types of monitoring equipment that are available at the scanning facility and ensuring that it can be used with animal subjects are important considerations for these types of protocols. The ability to monitor the physiologic status of an animal during scanning is extremely valuable because direct observation and access to the animal may be reduced during image acquisition.
Maintaining an animal’s body temperature during transportation to and from the imaging facility and during scanning improves the maintenance of anesthesia. Warming blankets often have metallic components or require a power source, so the use of portable, nonmetallic warming devices is advisable. These devices produce heat as a result of a chemical reaction or after microwaving. Covering the animal with blankets and using one of these warming devices is an effective way to maintain a favorable body temperature during relatively short imaging sessions.
As many imaging facilities are utilized both for human and animal scanning, the potential for cross-contamination exists. Human B virus exposure is always a concern when macaques are involved (Cohen et al., 2002) and human allergies to rodents, dogs, and cats are common (Wolfle and Bush, 2001). In addition, some animals may be susceptible to zoonotic diseases from humans; for example Old World nonhuman primates, such as rhesus macaques, are particularly susceptible to tuberculosis (Aiello, 1998b). Therefore, thorough disinfection of the equipment before and after its use may be warranted, especially when nonhuman primates are involved.
Finally, in positioning an animal in the scanner, care should be taken to maintain airway patency. Animals are usually intubated with an endotracheal tube during scanning, and this helps to ensure that the airway is not obstructed. Care should be exercised to prevent occlusion of the endotracheal tube and to prevent it from being dislodged during positioning. On completion of the imaging procedure, the animal may be extubated once a gag reflex and the ability to swallow are regained. The intravenous line should be removed, and the animal should be observed as it recovers from anesthesia before it is returned to its home cage.
The use of radioactive materials in imaging studies (such as in PET and SPECT imaging) poses specific occupational-health risks that should be considered as part of protocol development. Laboratory staff should be trained in the proper handling and disposal of radioactive materials. Furthermore, the potential for exposure to radiation from the animal and its bodily excretions after injection of radioactive tracers may have to be evaluated and appropriate actions taken to minimize the associated human health risks. Other considerations include thorough disinfection of the equipment if it is also to be used with human subjects or
patients; the potential, during transportation of an animal to and from the imaging facility, for exposure of people who are not involved in the study; and determining whether the air exhausted from the imaging facility is recycled into other building areas.
STEM CELL AND GENE-THERAPY STUDIES
Gene therapy is a technique involving the transfer of genetic material to an individual animal. Transfer can occur directly by administration of a foreign gene to an animal (in vivo) or indirectly through the introduction of genetically modified cells that contain a foreign gene (ex vivo) (NIH, 1995).
During in vivo gene therapy, foreign genes are introduced by administering DNA (naked or complexed with liposomes or proteins) (Cristiano, 2002; Lu et al., 2003; Templeton, 2002), RNA viruses (Quinonez and Sutton, 2002), or DNA viruses (Burton et al., 2002; Lai et al., 2002). To target the nervous system, the virus or DNA can be administered by microinjection into a specific region of the nervous system or by infusion into the bloodstream. Host cells are infected by the virus or will take up the DNA containing the foreign gene. The foreign gene will then exist in the host cells either episomally or integrated into a chromosome. The foreign gene may be chosen because it codes a desired protein, an antisense RNA (Sazani et al., 2002), or a potentially toxic protein (Dilber and Gahrton, 2001). The host cells will then express the foreign gene, changing the genetic profile of the host cells (NIH, 1995).
During ex vivo gene transfer, cells, such as fibroblasts, are removed from the body and genetically modified, often with the same methods used for in vivo gene therapy. The modified cells are then placed in a host animal (Murray et al., 2002).
Stem cell therapy is very similar to ex vivo gene transfer, except that the stem cell is the therapy, rather than a vehicle for a foreign gene. Stem cells have an extensive capacity for self-renewal and are multipotent, giving rise to neurons, astrocytes, and oligodendrocytes (Ostenfeld and Svendsen, 2003). The nervous system does not have the regenerative potential of other cell types, making stem cells a potential therapy for diseases and injuries of the nervous system.
Stem cell and gene therapy are powerful research methods showing promise in animal studies of Parkinson’s disease (Isacson et al., 2003; Sanchez-Pernaute et al., 2001), lysosomal storage disorders (Jung et al., 2001), stroke (Savitz et al., 2003), retinal degeneration (Chacko et al., 2003), and alcoholism (Thanos et al., 2001).
Animal Care and Use Concerns Associated with Stem Cell and Gene Therapy
There are unique animal-care issues related to stem cell and gene therapy. Although the brain is relatively isolated from the immune system, immune and
inflammatory responses do occur when viral gene therapy is used (Thomas et al., 2001). The most common viruses utilized as gene therapy vectors are lentiviruses (Quinonez and Sutton, 2002), herpes simplex viruses (Burton et al., 2002), adenoviruses (Lai et al., 2002), and adeno-associated viruses (Lai et al., 2002). However, newer generations of viral vectors seem to provoke less serious immune responses (Anonymous, 1996).
Stem cells have also been shown to cause an adverse immune response. This immune response is termed graft-versus-host disease and can occur acutely or chronically in a large percentage of patients (Abo-Zena and Horwitz, 2002). Clinical signs of an immune reaction depend on the species of animal, the type of reaction, and the organs affected. The reactions may result in local or systemic symptoms, including such vague symptoms as fever, vomiting, diarrhea, ataxia, and behavior changes and such dramatic symptoms as anaphylactic shock (Aiello, 1998c) or degeneration of the target organ (Yang et al., 1994). Many times immunosuppressive drugs or irradiation is used in combination with stem cell therapies. These can have significant adverse consequences on an animal’s health and well-being, including causing opportunistic infections and cancers due to the immune suppression (Junghanss and Marr, 2002).
Gene therapy also has the potential to be tumorigenic (Donsante et al., 2001) and stem cells have tumorigenic tendencies (Le Belle and Svendsen, 2002; Ruiz et al., 2002). Stem cell transplantation into the brain has also been shown to result in hyperplasia and atypical integration (Zheng et al., 2002). As a result, animals that undergo stem cell or gene therapy should be monitored acutely for immune reactions and chronically for tumor development and neurological dysfunction caused by hyperplasia or atypical integration. A plan for monitoring expected and unexpected consequences should be developed (see Chapter 3).
Occupational Health and Safety
The potential for unexpected consequences of gene therapy extends to the potential for infection of researchers, animal-care technicians, and other laboratory-animal species with the recombinant DNA under investigation. To manage the potential risks, NIH produced Guidelines for Research Involving Recombinant DNA Molecules (NIH, 1998). That document identified which kinds of experiments involving recombinant DNA required institutional biosafety committee approval or notification. Some of the designated experiments include research involving transgenic rodents and the use of infectious DNA or RNA viruses. Appendix Q of the document identifies the physical and biologic containment requirements for handling animals involved in recombinant-DNA research.