Anesthesia and immobilization of small mammals

Introduction


Anesthesia of small mammals provides some unique challenges to veterinary anesthetists. In addition to the usual factors to be considered when selecting an anesthetic plan, the potential interactions between the anesthetic and the particular research protocol needs to be weighed when working in laboratory animal medicine. It is important to discuss the proposed anesthetic regimen with the research group concerned and try to indicate any specific pharmacological properties of the anesthetic that are likely to be relevant.


Laboratory mammal anesthesia


General considerations


The majority of laboratory animals will be young, healthy adults, although in some circumstances animals with concurrent disease will be encountered. Laboratory veterinarians should be able to provide information on the health status of the animals and the incidence of clinical and subclinical disease. Most facilities require that animals undergo a period of acclimatization, usually for 1–2 weeks, prior to their use in research procedures. This provides an excellent opportunity for habituation to handling and restraint. It also provides time for the anesthetist and the animal care staff to assess the behavior and temperament of animals, perform a general clinical examination, and obtain background data such as growth rate and food and water consumption. This information is of considerable value when assessing postoperative recovery. Some basic biologic data for common rodents and rabbits are provided in Table 9.1.



Table 9.1. Physiological data for rodents and rabbits


Source: Flecknell P.A., Richardson C.A., Popovic A. 2007. Laboratory animals. In: Lumb and Jones’ Veterinary Anesthesia and Analgesia, 4th ed. W.J. Tranquilli, J.C. Thurmon, and K.A. Grimm, eds. Ames, IA: Blackwell Publishing, p. 766.

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Anesthetic or analgesic administration may cause discomfort, irritation, and/or ulceration of the skin, mucous membranes, vascular endothelium, or muscles due to low or high pH, temperature (straight from the refrigerator) and/or administered by an inappropriate route (e.g., pentobarbital by intramuscular [IM] route).


Intravenous (IV) injection or placement of IV catheters for anesthetic administration in conscious rodents may be challenging even for experienced clinicians. The use of physical restraint (e.g., restraint tubes) or volatile anesthetics for induction may provide the desired restraint to facilitate this task.


Anesthetic delivery systems


A major advantage of using volatile anesthetics in small mammals is the ease of administration using an anesthetic induction chamber. Ideally, chambers of different sizes should be available (e.g., for animals weighing less than 100 g and for animals weighing up to 1 kg). To reduce the period of involuntary excitement during induction, the chamber should be filled rapidly with the maximum safe induction concentration of the agent. After loss of consciousness, the animal can then be removed from the chamber, and maintained by using a face mask, at a reduced concentration of agent. Providing effective gas scavenging when a face mask is being used can be difficult, but several systems are available commercially that assist with this, for example, the double-mask system.


Anesthesia can also be induced by face mask, and this can be a rapid and convenient technique when using sevoflurane in rats and mice. Volatile anesthetics generally provoke a breath-holding response in rabbits that is often associated with violent struggling unless preanesthetic agents are given. After sedation, the animal should be observed carefully during administration of the anesthetic, and the mask removed briefly if breath holding occurs.


Some laboratory animal units are only equipped to provide compressed air as the anesthetic carrier gas, and this is inadvisable. All of the currently available agents produce some degree of respiratory depression, but hypoxia can be prevented by delivery in oxygen. During recovery from anesthesia, oxygen should continue to be provided until the animal has begun to regain consciousness. If this is not done, then severe hypoxia can occur in some individuals.


Injectable agents


One noteworthy difference relating to the use of injectable anesthetics in small rodents in comparison to dogs and cats is the difficulty of IV access. This results in anesthetic combinations often being administered as single injections by the intraperitoneal (IP), subcutaneous (SC), or IM route, rather than IV, to effect. Although this is a simple and rapid means of producing anesthesia, it has inevitable consequences in relation to the safety of certain anesthetic agents, especially those in which the anesthetic dose is close to the lethal dose. Since there is considerable variation between different strains of rodents in their response to anesthetic agents, anesthetic combinations that either have a broad safety margin or are wholly or partially reversible are preferred when available.


The high metabolic rate of small mammals can result in relatively high dose rates of some anesthetic agents being required to achieve unconsciousness. When coupled with the relative lack of efficacy of agents such as ketamine, this can lead to very high doses being administered (e.g., 100 mg/kg ketamine). Since the drug formulations for veterinary use are normally optimized to give convenient volumes for a dog or a cat, the volume of drug to be injected can be high and, if given IM, can damage tissue and cause pain on injection. Anesthetic combinations that are most widely used in rodents and rabbits are discussed below, and suggested dose rates are listed in Table 9.2.


Ketamine cannot be recommended as the sole anesthetic agent in rodents and rabbits, but when combined with adjunctive drugs such as acepromazine, dexmedetomidine, or opioids, varying planes of anesthesia can be produced. Combinations with tranquilizers often produce only light anesthesia, which is insufficient for surgical procedures, whereas combinations with alpha2 agonists such as dexmedetomidine and xylazine may produce surgical anesthesia. In contrast, in rabbits, combinations of ketamine with acepromazine and diazepam often produce surgical planes of anesthesia.


The use of dexmedetomidine or xylazine with ketamine has the advantage that the sedative–analgesic component of the combination can be reversed with alpha2 antagonists such as atipamezole. Since this anesthetic combination produces cardiovascular and respiratory depression, in addition to other systemic effects such as hyperglycemia and diuresis, it is common to administer the antagonist. Reversing the alpha2 agonist will, of course, reduce the level of postoperative analgesia provided, so that additional agents (e.g., carprofen or buprenorphine) should be administered.


In guinea pigs, the effects of ketamine and xylazine and/or dexmedetomidine are more variable, and surgical anesthesia may not be produced. In all species, if the plane of anesthesia is insufficient, then administering additional doses of the combination may have unpredictable effects. It is preferable to administer a low concentration of an inhalant anesthetic to deepen anesthesia. This same approach can be used to prolong the period of surgical anesthesia. As an alternative, the surgical site can be infiltrated with local anesthetic. As in other species, it is inadvisable to administer atropine routinely when using high doses of alpha2 agonists. Severe hypertension causing mortality has been reported in rats.


Tiletamine in combination with zolazepam (Telazol®, Pfizer Animal Health, New York) has been recommended as an anesthetic for use in rodents and rabbits. As with ketamine–benzodiazepine combinations, the depth of anesthesia produced is not always sufficient to enable surgical procedures. Combining the mixture with xylazine increases anesthetic depth, but the effects are still variable.



Table 9.2. Anesthetic and related drugs for use in rodents and rabbitsa


Source: Flecknell P.A., Richardson C.A., Popovic A. Laboratory animals. In: Lumb and Jones’ Veterinary Anesthesia and Analgesia, 4th ed. W.J. Tranquilli, J.C. Thurmon, and K.A. Grimm, eds. Ames, IA: Blackwell Publishing, p. 775.


IM, intramuscularly; IP, intraperitoneally; IV, intravenously; SC, subcutaneously.

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aNote that considerable between-strain variation occurs, so dose rates should be taken only as a general guide.


bDose (in milliliters per kilogram) of a mixture of one part Hypnorm (fentanyl–fluanisone) plus two parts water for injection, and one part midazolam (5-mg/mL initial concentration).


Etorphine–methotrimeprazine (Immobilon, VetaPharma Ltd. Sheburn-in-Elmet, LEEDS, UK), fentanyl–fluanisone (Hypnorm, VetaPharma Ltd.), and fentanyl–droperidol (Innovar-Vet, Shering-Plough Animal Health, Union, NJ) have been used in rodents and rabbits. All of these agents produce immobility and profound analgesia when used alone, but also cause significant respiratory depression. Fentanyl–fluanisone, when combined with midazolam or diazepam, produces surgical anesthesia in all species. Attempts to develop similar mixtures with the other commercially available neuroleptanalgesic combinations have been less successful.


Fentanyl–fluanisone–midazolam has the advantage that it can be mixed and administered as a single injection, but the active components must be diluted with sterile water before being combined. The mixture is stable for several weeks, but on occasion can crystallize. If this is noted, the mix should be discarded. The fentanyl component can be reversed by using naloxone, but this also reverses all analgesic effects of the combination. It is preferable to reverse the fentanyl with a mixed agonist–antagonist such as butorphanol, nalbuphine, or the partial agonist buprenorphine. This reverses any respiratory depression, although full recovery may be prolonged because of the sedative effects of the midazolam and fluanisone. Flumazenil will reverse the midazolam, but its relatively short half-life means that resedation can occur.


In rabbits, the combination is best administered separately, fentanyl–fluanisone initially to produce sedation, analgesia, and peripheral vasodilation. This makes placement of an IV catheter, for example, in the marginal ear vein, simple and enables slow IV administration of the midazolam to produce the desired effects.


In rats, rabbits, and guinea pigs, mixtures of potent opioids (e.g., fentanyl or sufentanil) can be combined with dexmedetomidine or other alpha2 agonists to produce surgical anesthesia. In some instances, the addition of a benzodiazepine improves the degree of muscle relaxation. These combinations have the advantage that they can be completely reversed by using specific antagonists.


Thiobutabarbital (Inactin, Sigma-Aldrich Corp., St. Louis, MO) has been extensively used to provide medium- to long-term anesthesia in rats. It is considered to have minimal effects on the cardiovascular system; in many respects, however, it resembles other barbiturates, producing reduction in cardiac output and organ blood flows.


Urethane is a hypnotic agent that produces long-lasting and stable anesthesia with minimal cardiovascular and respiratory system depression. Urethane provides good narcosis and muscle relaxation, but the analgesic component may not be adequate. It is commonly used in terminal experiments for central and peripheral neural function studies where reflex responses need to be preserved. When administered IP, the most common route used, it has profound endocrine and metabolic effects, producing superficial damage and necrosis of intra-abdominal organs and massive leakage of plasma into the peritoneal cavity. The onset of the aforementioned effects is rapid. Similar effects have not been observed when urethane was administered SC, IV, or intra-arterially. Urethane is carcinogenic and potentially mutagenic; therefore, it should only be used if other suitable alternatives are not available and only for nonrecovery studies.


Chloralose is used to provide long-lasting anesthesia, particularly in studies in which maintenance of cardiovascular responses is required. Chloralose is a hypnotic, and the anesthesia depth produced may be insufficient to enable surgical procedures to be undertaken. Induction and recovery from chloralose are very prolonged, so the agent is normally used only for terminal procedures. To avoid problems associated with a prolonged onset of action, anesthesia is often induced using another agent (e.g., isoflurane). Following IV cannulation and any other surgical procedures, chloralose is then administered.


Although now rarely used, chloral hydrate, because of its minimal effects on the cardiovascular system, is still used to anesthetize laboratory animals. It is also used in neuropharmacology studies, because it is thought to have a reduced likelihood of interacting with other compounds. It produces medium-duration anesthesia. The anesthesia depth varies between different strains of rodent and can be sufficient for surgical procedures to be undertaken. In some strains of rat, chloral hydrate can cause postanesthetic ileus, which can be fatal. Using a dilute solution of chloral hydrate (36mg/mL) can reduce the incidence of ileus.


Tribromoethanol (Avertin, Aldrich Chemical Co., Milwaukee, WI) is a hypnotic that produces surgical anesthesia in rats and mice that lasts approximately 15–20 minutes. It has become extremely popular for anesthesia of mice for embryo transfer and for the production of transgenic animals, and has been reported to be safe and effective. However, if improperly prepared or poorly stored, tribromoethanol can cause gastrointestinal disturbances. More recently, it has been reported that tribromoethanol can cause low-grade peritoneal irritation, even when correctly prepared and stored. In view of these potential adverse effects, tribromoethanol is better replaced with other anesthetic combinations.


Monitoring and intraoperative care


It is particularly important to provide high standards of perioperative care with laboratory animals, since not only can problems such as hypothermia prolong recovery, they cause widespread physiological effects that may interfere with particular research objectives. As in veterinary clinical practice, one staff member may need to act as both anesthetist and surgeon, so detailed clinical monitoring may be lacking. Use of electronic monitoring devices can therefore be of considerable value. The type of monitoring used should be selected based on the species, duration of anesthesia, type of surgery, and assessment of the risk of complications or emergencies.


Small size is correlated with a rapid heart rate (>300 beats per minute) that may exceed the upper limits of some monitors, and the low signal strength may not be detectable. In addition, small body size limits such procedures as invasive blood pressure monitoring and makes most noninvasive devices ineffective. Some equipment is now available that can function despite these problems, and routine electronic monitoring is becoming increasingly commonplace.


Assessment of respiratory function


Clinical observation of respiratory rate and pattern is relatively straightforward, but can be complicated by placement of surgical drapes, especially in small rodents. In these smaller species, the anesthetic circuit will not normally contain a reservoir bag, so observation of bag movements cannot be used to monitor respiration. Unfortunately, many electronic monitors do not respond to the relatively small respiratory movements and low tidal volumes, especially when used with animals weighing less than 200 g. In these small mammals, direct observation of respiratory rate and pattern may be the only option available.


In common with other species, the pattern, rate, and depth of anesthesia vary both with anesthetic depth and with the anesthetic regimen used. With inhalant anesthetics and the majority of injectable regimens, respiratory rate falls. Typical respiratory rates during anesthesia are 50–100 breaths per minute for small rodents and 30–60 breaths per minute for rabbits. Since many of these animals show a very marked stress-related tachypnea prior to induction, assessment of the degree of respiratory depression should either be based on estimates of normal resting rate (Table 9.1) or established by observing the animals preoperatively when undisturbed. A reduction to less than 50% of the estimated normal respiratory rate should cause concern. Gradual changes in rate, rather than a sudden reduction, are more usual, so keeping an anesthetic record is advisable.


The adequacy of oxygenation and pulse rate can be assessed by using a pulse oximeter, but the high heart rates in rodents may exceed the upper limits of the monitor. A monitor with an upper limit of at least 350 beats per minute is needed, and successful operation may also depend on the type of probe used. It is advisable to try several instruments, probes, and probe positions to find the most reliable combination. In the authors’ experience, a signal can usually be obtained from the hind foot in rodents or the base of the tail. In rabbits, the toe, tail, tongue, and ear are also useful. In particular, the use of an angled probe placed in the mouth has proven particularly reliable.


End-tidal carbon dioxide is difficult to measure in small mammals. The gas volume sampled by side-stream capnographs may be very large in relation to the animal’s tidal volume, and mainstream capnographs usually introduce too much dead space into the anesthetic-breathing circuit. In rabbits, equipment designed for pediatric use in people usually functions well.


Maintenance of an airway may be assisted by placement of an endotracheal tube, but the small size of many laboratory species makes this technically difficult. Rabbits can be intubated by using either an otoscope to visualize the larynx or a blind technique. Prior to intubation, the animal should breathe 100% oxygen for 1–2 minutes. Uncuffed endotracheal tubes should be used to maximize the airway internal diameter (ID); a 3–3.5-mm-diameter tube is usually suitable for a 3–4-kg rabbit. Tubes with a diameter of less than 2.5 mm are required for very small rabbits (<800 g), and these should be purchased from specialty suppliers.


When an otoscope is used the rabbit is positioned on its back. The mouth is opened and its tongue pulled forward into the gap between the incisors and premolars. Care must be used not to injure the tongue on the edges of the incisors. The otoscope speculum should be inserted into the gap between the teeth on the opposite side of the mouth to the tongue and advanced until the end of the soft palate or the larynx is visible. In some animals, the epiglottis will be positioned behind the soft palate, hiding the larynx from view. To expose the larynx, an introducer is used to reposition the epiglottis and soft palate. The larynx should then be sprayed with local anesthetic. The introducer is then advanced through the otoscope speculum, through the larynx, and into the trachea. The otoscope is removed and the endotracheal tube threaded onto the introducer and into the trachea. The introducer is then withdrawn.



Figure 9.1. Intubation of a rabbit by using the “blind technique.” The rabbit is in sternal recumbency, and oxygen is administered for at least 2 minutes prior to intubation. To intubate, the endotracheal tube is placed in the rabbit’s mouth to the level of its larynx. On inspiration, the endotracheal tube is gently advanced into the larynx in the direction of the loudest breath sounds.


Source: Flecknell P.A., Richardson C.A., Popovic A. 2007. Laboratory animals. In: Lumb and Jones’ Veterinary Anesthesia and Analgesia, 4th ed. W.J. Tranquilli, J.C. Thurmon, and K.A. Grimm, eds. Ames, IA: Blackwell Publishing, p. 778.

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To place a tube using the blind technique, the rabbit is positioned in sternal recumbancy, with its head and neck extended upward (Figure 9.1). The endotracheal tube is introduced into the gap between the incisors and premolars and advanced into the pharynx. When the larynx is reached, some increase in resistance is felt. The tube can then be advanced into the larynx and trachea. Successful placement is usually accompanied by a slight cough. In some cases, the tube passes into the esophagus and will need to be withdrawn and repositioned. Passage of the tube is often assisted by gently rotating it through 45° as it is advanced into the larynx. The tube position can be monitored by listening at the end of the tube. If breath sounds can be heard, the tube should be in the pharynx or the trachea. As an alternative to intubation, a laryngeal mask can be used. This technique is easier to master than endotracheal intubation, but manual or mechanical ventilation may not be effective. If only oxygen supplementation is required, a nasal catheter can be passed and positioned in the back of the pharynx.


Intubation of small rodents is made easier if a specialized apparatus is used. A technique using a modified otoscope speculum to visualize the larynx and an over-the-needle catheter as the endotracheal tube is relatively easy to master. Although a variety of other methods have been described, the modified otoscope enables rapid, atraumatic intubation and is supplied together with an instructional video.


After intubation, animals can be maintained on an appropriate anesthetic circuit; for example, a purpose-made low dead space T-piece for small rodents, or a pediatric T-piece or unmodified Bain’s circuit for rabbits.


Assessment of cardiovascular function


Clinical monitoring of the cardiovascular system is difficult in small rodents because of their size. Peripheral pulses are difficult or impossible to palpate, and heart rate is frequently greater than 250 beats per minute. In rabbits and guinea pigs, the chest can be auscultated or palpated, but this is difficult in smaller rodents. An esophageal stethoscope can be used in rabbits.


When using electronic monitoring equipment, the upper rate limits (e.g., 250 or 300 beats per minute) are often exceeded, and some instruments will not detect the low-amplitude electrocardiographic signal.


Other clinical assessments, such as use of capillary refill time, are practical. In all species, assessment of the color of the mucous membranes enables some assessment of peripheral perfusion and oxygen saturation of hemoglobin.


Arterial blood pressure can be measured by using noninvasive systems in larger rabbits or by using pediatric-sized cuffs or specially designed veterinary equipment. Blood pressure can be measured in this way in rats, using a tail cuff, but special apparatus is required. Invasive blood pressure monitoring is possible in all species, but surgical exposure of the vessel is needed in rodents, which tends to limit the use of this technique to nonrecovery procedures. In rabbits, an over-the-needle catheter can be placed in the central ear artery.


Blood volume in all of these species is approximately 70mL/kg of body weight, so small rodents will have very low total blood volumes (e.g., 2 mL for a 30-g mouse). It is therefore critically important to minimize blood loss by careful hemostasis and to monitor blood loss by accurate weighing of swabs and assessing other losses at the surgical site.


Thermoregulation


Small mammals have an increased surface area/body weight ratio that results in rapid cooling during anesthesia. Maintaining body temperature and careful monitoring to ensure this is being achieved effectively are important. Hypothermia can cause delayed recovery from anesthesia and, if severe, can cause cardiac arrest. Rectal temperature should be monitored with an electronic thermometer. The probe size of less expensive instruments is usually appropriate for animals weighing 250 g or more, but specialized instruments are needed for very small rodents (e.g., mice and hamsters). To reduce loss, the area of fur shaved during preparation of the surgical site should be minimized, and use of skin disinfectants should be limited to the minimum necessary to maintain asepsis.


Animals should be placed on a heating pad maintained at 37–39°C. It is important that measures to maintain body temperature are continued into the postoperative period.


Emergencies


All of the measures for coping with anesthetic emergencies applicable to companion animals can be used in laboratory species, but as with many other techniques, small body size can limit or complicate some of these procedures.


To assist ventilation if an animal has not been intubated, its head and neck should be extended, the tongue pulled forward, and the chest gently squeezed between the anesthetist’s thumb and forefinger. If the tongue is difficult to grasp, it can be rolled forward using a cotton swab.


When an animal has been intubated, respiration can be assisted relatively easily. Attempting to assist ventilation by using a face mask is usually unsuccessful, but in small rodents a soft piece of rubber tubing can be placed over the nose and mouth, and the lungs inflated by gently blowing down the tube.


As mentioned earlier, since total blood volume is low in small rodents, every effort should be made to minimize blood loss and avoid overhydration. If fluid therapy is required, this can be delivered via an over-the-needle catheter in the tail vein of rats, the medial tarsal vein in guinea pigs, or the marginal ear vein, cephalic vein, or jugular vein in rabbits.


I f whole blood is required, then a suitable donor may be available in the research facility. All of the commonly available fluid products can be administered safely to small mammals and other laboratory species. In smaller rodents in which IV access is not practicable, IP or SC administration of warmed electrolyte solutions can slowly replace fluid deficits, but will be of minimal benefit if rapid hemorrhage is occurring. In these smaller species, placement of an intraosseous catheter can provide an alternative route for fluid replacement.


If cardiac arrest occurs, external cardiac massage and emergency drugs such as epinephrine can be used when attempting resuscitation.


Postoperative care


If possible, a separate recovery area should be provided, because this makes it easier to provide an optimal environment during this period. It also encourages individual attention and special nursing, if those are required. Most of the commonly used anesthetics will continue to cause some degree of respiratory depression in the immediate postoperative period. In addition to continuing to monitor respiratory function, care must be taken that respiratory obstruction does not occur. Small rodents and rabbits may attempt to hide and push into the corner of a recovery cage, and this can result in airway obstruction. Also, when allowed to recover in a group, rodents may huddle together, which can decrease oxygen availability for the animals at the bottom of the group. Although recovery is often more rapid after use of inhalational agents, significant hypoxia (oxygen saturation of less than 90%) can occur, and this should be prevented by maintaining the animal in an oxygen-enriched environment, either by using a face mask or by delivering oxygen into the incubator until respiratory function is judged to be appropriate.


It is important that measures to maintain normal body temperature are continued in the recovery period. This can often be achieved by allowing animals to recover in a pen or cage in a recovery room (maintained at a high ambient temperature, with supplemental heating of the cage as necessary) or inside an incubator. A temperature of 25–30°C is needed for adult animals and 35–37°C for neonates. If an incubator is unavailable, heating pads and lamps should be provided. Since small mammals can be heated rapidly, care must be taken not to overheat or burn the patient, and a thermometer should be placed next to the animal to monitor the temperature in its immediate environment.


During recovery from anesthesia, animals should be provided with bedding, such as synthetic sheepskin. If this is not available, then towels or a blanket should be used. Sawdust or wood shavings are unsuitable because this type of bedding will often stick to an animal’s eyes, nose, and mouth. Tissue paper is often provided as bedding for small rodents, but it is relatively ineffective because animals usually push it aside during recovery from anesthesia and end up lying in the bottom of a plastic cage soiled with urine and feces.


Drinking water should be available, but care must be taken that this not be spilled or placed in a bowl that the animal may drown in. If the animal’s skin becomes wet, it will lose heat rapidly. Small rodents are usually accustomed to using water bottles, so this is rarely a problem, but it can present difficulties with rabbits, guinea pigs, ferrets, and larger species.


It may also be necessary to provide fluid therapy postoperatively. This can be given by IV infusion in larger animals, but it is most convenient in small rodents to give warmed (37°C) SC or IP dextrose/saline at the end of surgery (Table 9.3).


Food should be provided for most laboratory species immediately after they regain consciousness and are at a minimal risk of aspiration. A mash made by soaking pelleted diet in warm water is often rapidly consumed by small rodents, providing both additional fluid as well as food intake.


Pain assessment


If analgesics are administered appropriately, then it is essential that attempts are made to assess the severity of postoperative pain. Only when this is done can one determine whether an appropriate dose of analgesic has been administered. Pain assessment is also essential to judge whether an appropriate type of analgesic has been selected, when to repeat dosing, and when to discontinue therapy.



Table 9.3. Volumes of fluid for administration to rodents and rabbits


Source: Flecknell P.A., Richardson C.A., Popovic A. 2007. Laboratory animals. In: Lumb and Jones’ Veterinary Anesthesia and Analgesia, 4th ed. W.J. Tranquilli, J.C. Thurmon, and K.A. Grimm, eds. Ames, IA: Blackwell Publishing, p. 781.

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