In: Recent Advances in Anesthetic Management of Large Domestic Animals , E. P. Steffey (Ed.) Publisher: International Veterinary Information Service (www.ivis.org), Ithaca, New York, USA. Anesthetic Management of Cattle (Last Updated: 15-Feb-2001 ) T. Riebold Veterinary Teaching Hospital, College of Veterinary Medicine, Oregon State University, Corvallis, Oregon, USA. Introduction As in other species, sedation and anesthesia are required in cattle for surgical or diagnostic procedures. The decision to induce general anesthesia may be influenced by cattle’s anatomic and physiologic characteristics. Cattle usually accept physical restraint well and that, in conjunction with local or regional anesthesia, is often sufficient to allow completion of many procedures. Other diagnostic and surgical procedures that are more complex require general anesthesia. The intent of this chapter is to review current knowledge and techniques of general anesthetic management of cattle as applied in clinical practice in the United States. Preanesthetic Preparation Considerations for preanesthetic preparation include fasting, assessment of hematologic and blood chemistry values, venous catheterization, and estimation of body weight. Cattle as ruminants are susceptible to complications associated with recumbency and general anesthesia; tympany, regurgitation, and aspiration pneumonia. Accordingly, it is recommended that calves be fasted 12 - 18 hours and deprived of water for 8 - 12 hours. Adult cattle should be fasted 18 - 24 hours and deprived of water for 12 - 18 hours. In nonelective cases, this is often not possible and precautions should be taken to avoid aspiration of gastric fluid and ingesta. Fasting neonates is not advisable because hypoglycemia may result. Fasting and water deprivation will decrease the incidence of tympany and regurgitation by decreasing the volume of fermentable ingesta but may also produce bradycardia in cattle [1]. Additionally, pulmonary functional residual capacity maybe better preserved in the fasted anesthetized ruminant [2]. Even with these precautions, some cattle will become tympanitic while others will regurgitate. Hematologic and blood chemistry values should be determined and evaluated as appropriate before induction of anesthesia. Venipuncture and catheterization of the jugular vein are often performed prior to anesthesia. Catheters up to 16 gauge are appropriate for calves while adult cattle require 12 - 14 gauge catheters. Infiltration of a local anesthetic at the site of catheterization is recommended. Anticholinergics are usually not administered to cattle prior to induction of anesthesia as they do not consistently decrease salivary secretions unless used in high doses and repeated frequently. Usual doses of atropine to prevent bradycardia in cattle (0.06 - 0.1 mg/kg IV) do not prevent salivation during anesthesia. Glycopyrrolate (0.005 - 0.01 mg/kg IM or 0.002 - 0.005 mg/kg IV) may be substituted for atropine [3]. Sedation/Restraint The alpha-2 agonist drugs are currently most commonly used to induce tranquilization and/or sedation in cattle. Other drugs such as acepromazine, chloral hydrate, and pentobarbital have long histories of use with cattle and continue to be commercially available, however, their contemporary importance has become mostly limited to special circumstances. Diazepam can also be used in calves and other small ruminants. Readers are referred to available texts for further discussion on these older drugs [4-8]. Xylazine, detomidine, medetomidine, and romifidine are alpha-2 agonists. Of these, xylazine is presently most often used in the U.S. to provide sedation or, in higher doses, restraint (recumbency and light planes of general anesthesia) in cattle [5, 9,10]. There appears to be some variation in response to xylazine within a species. Hereford cattle have been shown to be more sensitive to xylazine than Holstein cattle [11] and anecdotal evidence indicates that Brahmans are the most sensitive of the breeds [7]. High environmental ambient temperature will cause a pronounced and prolonged response to xylazine in cattle [12]. Xylazine also will cause hyperglycemia and hypoinsulinemia in cattle and sheep [13-16]. Additionally, it will cause hypoxemia and hypercarbia in cattle [11,17] and can cause pulmonary edema [18]. Finally, xylazine has an oxytocin-like effect on the uterus of pregnant cattle [19] and sheep [20]. Detomidine is used to a lesser extent in the U.S. but is also effective for providing sedation and/or analgesia in cattle [8, 21]. It does not appear to have the same effect on the gravid uterus as xylazine in cattle [21] and is the drug of choice when sedation is needed in pregnant cattle. The degree of sedation or restraint produced by xylazine depends on the route of injection, dosage given, and the animal’s temperament. Low doses (0.015 - 0.025 mg/kg IV or IM) will provide sedation without recumbency in cattle. Detomidine is given at 2.5 - 10.0 µg/kg IV in cattle to provide standing sedation of approximately 30 - 60 minutes
Transcript
In: Recent Advances in Anesthetic Management of Large Domestic Animals , E. P. Steffey (Ed.) Publisher: International Veterinary Information Service (www.ivis.org), Ithaca, New York, USA.
Anesthetic Management of Cattle (Last Updated: 15-Feb-2001 ) T. Riebold
Veterinary Teaching Hospital, College of Veterinary Medicine, Oregon State University, Corvallis, Oregon, USA.
Introduction As in other species, sedation and anesthesia are required in cattle for surgical or diagnostic procedures. The decision to induce general anesthesia may be influenced by cattle's anatomic and physiologic characteristics. Cattle usually accept physical restraint well and that, in conjunction with local or regional anesthesia, is often sufficient to allow completion of many procedures. Other diagnostic and surgical procedures that are more complex require general anesthesia. The intent of this chapter is to review current knowledge and techniques of general anesthetic management of cattle as applied in clinical practice in the United States.
Preanesthetic Preparation Considerations for preanesthetic preparation include fasting, assessment of hematologic and blood chemistry values, venous catheterization, and estimation of body weight. Cattle as ruminants are susceptible to complications associated with recumbency and general anesthesia; tympany, regurgitation, and aspiration pneumonia. Accordingly, it is recommended that calves be fasted 12 - 18 hours and deprived of water for 8 - 12 hours. Adult cattle should be fasted 18 - 24 hours and deprived of water for 12 - 18 hours. In nonelective cases, this is often not possible and precautions should be taken to avoid aspiration of gastric fluid and ingesta. Fasting neonates is not advisable because hypoglycemia may result. Fasting and water deprivation will decrease the incidence of tympany and regurgitation by decreasing the volume of fermentable ingesta but may also produce bradycardia in cattle [1]. Additionally, pulmonary functional residual capacity maybe better preserved in the fasted anesthetized ruminant [2]. Even with these precautions, some cattle will become tympanitic while others will regurgitate. Hematologic and blood chemistry values should be determined and evaluated as appropriate before induction of anesthesia. Venipuncture and catheterization of the jugular vein are often performed prior to anesthesia. Catheters up to 16 gauge are appropriate for calves while adult cattle require 12 - 14 gauge catheters. Infiltration of a local anesthetic at the site of catheterization is recommended. Anticholinergics are usually not administered to cattle prior to induction of anesthesia as they do not consistently decrease salivary secretions unless used in high doses and repeated frequently. Usual doses of atropine to prevent bradycardia in cattle (0.06 - 0.1 mg/kg IV) do not prevent salivation during anesthesia. Glycopyrrolate (0.005 - 0.01 mg/kg IM or 0.002 - 0.005 mg/kg IV) may be substituted for atropine [3].
Sedation/Restraint The alpha-2 agonist drugs are currently most commonly used to induce tranquilization and/or sedation in cattle. Other drugs such as acepromazine, chloral hydrate, and pentobarbital have long histories of use with cattle and continue to be commercially available, however, their contemporary importance has become mostly limited to special circumstances. Diazepam can also be used in calves and other small ruminants. Readers are referred to available texts for further discussion on these older drugs [4-8]. Xylazine, detomidine, medetomidine, and romifidine are alpha-2 agonists. Of these, xylazine is presently most often used in the U.S. to provide sedation or, in higher doses, restraint (recumbency and light planes of general anesthesia) in cattle [5, 9,10]. There appears to be some variation in response to xylazine within a species. Hereford cattle have been shown to be more sensitive to xylazine than Holstein cattle [11] and anecdotal evidence indicates that Brahmans are the most sensitive of the breeds [7]. High environmental ambient temperature will cause a pronounced and prolonged response to xylazine in cattle [12]. Xylazine also will cause hyperglycemia and hypoinsulinemia in cattle and sheep [13-16]. Additionally, it will cause hypoxemia and hypercarbia in cattle [11,17] and can cause pulmonary edema [18]. Finally, xylazine has an oxytocin-like effect on the uterus of pregnant cattle [19] and sheep [20]. Detomidine is used to a lesser extent in the U.S. but is also effective for providing sedation and/or analgesia in cattle [8, 21]. It does not appear to have the same effect on the gravid uterus as xylazine in cattle [21] and is the drug of choice when sedation is needed in pregnant cattle. The degree of sedation or restraint produced by xylazine depends on the route of injection, dosage given, and the animal's temperament. Low doses (0.015 - 0.025 mg/kg IV or IM) will provide sedation without recumbency in cattle. Detomidine is given at 2.5 - 10.0 µg/kg IV in cattle to provide standing sedation of approximately 30 - 60 minutes
duration [5, 8]. Higher doses of xylazine (0.1 mg/kg IV or 0.2 mg/kg IM) will provide recumbency and light planes of general anesthesia in cattle for approximately one hour [5,7]. Higher doses can be expected to induce longer periods of recumbency. Detomidine at 40 µg/kg IV will produce profound sedation and recumbency [5]. Higher doses of detomidine (100 µg/kg) administered by dart have been used to immobilize free ranging cattle [22]. Approximately 15 minutes were required for onset of action [22]. Medetomidine has been given at 30.0 µg/kg IM to produce recumbency in calves lasting 60 - 75 minutes [23]. If one was to extrapolate from use of alpha-2 agonists in sheep, romifidine at 50 µg/kg IV or medetomidine at 10 µg/kg IV could be expected to produce recumbency in cattle [24]. Combinations of xylazine and butorphanol have been used in cattle to provide neuroleptanalgesia. Doses are 0.01 - 0.02 mg/kg IV of each drug given separately in cattle [25]. Duration of action is approximately 1 hour. Combinations of detomidine (0.07 mg/kg) and butorphanol (0.04 mg/kg) have also been used to immobilize free ranging cattle [22]. Sedation following use of alpha-2 agonists can be reversed by alpha-2 adrenoceptor antagonists, yohimbine, tolazoline, atipamezole, and idazoxan. Yohimbine is given at 0.12 mg/kg IV although there is some variability in response to its administration in cattle [26,27] Tolazoline is given at 0.5 - 2.0 mg/kg IV [27]. There are anecdotal reports of death associated with tolazoline administration in cattle, usually following higher doses of the drug given to animals with compromised physical status. Tolazoline should be given at 0.5 - 1.0 mg/kg IV. If sufficient arousal does not occur, additional tolazoline could be given. Tolazoline given at 2.0 mg/kg IV will cause hyperesthesia in unsedated cattle. [28] Idazoxan is given at 0.05 mg/kg IV to calves [29]. Atipamezole is given at 20 - 60 µg/kg IV [30]. Doxapram, an analeptic, can be used to augment the response of yohimbine or tolazoline to alpha-2 agonist sedated cattle. Doxapram (1.0 mg/kg IV) given alone has been shown to be effective in cattle [31].
Induction General anesthesia can be induced by either injectable or, especially in young calves, inhalation techniques. Available drugs include: thiobarbiturates, ketamine, guaifenesin, tiletamine-zolazepam, propofol, halothane, isoflurane, and sevoflurane [5].
Thiopental - The thiobarbiturates, thiopental and thiamylal, have been used extensively in veterinary anesthesia, either alone and in combination with guaifenesin to quickly induce general anesthesia. Thiamylal is similar to thiopental but more potent and is no longer commercially available in North America. Its use will not be further discussed. Recovery from induction doses of thiopental is based upon redistribution of the drug from the brain to other tissues in the body. Metabolism of the agent continues for some time following recovery until final elimination occurs. Maintenance of anesthesia with thiopental is not recommended because saturation of tissues with thiopental causes recovery to be dependent on metabolism and recovery will be prolonged. Concurrent use of nonsteroidal antiinflammatory drugs is contraindicated because they may displace the thiobarbiturates from plasma protein and delay recovery [32]. Thiopental is given at 6 - 10 mg/kg IV in unsedated animals and will provide approximately 10 - 15 minutes of anesthesia. Ketamine - Ketamine is commonly used in veterinary anesthesia. It also provides mild cardiovascular stimulation and is safer than thiopental in sick animals. Following anesthetic induction doses, ketamine often does not eliminate the swallowing reflex however, tracheal intubation can be accomplished. While ketamine will induce immobilization and incomplete analgesia when given alone, addition of a sedative or tranquilizer will improve muscle relaxation and quality of anesthesia. Most commonly xylazine or diazepam is recommended, although the availability of detomidine offers another alternative. Xylazine (0.1 - 0.2 mg/kg IM) can be given followed by ketamine (10 - 15 mg/kg IM) in calves [33]. Anesthesia usually lasts about 45 minutes and can be prolonged by injection of additional ketamine at 3 - 5 mg/kg IM or 1 - 2 mg/kg IV. The longer duration of action of xylazine obviates the need for readministration of xylazine in most cases. Alternatively, xylazine (0.03 - 0.05 mg/kg IV) followed by ketamine (3 - 5 mg/kg IV) can be used to provide anesthesia of 15 - 20 minutes duration [5,9,10]. Adult cattle can be anesthetized with xylazine (0.1 - 0.2 mg/kg IV) followed by ketamine (2.0 mg/kg IV) [34]. The lower dose of xylazine is used when cattle weigh greater than 600 kg [34]. Duration of anesthesia is approximately 30 minutes; anesthesia can be prolonged for 15 minutes with additional ketamine (0.75 - 1.25 mg/kg IV). Diazepam (0.1 mg/kg IV) followed immediately by ketamine (4.5 mg/kg IV) can be used in cattle. Muscle relaxation is usually adequate for tracheal intubation although the swallowing reflex may not be completely obtunded. Anesthesia usually is of 10 - 15 minutes duration following diazepam-ketamine with recumbency of up to 30 minutes [4-6,10]. Medetomidine has been combined with ketamine to provide anesthesia in calves. Because medetomidine (20 µg/kg IV) is much more potent than xylazine, very low doses of ketamine (0.5 mg/kg IV) can be used [30]. However, use of a local anesthetic at the surgical site may be required when ketamine is used at this dose [30]. Consequently, complete reversal of anesthesia can be achieved with alpha-2 antagonists without excitement occurring during recovery. Guaifenesin - Guaifenesin, a centrally acting skeletal muscle relaxant, can be used alone to induce recumbency in cattle. Addition of ketamine or thiopental to guaifenesin improves induction quality and decreases the volume required for induction and improves muscle relaxation when compared to induction with ketamine or the thiobarbiturates given alone. Typically, 5% guaifenesin solutions are used as hemolysis can occur with 10% guaifenesin solutions [35,36]. Commonly these solutions are given rapidly to effect in either tranquilized or untranquilized patients. The calculated dose is 2.0 ml/kg. The amount of ketamine added to guaifenesin varies but is commonly 1.0 - 2.0 mg/ml. The amount of
thiobarbiturate added to guaifenesin varies but is commonly 2.0 - 4.0 mg/ml. For convenience, guaifenesin-based mixtures may be injected with large syringes rather than administered by infusion to calves. Following induction, guaifenesin-based solutions can be infused to effect to maintain anesthesia. Xylazine may also be added to ketamine-guaifenesin solutions for induction and maintenance of anesthesia in cattle [34,37]. Final concentrations are guaifenesin (50 mg/ml), ketamine (1 - 2 mg/ml), and xylazine (0.1 mg/ml). This solution is infused at 0.5 - 1 ml/kg IV for induction. Anesthesia can be maintained by infusion of the mixture at 1.5 ml/kg/hr for calves [37] and at 2 ml/kg/hr for adult cattle [34,37] although final administration rate will vary depending upon circumstances. If the procedure requires greater than 2 ml/kg of the guaifenesin-ketamine-xylazine mixture to allow completion of the surgical procedure, the amount of xylazine added should be decreased by 50% because its duration of action is longer than the other two agents. [25] Alternatively, a mixture of guaifenesin (50 mg/ml), ketamine (1 mg/ml), and xylazine (0.05 mg/ml) could be infused at 2.0 ml/kg following induction for maintenance. If anesthetic depth becomes insufficient, the infusion rate should be increased by 10 - 20%. Use of an infusion pump allows administration to be more precise and convenient. Recovery usually occurs within 30 - 45 minutes. Tiletamine-Zolazepam - Tiletamine-zolazepam is a proprietary combination available for use as an anesthetic agent in cats and dogs. In many respects tiletamine-zolazepam can be considered to be similar to ketamine pre-mixed with diazepam. Tiletamine-zolazepam can be used successfully with or without xylazine in cattle. However, addition of xylazine to tiletamine-zolazepam lengthens duration of effect. Tiletamine-zolazepam has been given at 4.0 mg/kg IV to healthy untranquilized calves and found to cause minimal cardiovascular effects and provided anesthesia of 45 - 60 minutes duration [38]. Xylazine (0.1 mg/kg IM) followed immediately by tiletamine-zolazepam (4.0 mg/kg IM) produced onset of anesthesia within 3 minutes and duration of anesthesia of approximately 1 hour [39]. Calves were able to stand approximately 130 minutes following injection. Increasing xylazine to 0.2 mg/kg IM increased duration of anesthesia and recumbency and the incidence of apnea [39]. The drugs can be administered intravenously. Xylazine can be given at 0.05 mg/kg IV followed by tiletamine-zolazepam at 1.0 mg/kg IV [34]. Propofol - Propofol is a nonbarbiturate, nonsteroidal hypnotic agent and can be used to provide brief periods of anesthesia. Economic considerations limit the applicability of propofol as it is an expensive drug to use in adult cattle. Inhalation Agents - Anesthesia can be induced by mask in calves weighing less than 150 kg. If a commercial mask is unavailable a mask can be made by cutting the bottom out of a one gallon plastic jug or other container of appropriate size and padding the edges with cotton and tape. The mask must fit tightly around the calf's muzzle to prevent inspiration of atmospheric air and dilution of anesthetic gases. Halothane and isoflurane are agents of choice for use in cattle because they give short induction and recovery times. Sevoflurane and desflurane are also excellent choices for mask induction but their use carries considerable expense. The addition of nitrous oxide to the gas mixture hastens induction. Unless its use is contraindicated, nitrous oxide may be used as 50% of the total gas flow with one of the inhalation agents for mask induction of calves and then discontinued after intubation. It is recommended that nitrous oxide be discontinued after induction to avoid its accumulation in the rumen. Normal oxygen flow rates during induction are 3 - 6 liters/minute. Normal vaporizer settings are 3 - 5% halothane or isoflurane and 5 - 7% for sevoflurane during induction. The higher flow rates and vaporizer settings are used for bigger calves.
Maintenance Tracheal intubation is recommended in cattle to provide a secure airway and prevent aspiration of salivary and ruminal contents if passive regurgitation occurs. Several techniques (blindly, digital palpation, or direct laryngoscopy) can be used to accomplish intubation and the reader is referred to descriptions of those techniques. An endotracheal tube of appropriate size is inserted and manipulated into the larynx (Table 1). The technique is similar to that performed in small ruminants [4-8,40].
Body Weight (kg) Endotracheal Tube Size (mm id.)
< 30 4 - 7
30 - 40 8 - 10
60 - 80 10 - 12
100 12
200 - 300 14 - 16
300 - 400 16 - 22
400 - 600 22 - 26
> 600 26
> 600 26
Anesthesia in cattle can be maintained with halothane, isoflurane, or sevoflurane. Economic issues often dictate which agent is used. Conventional small animal anesthetic machines can be used to anesthetize calves weighing less than 40 kg. Conventional human anesthetic machines or small animal machines with expanded soda lime canisters are adequate for animals up to about 200 kg. Oxygen flow rates of 20 ml/kg/minute during induction and 12 ml/kg/minute during maintenance with minimal flow rates of 1 L/minute are adequate. Anesthesia is usually maintained with halothane at 1.5 - 2.5% or isoflurane at 1.5 - 3% or sevoflurane at 2.5 - 4%. Because cattle have a respiratory pattern characterized by rapid respiratory rate and small tidal volume, higher vaporizer settings (e.g., halothane 2 - 3%) may be required to maintain anesthesia in spontaneously breathing patients. Vaporizer setting can be decreased if controlled ventilation is used.
Supportive Therapy Supportive therapy is an important part of anesthetic practice and can exert a strong influence on recovery and patient morbidity/mortality. Supportive therapy includes patient positioning, fluid administration, mechanical ventilation, cardiovascular support, good monitoring techniques, and oxygen administration to cattle under intravenous anesthesia. These techniques are addressed more completely in other chapters and other texts [4-8,41-43]. The reader is encouraged to consult those references for additional information. As with any species, good anesthetic techniques require monitoring to allow drug administration to meet the animal's requirements and prevent excessive insult to the cardiovascular, respiratory, central nervous, and musculoskeletal systems, thereby decreasing risk of complications. Monitoring techniques are similar to those employed in horses [4,6,44,45]. In healthy anesthetized adult cattle, heart rate is usually 70 - 90 beats/minute. Animals that have received an anticholinergic will have an increased heart rate. Normal heart rate for calves varies with age. Younger calves will have a heart rate of 90 - 130 beats/minute, decreasing as they mature. Pulse pressure can be ascertained by palpating the common digital, caudal auricular, radial, and saphenous arteries. Mucous membranes should be pink although the mucous membranes of some cattle are pigmented, making assessment difficult. Electrocardiography (ECG) is used with either standard limb leads (I, II, III) or a dipole lead for detection of cardiac rate and rhythm disturbances. Arterial pressure provides an accurate variable for assessing depth of anesthesia and can be monitored with either direct or indirect techniques. Normal arterial pressure values in anesthetized cattle are systolic pressure, 120 - 150 mmHg; diastolic pressure, 80 - 110 mmHg; and mean arterial pressure, 90 - 120 mmHg; and exceed those of standing cattle [46]. The respiratory system is evaluated by monitoring respiratory rate and tidal volume. Spontaneous breathing rates are usually 20 - 30 breaths/minute in adult cattle and usually 20 - 40 breaths/minute in calves. Awake cattle have a decreased tidal volume when compared to horses and that relationship persists during anesthesia [47]. Normal arterial blood gas values for anesthetized cattle are similar to those of anesthetized horses. Respiratory gas analysis to determine end tidal CO2 and anesthetic agent concentration can be performed to provide additional information. Care must be exercised during selection of agent analyzers. Ruminants often have detectable amounts of methane (and other gases) in their expired gas. Methane (and perhaps other gases) will be interpreted as the anesthetic agent by some infrared monitors and falsely report anesthetic concentration [48]. The central nervous system can be monitored by observation of ocular reflexes. The palpebral reflex disappears with minimal depth of anesthesia in cattle and is usually of no value during anesthesia. Dorsoventral rotation of the globe will occur as anesthetic depth changes in cattle [49]. The eyeball is normally centered between the palpebra. As anesthesia is induced the eyeball rotates ventrally, with the cornea being partially obscured by the lower eye lid. As depth of anesthesia is increased, the cornea becomes completely hidden by the lower eyelid; this sign usually indicates adequate depth of surgical anesthesia. A further increase in anesthesia is accompanied by dorsal rotation of the eyeball. Dorsal movement is complete when the cornea is centered between the palpebra; this sign indicates deep surgical anesthesia with profound muscle relaxation. During recovery eyeball rotation occurs in reverse order to that occurring during induction [49]. Nystagmus usually does not occur during anesthesia of cattle and when it does occur, can not be correlated with changes in depth of anesthesia. The corneal reflex should be present.
Recovery Cattle recover well from general anesthesia and seldom experience emergence delirium or make premature attempts to stand. When an alpha-2 agonist is used as part of the anesthetic regimen, an antagonist can be used to hasten recovery [30,50,51] Extubation of cattle should not occur until the laryngeal reflex has returned and the animal begins to chew. If the patient has regurgitated the buccal cavity and pharynx should be lavaged to prevent aspiration of the material. In these instances, the endotracheal tube should be withdrawn with the cuff inflated in an attempt to remove any material that may have located in the trachea. Although cattle recover well from general anesthesia with minimal assistance, an attendant should be available.
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Therapy. St. Louis: CV Mosby, 1991; 153-179. 45. Riebold, TW. Monitoring equine anesthesia. In: Principles and Techniques of Equine Anesthesia. Vet Clin N Am (Eq Pract). Philadelphia: WB Saunders Company. 1990; 6:607-624. 46. Matthews NS, Gleed RD, Short CE. Cardiopulmonary effects of general anesthesia in adult cattle. Mod Vet Pract 1986; 67:618-620. 47. Gallivan GJ, McDonell WN, Forrest JB. Comparative ventilation and gas exchange in the horse and cow. Res Vet Sci 1989; 46:331-336. 48. Moens YPS. The reliability of recent monitoring developments in veterinary anaesthesia. In: Proceedings of the 5th International Cong Vet Anes 1994; 33-35. 49. Thurmon JC, Romack FE, Garner HE. Excursion of the bovine eyeball during gaseous anesthesia. Vet Med/Sm An Clin 1968; 63:967-970. 50. Kruse-Elliott KT, Riebold TW, Swanson CR. Reversal of xylazine-ketamine anesthesia in goats. Vet Surg 1987; 16:321. 51. Young DB, Shawley RV, Barron SJ. Tolazoline reversal of xylazine-ketamine anesthesia in calves. Vet Surg 1988; 18:171.
All rights reserved. This document is available on-line at www.ivis.org. Document No. A0603.0201.
In: Recent Advances in Anesthetic Management of Large Domestic Animals, SteffeyE.P. (Ed.)Publisher: International Veterinary Information Service (www.ivis.org)
Anesthetic Management of the Horse: Intravenous Anesthesia (31 Oct2000)
K.R. MamaDepartment of Clinical Sciences, College of Veterinary Medicine and Biomedical Sciences, ColoradoState University, Fort Collins, Colorado, USA. IntroductionFor decades thiobarbiturates have been used to induce and maintain short term (15 - 30 minutes) generalanesthesia in equids. Thiamylal is no longer commercially available, at least in the United States, but useof thiopental remains widespread. In earlier years these ultrashort acting barbiturates were the agents ofchoice to induce short term recumbency in tranquilized horses. Later guaifenesin and more recentlyxylazine (or other alpha-2 agonist type drugs) were administered in combination with or immediately priorto the intravenous injection of the thiobarbiturates to improve upon the quality of anesthetic induction andrecovery. These improved techniques of general anesthesia facilitated surgical and other therapeuticprocedures requiring more prolonged recumbency (e.g. 30 - 60 minutes). However, as duration ofbarbiturate administration increased, the duration of recovery from anesthesia became longer and thequality of recovery became more unpredictable and in some cases dangerous to the animal and associatedpersonnel.Today, dissociative drugs such as ketamine have largely replaced thiobarbiturates for routine equineanesthetic management. This change has improved upon the qualitative consistency of both anestheticinduction and recovery. Because the dissociative agents have undesirable central nervous systemexcitatory properties their use in equids requires concurrent administration of other behavior modifyingdrugs such as an alpha-2 agonist (e.g. xylazine). A brief review and update of specific contemporarytechniques follow.
Anesthetic Induction and Maintenance - Dissociative Drug BasedXylazine/ketamine and xylazine/guaifenesin/ketamine - the use of xylazine and ketamine for induction andshort-term anesthetic maintenance in the horse has been extensively described [1-5]. Recently,investigators evaluated the behavioral and cardiopulmonary responses associated with varied dosecombinations of xylazine and ketamine during anesthetic maintenance [6-8].Response to noxious stimulation varied with drug and dose as did speed of recovery from anesthesia;quality of recovery was good to excellent in all horses. Bradydysrhythmias and relative hypoxemia werecommonly recorded during xylazine/ketamine maintenance. Blood pressure recorded in horses receivingxylazine and ketamine was higher than that previously reported for inhalation anesthesia; cardiac outputwas similar (low) [8]. Low cardiac output in the face of good blood pressure is likely a direct result ofdrug actions (e.g., bradycardia, vasoconstriction). In support of this interpretation, Singh et al., showedthat pretreatment with glycopyrrolate (2.5 µg/kg) minimized the negative influence (likely due to thedecreased heart rate) of xylazine and ketamine on cardiac output [9]. The stimulatory influence of PaO2on cardiac output must also be considered. Results from a study by Mama et al., indicate that duringmaintenance with equipotent doses of xylazine and ketamine, cardiac output was significantly higher inhorses breathing room air (likely due to sympathetic stimulation resulting from hypoxemia) as comparedto those breathing 100% oxygen [6].The addition of guaifenesin to xylazine and ketamine for short-term equine anesthetic maintenance wasfirst described in the mid 1970’s [3]. This combination of drugs provides desirable characteristics(analgesia, unconsciousness and muscle relaxation) associated with general anesthesia, and horses tend torecover well and in a predictable manner after drug administration is discontinued. Hypotension, which is
commonly observed during inhalation anesthetic maintenance, is rarely observed withxylazine/guaifenesin/ketamine maintenance when horses breathe air. These positive attributes have led tothe common use of these three drugs in clinical veterinary practice both for induction of anesthesia andduring maintenance of anesthesia for procedures (e.g. laceration repair, castration, etc.) lasting up to 1hour.Maintenance of anesthesia for greater than 1 hour with injectable anesthetic techniques is not usuallyrecommended due to the potential for cumulative drug effects, which in turn prolong recovery fromanesthesia and may negatively influence its quality. The potential for hypoxemia during recumbencymaintained with this drug combination also limits its long-term use if oxygen is not supplemented (as inmost circumstances of anesthesia performed in a non-hospital [i.e. field] setting) [3,5]. With widespreaduse of this injectable technique, it has also become apparent that due to the presence of reflex activity (e.g.blinking, swallowing), surgical conditions are not ideal for procedures involving the eye or upper airway[5]. The presence of reflex activity at a surgical plane of anesthesia can also confound evaluation ofanesthetic depth and may lead to inappropriate drug dosing.Detomidine/ketamine and detomidine/guaifenesin/ketamine - detomidine (20 µg/kg) plus ketamine (2mg/kg) and detomidine plus guaifenesin and ketamine (varying infusion doses) have been studied forinduction and maintenance of anesthesia in ponies and to a limited degree in horses [10-13]. While theconcurrent administration of other drugs (e.g. acepromazine, flunixin) with potential behavioral,cardiovascular or analgesic effects may influence interpretation of the results from these studies, authorsreport that blood pressure and cardiac index was well maintained. When compared to halothane anesthesiaduring surgical castration authors report that cortisol levels increased over pre-surgical levels to a greaterdegree during inhalation anesthesia than during maintenance with detomidine, guaifenesin and ketamine.Surgical conditions and recovery from anesthesia were comparable for the two protocols.Romifidine/ketamine and romifidine/guaifenesin/ketamine - the use of romifidine (100 µg/kg) andketamine (2.2 mg/kg) prior to maintenance of anesthesia with halothane was first described in the early90’s [14]. While induction of anesthesia was rated excellent in 33 of 45 horses, swallowing, rigidity andmild muscle tremors were observed early in the recumbency period. In another study anesthesia wasinduced in a similar manner and then maintained with an additional intravenous bolus of both romifidine(40 µg/kg) and ketamine (1.1 mg/kg) administered approximately 18 - 20 minutes after the initial dose ofketamine [15]. A positive response (purposeful skeletal muscle movement) to a pinprick was observed at35 minutes after the initial ketamine dose; lateral recumbency was maintained for an average of 43minutes. Compared to pre-sedation values heart rate and arterial oxygen tensions were decreased andmean arterial pressure was increased during anesthetic induced recumbency.McMurphy et al., compared cardiopulmonary effects of halothane and total intravenous anesthesiamaintained with romifidine (82.5 µg/kg/h), ketamine (6.6 mg/kg/h) and guaifenesin (100 mg/kg/hour for30 min, then 50 mg/kg/h) [16]. Although differences in some recorded variables (e.g. heart rate, meanarterial pressure) were observed at various time points during the study, authors concluded that except forchanges in pulmonary artery pressures, there were no significant differences in recorded cardiopulmonaryvariables between the two anesthetic techniques.Xylazine/diazepam/ketamine, romifidine/tiletamine-zolazepam and xylazine/climazolam/ketamine - theuse of benzodiazepines in place of guaifenesin has been evaluated for anesthetic maintenance. In the early90’s Brock et al., characterized behavioral and cardiopulmonary effects associated with use of two dosesof diazepam (0.05 and 0.1 mg/kg) in horses also receiving xylazine (0.3 mg/kg) and ketamine (2.0 mg/kg)for anesthetic induction [17]. Diazepam (0.1 mg/kg) was felt to be equivalent to guaifenesin (100 mg/kg)in this protocol.The use of tiletamine (dissociative) and zolazepam (benzodiazepines) has also been evaluated in horsespremedicated with romifidine [15]. The quality of anesthesia was smooth and horses remained in lateralrecumbency an average of 45 minutes. In another study anesthesia was maintained for 120 minutes withclimazolam (0.4 mg/kg/h) and ketamine (6 mg/kg/h) in ponies premedicated with xylazine andacepromazine [18]. Although recovery quality was not as good as that reported with previously describedtechniques, authors felt that the cardiopulmonary function was better maintained.
Anesthetic Induction and Maintenance - Propofol BasedPropofol is an anesthetic agent characterized as having a rapid onset and short duration of action. Due tothese beneficial drug characteristics, its use in the anesthetic management of human beings and smallanimal patients is now routine. Anesthetic induction and maintenance with propofol in ponies was firstdescribed in the 1980’s [19]. Since that time it has also been evaluated for use in foals and adult horses.As with ketamine, it has generally been used in combination with alpha-2 agonists and/or musclerelaxants.
Propofol for Anesthesia in Foals - foals anesthetized with propofol (2 mg/kg) after premedication withxylazine (1.1 mg/kg) and butorphanol (0.01 mg/kg) were recorded as having higher heart rates and lowerblood pressures than foals induced with ketamine (2mg/kg) [20]. While surgical castration was performedsuccessfully with both drug protocols, the time to sternal recumbency and standing was shorter in foalsreceiving propofol; mean time to standing 12.3 minutes versus 19.7 minutes. In another study anesthesiawas maintained with an infusion of propofol (0.26 - 0.47 mg/kg/min) for non-invasive diagnosticprocedures after induction with propofol (2mg/kg) in foals premedicated with xylazine (0.5 mg/kg) [21].Quality of anesthetic induction, maintenance and recovery was satisfactory and foals took an average of 27minutes to stand following discontinuation of the infusion. Heart rate and mean blood pressure rangedfrom 84 - 92 beats per minute and 98 - 123 mm Hg, respectively. In foals breathing room air the PaCO2
ranged from 45 - 60 mm Hg and the PaCO2 ranged from 65 - 103 mm Hg.Propofol for Anesthesia in Horses (and Ponies) - recorded behavioral and cardiopulmonary characteristicsassociated with propofol in adult horses have varied. In otherwise unmedicated horses, the anestheticinduction quality was unpredictable and ranging from good to poor [22]. Surprisingly, behavioralcharacteristics were not significantly improved following premedication with xylazine (0.5 and 1.0mg/kg), detomidine (0.015 and 0.030 mg/kg), or medetomidine (7 µg/kg) [23,24]. However, with theaddition of guaifenesin to the anesthetic induction protocol, induction was rated as good to excellent [6].Although induction quality varied and differed from previous reports indicating good inductions withpropofol use in ponies and Brazilian horses [25], recovery quality was good to excellent with all protocols.Selected cardiopulmonary responses were recorded during xylazine/propofol and detomidine/propofolanesthesia [23]. Heart rate decreased after xylazine and detomidine administration and remained lowerthan pre-drug values during recumbency. The overall trend was toward a decrease in respiratory rate andincrease in PaCO2 during recumbency. The PaCO2 decreased significantly from pre-drug values duringrecumbency induced by both xylazine/propofol and detomidine/propofol.Similar findings (e.g. low heart rate, relative hypoxemia, etc.) were described during anestheticmaintenance with xylazine and low dose (0.15 mg/kg/min) propofol infusion [6]. Cardiac index wassimilar to that previously described for halothane anesthetized horses [6]. Anesthesia with high dose (0.25mg/kg/min) propofol infusion was characterized by marked respiratory depression and absence ofresponse to noxious stimulus. Despite the increased anesthetic depth, and likely the result of the indirectsympathomimetic effect of an increased arterial carbon dioxide tension, heart rate and cardiac index weremaintained within the normal ranges described for unanesthetized horses.Propofol/ketamine - the use of propofol and ketamine together for maintenance of anesthesia in poniesanesthetized for castration with detomidine/ketamine has also been evaluated [26]. Authors report verygood operating conditions and quiet recoveries from anesthesia following an average of ketamine (0.04mg/kg/min) and propofol (0.12 mg/kg/min).
Injectable Drugs as Part of a Balanced TechniqueThe purpose of a balanced anesthetic technique is to achieve all of the components of general anesthesiawhile minimizing the negative aspects of individual drugs on cardiopulmonary function. While thistechnique is commonly used in human beings and small animal patients, its use in the horse has beenlimited. Recent investigative work provides information that may facilitate increased clinical use of thistechnique in the anesthetic management of horses.Halothane/xylazine and halothane/detomidine - alpha-2 agents are known for their sedative and analgesicproperties in horses. It is therefore not unreasonable to expect that they would influence anestheticrequirements of concurrently administered drugs. Two reports with different alpha-2 agonist drugssubstantiate this.Steffey et al., report a 25 - 34% reduction in isoflurane dose requirement measured 40 to 60 minutesfollowing xylazine (0.5 mg/kg and 1.0 mg/kg, IV) administration to horses [27].Dunlop et al., demonstrated the halothane sparing effect of detomidine in horses [28]. Their reportindicates that halothane requirements decreased to approximately 55% from control as detomidine dose(and plasma concentration) increased.Halothane/ketamine and halothane/guaifenesin/ketamine - Muir et al., describe a reduction in halothanedose requirement and improvement in cardiovascular function with increasing plasma ketamineconcentrations [29]. While these aspects of combining these two drugs are favorable, authors describe apoor and prolonged recovery from anesthesia and suggest further clinical based evaluation of thistechnique. In a clinical study a combination of guaifenesin and ketamine was used to reduce the doserequirement of halothane in horses presented for diagnostic evaluation and emergency surgery [30]. Theyreport stable anesthetic conditions and predominantly good recoveries from anesthesia with this technique.
Halothane/lidocaine - another drug that has been evaluated for its effect on halothane minimum alveolarconcentration (MAC) for equine patients (ponies) is lidocaine [31]. Reduction in halothane MACcorrelated with increasing plasma lidocaine concentrations and ranged from 20 to 70%. Cardiopulmonaryeffects of this combination have not been fully evaluated.
Injectable Drugs as Modifiers of Recovery Following Inhalation AnesthesiaIn the 1980’s Rose et al., reported that recovery following isoflurane anesthesia in the adult horse wasunpredictable and less than ideal [32]. Because of continued observations of unpredictable recoveries frominhalation anesthesia, but predictably good recoveries following especially short and intermediate durationinjectable anesthesia, there is an interest in modulating recovery from inhaled agents by using injectabledrugs. Clinically this is commonly manifested by administration of an alpha-2 agonist early in the recoveryphase especially when using an inhaled anesthetic agent with a low blood gas solubility. The potentialbenefit is also supported by results of investigative efforts [33].Preliminary data suggest that recovery from isoflurane anesthesia can also be improved upon by use ofpropofol in the early recovery period; recovery quality was better with fewer attempts to stand in horsesthat received propofol [34].
SummaryInjectable anesthetic techniques for horses have improved and we now have many more choices availableto address needs in a variety of clinical circumstances. Despite these improvements however, deficienciesexist and the quest for the "ideal application" of injectable drugs for horses continues.
References1. Muir WW, Skarda RT, Milne DW. Evaluation of xylazine and ketamine hydrochloride for anesthesiain horses. Am J Vet Res 1977; 38:195-201. - PubMed -2. Matthews NS, Miller SM, Slater MR, et al. A comparison of xylazine-ketamine anddetomidine-ketamine anaesthesia in horses. J Vet Anaesth 1993; 20:68-72.3. Muir WW, Skarda RT, Sheehan W. Evaluation of xylazine, guaifenesin, and ketamine hydrochloridefor restraint in horses. Am J Vet Res 1978; 38:1274-1278. - PubMed -4. Greene SA, Thurmon JC, Tranquilli WJ, et al. Cardiopulmonary effects of continuous intravenousinfusion of guaifenesin, ketamine, and xylazine in ponies. Am J Vet Res 1986; 47:2364-2367. - PubMed -5. Young LE, Bartram DH, Diamond MJ, et al. Clinical evaluation of an infusion of xylazine, guaifenesinand ketamine for maintenance of anaesthesia in horses. Equine Vet J 1993; 25:115-119. - PubMed -6. Mama KR, Wagner AE, Steffey EP, et al. Evaluation of xylazine and ketamine for maintenance ofanesthesia in horses. In: Proceedings of the Ann Mtg Am Coll Vet Anes 1999; 18.7. Mama KR, Wagner AE, Steffey EP, et al. Behavioral response associated with xylazine and ketamineanesthesia in horses. In: Proceedings of the AVA Autumn Mtg 1999; 24.8. Mama KR, Pascoe PJ, Steffey EP, et al. Comparison of two techniques for total intravenous anesthesiain horses. Am J Vet Res 1998; 59:1292-1298. - PubMed -9. Singh K, McDonell WN, Young SS, et al. Cardiopulmonary and gastrointestinal motility effects ofxylazine/ketamine-induced anesthesia in horses previously treated with glycopyrrolate. Am J Vet Res1996; 57:1762-1770. - PubMed -10. Matthews NS, Hartsfield SM, Cornick JL, et al. A comparison of injectable anesthetic combinationsin horses. Vet Surg 1991; 20:268-273. - PubMed -11. Taylor PM, Luna SP, Sear JW, et al. Total intravenous anaesthesia in ponies using detomidine,ketamine and guaiphenesin: pharmacokinetics, cardiopulmonary and endocrine effects. Res Vet Sci 1995;59:17-23. - PubMed -12. Taylor PM, Kirby JJ, Shrimpton J, et al. Cardiovascular effects of surgical castration duringanaesthesia maintained with halothane or infusion of detomidine, ketamine and guaifenesin in ponies.Equine Vet J 1998; 30:304-309. - PubMed -13. Taylor PM, Luna SPL, Brearley JC, et al. Physiological effects of total intravenous surgicalanaesthesia using detomidine-guaiphenesin-ketamine in horses. J Vet Anaesth 1992; 19:24-31.14. Young LE. Clinical evaluation of romifidine/ketamine/halothane anaesthesia in horses. J Vet Anaesth1992; 19:89.15. Marntell S, Nyman G. Prolonging dissociative anaesthesia in horses with a repeated bolus injection. JVet Anaesth 1996; 23:64-69. 16. McMurphy RM, Young LE, Marlin DJ, et al. Comparison of the cardiopulmonary effects of totalintravenous anesthesia with romifidine, guaiphenesin, and ketamine vs halothane in horses. In:Proceedings of the Ann Mtg Am Coll Vet Anes 1998; 13.
17. Brock N, Hildebrand SV. A comparison of xylazine-diazepam-ketamine andxylazine-guaifenesin-ketamine in equine anesthesia. Vet Surg 1990; 19:468-474. - PubMed -18. Bettschart-Wolfsenberger R, Taylor PM, Sear JW, et al. Physiologic effects of anesthesia induced andmaintained by intravenous administration of climazolam-ketamine combination in ponies premedicatedwith acepromazine and xylazine. Am J Vet Res 1996; 57:1472-1477. - PubMed -19. Nolan AM, Chambers JP. The use of propofol as an induction agent after detomidine premedicationin ponies. J Assoc Vet Anaesth 1989; 16:30-32.20. Donaldson LL, Dunlop GS, Cooper WL. A comparison of propofol with ketamine after xylazine andbutorphanol as field anesthesia for young foals. In: Proceedings of the Ann Mtg Am Coll Vet Anes 1998;11.21. Matthews NS, Chaffin MK, Hartsfield SM. Propofol immobilization of neonatal foals. In:Proceedings of the Ann Mtg Am Coll Vet Anes 1993; 12.22. Mama KR, Steffey EP, Pascoe PJ. Evaluation of propofol as a general anesthetic for horses. Vet Surg1995; 24:188-194. - PubMed -23. Mama KR, Steffey EP, Pascoe PJ. Evaluation of propofol for general anesthesia in premedicatedhorses. Am J Vet Res 1996; 57:512-516. - PubMed -24. Bettschart-Wolfsenberger R, Freeman S, Bettschart RW, et al. Assesment of medetomidine/propofoltotal intravenous anaesthesia (TIVA) for clinical anaesthesia in equidae. In: Proceedings of the AVASpring Mtg 2000.25. Aguiar AJA, Hussni CA, Luna SPL, et al. Propofol compared with propofol/guaiphenesin afterdetomidine premedication for equine surgery. J Vet Anaesth 1993; 20:26-28.26. Flaherty D, Reid J, Welsh E et al. A pharmacodynamic study of propofol or propofol and ketamineinfusions in ponies undergoing surgery. Res Vet Sci 1997; 62:179-184. - PubMed -27. Steffey EP, Pascoe PJ, Woliner MJ, et al. Effects of xylazine hydrochloride during isoflurane-inducedanesthesia in horses. Am J Vet Res 2000; 61:1225-1231. - PubMed -28. Dunlop CI, Daunt DA, Chapman PL, et al. The anesthetic potency of 3 steady-state plasma levels ofdetomidine in halothane anesthetized horses. In: Proceedings of the 4th ICVA 1991; 7.29. Muir WW, Sams R. Effects of ketamine infusion on halothane minimal alveolar concentration inhorses. Am J Vet Res 1992; 53:1802-1806. - PubMed -30. Spadavecchia C, Stucki F, Schatzmann U. Ketamine-guaiphenesin infusion to maintain generalanaesthesia in horses receiving halothane in subanaesthetic dose: a clinical study. In: Proceedings of theAVA Autumn mtg 1999; 23.31. Doherty TJ, Frazier DL. Effect of intravenous lidocaine on halothane minimum alveolar concentrationin ponies. Equine Vet J 1998; 30:300-303. - PubMed -32. Rose JA, Rose EA. Clinical experience with isoflurane anesthesia in foals and adult horses. In:Proceedings of the AAEP 1988; 555-561.33. Carroll GL, Hooper RN, Rains CB, et al. Maintenance of anaesthesia with sevoflurane and oxygen inmechanically-ventilated horses subjected to exploratory laparotomy treated with intra- and post operativeanaesthetic adjuncts. Equine Vet J 1998; 30:402-407. - PubMed -34. Mama KR, Steffey EP, Pascoe PJ. A preliminary study comparing anesthetic recovery in horsesfollowing isoflurane of isoflurane propofol. In: Proceedings of the Ann Mtg Am Coll Vet Anes 1995; 13.
All rights reserved. This document is available on-line at www.ivis.org. Document No. A0604.1000 .
In: Recent Advances in Anesthetic Management of Large Domestic Animals, E.P. Steffey (Ed.) Publisher: International Veterinary Information Service (www.ivis.org), Ithaca, New York, USA. Anesthetic Management of the Horse: Inhalation Anesthesia (31-Jan-2003) P. W. Kronen
Division of Anaesthesiology, Department of Clinical Veterinary Medicine, University of Berne, Berne, Switzerland.
Introduction Management of intermediate and long-term (more than 30 - 60 minutes) anesthetic procedures most commonly involves the use of inhalational anesthetic methods in many species. Over the last 40 years, this has become true also for equine anesthesia. The delivery of anesthetic drugs via the lungs offers some advantages - in part caused by the pharmacokinetics of the volatile agents in clinical practice. Among these advantages, the relatively quick and easy adjustment of anesthetic depth ranks high. Rapid changes in anesthetic delivery and tissue concentrations of volatile anesthetics allow for the anesthetist to quickly react to alterations in organ system functions (central nervous, cardiovascular, and respiratory) and thereby control functionality within physiologic limits. This is even more important when taking clinical conditions and diseased patients into consideration. Furthermore, inhalation anesthesia with the modern volatile anesthetics may produce a relatively short recovery period, a factor of growing interest in the literature of equine anesthesia. Volatile anesthetics are potent drugs with a relatively low margin of safety (therapeutic index: 2 - 4). They must be used carefully as overdosing may result in severe depression of the cardiopulmonary, central nervous, and other system functions and lead to death. In the past, several surveys on morbidity and mortality rates related to anesthesia - even if not specific to horses - suggested that a major percentage of anesthetic deaths were avoidable [1,2]. Fortunately, a newer and equine specific study (CEPEF-1) does not show similar results [3], but unwanted effects - even though quantitatively reduced with halothane, isoflurane, desflurane, and sevoflurane - can still not be excluded. Therefore, understanding of principles and concepts of inhalational anesthesia in general is crucial to the employment of volatile anesthetics. These topics are considered basic to this chapter and are discussed elsewhere [4-12]. The scope of this chapter is, instead, to provide a focused review of selected topics, namely interactions with carbon dioxide absorbers, mechanism of action of inhaled anesthetics, and use of sevoflurane and desflurane in the equine patient.
Reactions with CO2-absorbers All volatile anesthetics react with the carbon dioxide absorbers currently used or tested. The degree of reaction varies greatly though, and significant amounts of research have been put forth in order to assess and minimize such reactions. Three types of reactions are of possible concern: Firstly, absorption of volatile anesthetics to components of the carbon dioxide absorbers (silica, polyvinylpyrrolidine; Table 3) may occur. This plays a minor role in the decreased anesthetic concentration leaving the CO2-absorber. Secondly, volatile agents containing a difluoromethoxy moiety (CHF2, Table 1) may react with the absorber to form carbon monoxide (CO) [13]. With classic soda lime (Table 3), the amount of CO produced decreases in the order: desflurane > enflurane > isoflurane [14]. The clinical significance of such carbon monoxide production is probably low in equine anesthesia, given the large volume of the anesthetic circuit and the high fresh gas flow rates that are commonly used. Even though halothane does not present the above mentioned difluoromethoxy component, Dodam et al., [15] found the carbon monoxide concentration in horse circle rebreathing systems to be higher with halothane (highest at 90 min with 54 + 33 ppm [parts per million]) than with isoflurane (highest at 90 min with 21 + 18 ppm). However, this group as well concluded that the detected concentrations were of no clinical significance with either agent. Also, trifluoromethane, a breakdown product of desflurane (and isoflurane) may react with carbon dioxide absorbers to form CO [7]. Thirdly, halothane and more vigorously sevoflurane may react with carbon dioxide absorbers and be degraded to haloalkenes. These substances, mainly BCDFE (2-bromo-2-chloro-1,1-difluoroethylene) from halothane and Compound A (FDVE, fluoromethyl-2,2-difluoro-1-(trifluoromethyl) vinyl ether) from sevoflurane degradation have been demonstrated to exert nephrotoxic effects in rats [16]. Thresholds for nephrotoxicity for compound A in rats are generally accepted to be within clinically relevant concentrations [17-19]. Other than in rats, thresholds for nephrotoxicity due to compound A have been demonstrated only in non-human primates and lay outside the range of clinically produced exposure [20]. Even though such nephrotoxicity has not been shown in other species to date, the potential for it and the lack of exclusion represents a
possible concern and further investigation in different species is required. However, carbon dioxide absorbers break down sevoflurane to Compound A with decreasing quantities: barium-OH lime > sodium-OH lime > KOH-free sodium-OH lime > calcium-OH lime. Both, carbon monoxide and haloalkene exposure is dependent on the presence of strong alkali (monovalent hydroxides: KOH, NaOH) in the CO2-absorber, the fresh gas flow as well as the degree of desiccation and the absorbent temperature [13,14,21]. Absorbers not containing strong alkali degradate halothane and sevoflurane to significantly lesser degrees [13,22,23], resulting in significantly lower haloalkane production (about 10% of that of other absorbers). Clinically, it is impossible to detect decreases in absorber humidity and therefore regular exchange of the absorber in the anesthetic circuit is warranted, even when unused. Several attempts have been made to develop methods for absorber cooling in order to decrease degradation of inhalants. Although cooling does effectively reduce the formation of carbon monoxide and haloalkenes in vitro [24,25], and the addition of dead space reduces absorber temperature in vivo [26], none of these methods has found a clinical application.
Mechanism of Action The mechanism of action of inhaled anesthetics remains undefined. However, a vast number of recent publications highlight the research effort put forth in an attempt to bring light to this particular topic. A review of the current literature is therefore included here.
Actions on Central Nervous System (Macroscopic) - Inhalant anesthetics act differently at different sites of the central nervous system (CNS). The reticular formation plays an important role in regulation of consciousness and motor activity, and therefore this brainstem region has often been proposed as an important part of anesthetic action of inhalant anesthetics. Generally, inhalant anesthetics cause a decrease in brain activity, but such a decrease in tone at the level of the reticular formation as mechanism for general anesthesia, today, is regarded as an oversimplification. In fact, anesthetics can cause decreases, but also increases or no changes in neuronal transmission, depending on the agent and the anatomical location. Furthermore, there is evidence for alterations in neuronal activity of the cerebral cortex and hippocampus [27], as well as the transmission from the thalamus to certain cortical regions under the influence of inhalant anesthetics [28]. Also at the level of the spinal cord, inhalant anesthetics cause changes in excitatory and inhibitory function of neuronal transmission. More precisely, they alter the dorsal horn mediated responses to noxious and non-noxious stimuli, decrease the activity of spinal motor neurons and decrease tonic descending input. To date, there is no evidence of peripheral receptor depression involved in the action of inhaled anesthetics. Historically, both main effects of general anesthesia, amnesia and immobility, were thought to be caused by the same mechanism - only at different concentrations, ie., amnesia occurring at lower and immobility at higher concentrations of an anesthetic drug. Today, there is evidence that these two main actions are mediated through different sites of action [29-31]: When comparing different anesthetics in their ratio between concentrations needed to produce amnesia and immobility, respectively, they differ markedly among different drugs, suggesting two different mechanisms or sites of action. Such differential activity has been found also in the horse [32]. Furthermore, the existence of so-called non-immobilizing drugs, ie., drugs that produce amnesia, but not immobility, implies that the site that mediates amnesia is not associated with the production of immobility. These non-immobilizers do not even reduce the requirements for conventional anesthetics to produce immobility (definition of MAC, Minimum Alveolar Concentration) [33]. Furthermore, functional separation of the spinal cord from the brain has been found not to decrease the anesthetic partial pressure required for immobility [34], suggesting that immobility is mediated at the level of the spinal cord [29].
Interruption of Neuronal Transmission (Microscopic) - The sensitivity of single neurons to the actions of anesthetics varies. Some are hypersensitive (inhibition of axonal firing at less than 1 MAC), some sensitive (inhibition at 1 MAC), and some are not sensitive. The understanding is further complicated as inhaled anesthetics may alter the neuronal transmission at several different levels. Axonal inhibition may be achieved at near-clinical concentrations, and even an incomplete inhibition of action potentials may result in a decreased neurotransmitter release at the subsequent synapsis. The degree to which inhaled anesthetics alter axonal transmission seems to be dependent on the impulse frequency of the axon (low frequency transmission is blocked, while higher frequencies are not), the fiber diameter (inversely proportional), and the axonal region (branching points > axons). However, sypnapses are circa 5 times more sensitive to the inhibitory actions of inhaled anesthetics than axons [35]. Such synaptic inhibition can be on the presynaptic site, decreasing the amount of neurotransmitter release or decreasing the rate of re-uptake from the synaptic cleft, or at the postsynaptic site, altering the binding of neurotransmitter or influencing the ionic conductance changes that follow activation of the postsynaptic receptors. Both, pre- and postsynaptic effects have been found and inhalant anesthetics may increase, decrease or not affect presynaptic neurotransmitter release and the postsynaptic response [36]. The variation of effects depends on the single neurotransmitter, the anesthetic drug, and the anatomical location, but no single neurotransmitter concentration seems to allow for an explanation of the anesthetic state.
Site of Action (Molecular) - The differential activity of inhalant anesthetics at several macro- and microscopic locations does not preclude a unitary molecular site of action [36]. Historically this has led to the idea of a unitary theory of narcosis [37]. In fact, despite the wide spectrum of substances that cause anesthesia (inert gases, simple organic and inorganic molecules, haloalkenes, ethers, etc.), there are astonishing correlations between physical properties of the various inhalant anesthetics and their anesthetic activity. The best correlation can be found between anesthetic potency and lipid solubility and is termed Meyer-Overton rule after the two independent dicoverers (Hans Horst Meyer 1899 in Marburg, Germany and Charles Ernest Overton 1901 in Zurich, Switzerland) [38]. Accordingly, the product of the anesthetizing partial pressure and the olive oil/gas partition coefficient varies only very little over a 100,000 fold range of anesthetizing partial pressures. Thereby, anesthesia is produced when a sufficient number of molecules (independent of their type) occupy the respective hydrophobic regions in the central nervous system. Since the Meyer-Overton postulates, several apparent exceptions to their rule have been found. Enflurane and isoflurane are structural isomers (Table 1), have similar oil / gas partition coefficients (Table 2), and would therefore be expected to show similar anesthetic potency, but the MAC for isoflurane is only about 62% of that for enflurane (in the horse, Table 4). Secondly, complete halogenation of the end-methyl groups on alkanes and ethers results in decreased anesthetic potency and increased convulsant activities of the compound, despite the respective increase in lipid solubility. Thirdly, the higher n-alkanes in a homologous series do not follow the Meyer-Overton rule [36]. Lastly, the lipophilicity of non-immobilizers would indicate that they produce immobility, but they do not [29]. In 1954, LJ Mullins hypothesized on the basis of the Meyer-Overton rule a possible molecular mechanism of anesthesia, the critical volume hypothesis [39]. Thereby, anesthetic molecules would be absorpted into the lipid bilayer of the cell membrane, cause a volume expansion beyond a critical volume, and obstruct ion channels or change electrical conductance of neurons. In fact, a volume expansion of membranes was found later on, as well as a reversal of anesthetic state with increases in hydrostatic pressure [40]. On the other hand, the increases of anesthetic requirements seen with increased temperature (and consequent increase in membrane volume) clearly contrast this hypothesis, as well as the fact that not all lipid soluble agents produce anesthesia and the non-linearity of the pressure reversal curve for some anesthetics. To date, the critical volume hypothesis is regarded as an oversimplification of the anesthetic state [36]. On the basis of the fact that inhalant anesthetics interrupt neuronal transmission and because this transmission occurs as an ion movement at the level of the neuronal membrane, the latter is commonly thought to be the primary site of action. Possible molecular sites of such membrane interference would be in the non-polar interior of the phospholipid bilayer, in hydrophobic pockets in proteins embedded or outside this bilayer, or at the interface between lipophilic sites and intrinsic membrane proteins. Several theories exist regarding changes in the neuronal membrane conformation (alteration in membrane dimension or physical state) as an attempt to explain anesthetic action, but to date most authors agree that ultimately the action of inhalant anesthetics is on neuronal membrane proteins that permit ion fluxes during membrane excitation [29,30,36,37,41-45]. Nevertheless, it remains unknown whether anesthetic molecules act primarily through an indirect alteration in surrounding lipids or via a second messenger system or directly by binding on channel proteins. Possibly, immobility is mediated by the binding to specific channel proteins (GABA A [46], glutamate [45], nicotinic acetylcholine [47] receptors), while amnesia may present with a different, unspecific, underlying molecular mechanism [29].
Sevoflurane History - B M Regan while working for Travenol Laboratories synthesized sevoflurane in 1968. His findings, though, were not published until 1971 [48] and apparent toxic effects (which later were found to be consequences of flawed study design) impeded further development of the compound [49]. Because at that time sevoflurane did not seem promising for release into practice, the rights for this compound first went from Baxter-Travenol to Anaquest (Ohmeda / BOC) and then to Maruishi in Japan for human anesthesia. The Japanese company continued research and development and released sevoflurane in 1990 in Japan. Within three years an estimated one million people received sevoflurane. Abbott subquently obtained the rights to sevoflurane, continued its development, and finally released sevoflurane in the United States in 1994. The first report of sevoflurane use in horses was published in 1994 by Aida et al., in Japan [50].
Physico-chemical Properties and Recovery Characteristics - Some physico-chemical properties of sevoflurane are included in Table 1. Sevoflurane is structurally related to isoflurane and enflurane (Table 1) and consequently shares many of the physico-chemical properties with these agents. The vapor pressure of this inhalant permits use of conventional, temperature-compensated vaporizer technology and, in fact, the vaporizers commercially available are similar to the ones for halothane and isoflurane. Sevoflurane is unstable in most carbon dioxide absorbers, resulting in the production of several compounds. The most prominent of these, Compound A, is a haloalkene with potential for nephrotoxicity (see paragraph on carbon dioxide absorbers). Carbon monoxide production from sevoflurane interaction with carbon dioxide absorbers is not significant [51]. The blood/gas partition coefficient, a measurement of solubility in this particular solvent, of sevoflurane (0.69, see Table 2) is significantly lower than that for halothane or isoflurane. Therefore, other conditions being equal, one would expect anesthetic induction, recovery, and intraoperative modulation of anesthetic depths to be notably faster than
with the other mentioned agents. This potential advantage over older compounds has been confirmed in a number of studies indicating a fast and smooth recovery from sevoflurane anesthesia in adult horses and humans [52-58]. In children, however, several reports describe more postanesthetic agitation with sevoflurane than with halothane [59-61], although Read et al., [62] found no difference in induction and recovery characteristics between isoflurane and sevoflurane when used as the sole anesthetic in foals.
Central Nervous effects - Sevoflurane is less potent than halothane or isoflurane, but more potent than desflurane, nitrous oxide or xenon, as reflected by MAC (Table 4). Sevoflurane, like other volatile anesthetics, produces a dose-related, generalized depression of the central nervous system, as reflected by burst suppression on the EEG, but isoelectric patterns seem to require concentrations exceeding 2 MAC (in dogs and rabbits) [11]. Also, the bispectral index (BIS), a numerical value derived from the EEG to assess CNS depression (not fully validated in Veterinary Medicine), seems to correlate well with delivered sevoflurane doses in dogs [63]. As other volatile anesthetics, sevoflurane causes dose-related decreases in cerebral vascular resistance and metabolic rate [64,65]. It may therefore increase cerebral blood flow (to a lesser degree than halothane [66]) and intracranial pressure to a similar degree as isoflurane, even though the latter can be prevented by hypocapnia [67,68]. In fact, cerebral autoregulation is maintained during sevoflurane anesthesia [69]. Sevoflurane is not seizuregenic [49,65].
Cardiovascular Effects - Sevoflurane induces dose-dependent cardiovascular depression to a degree similar to that of isoflurane, except for an inconsistantly reported positive chronotropic effect [11,52,54,57,70-75]. Much of this information derives from experiments with other species but in horses too, sevoflurane decreases cardiac output, systemic vascular resistance, arterial blood pressure and mean pulmonary artery pressure [76]. However, these effects are also affected by anesthetic duration, ie., arterial blood pressure may increase with anesthesia time [77], as has been reported for halothane [78]. Sevoflurane does not cause arrhythmias of the heart as halothane and the arrhythmogenic epinephrine dose in dogs is similar to that for isoflurane [79].
Respiratory Effects - Sevoflurane induces a dose-dependent respiratory depression to a similar degree as isoflurane [6,80]. Both agents, sevoflurane and isoflurane, cause greater increases in PaCO2, decreases in pH and ventilatory response to hypercapnia than does halothane in horses [71,81]. Respiratory rate is lower than with halothane, and minute ventilation decreases [50,52,73,74,82].
Biotransformation - Sevoflurane is metabolized to a moderate extent (5%, Table 5). Very little amounts of the drug is probably lost percutaneously, via surgical incisions, in the urine and feces [83], and the remainder of the total administered dose is exhaled unchanged (as for the other volatile agents). Most published data reflect findings from humans or laboratory species. It is unknown whether or not there are substantial differences in volatile drug metabolism in the horse. However, it is known that the anatomical site for sevoflurane metabolism is the endoplasmatic reticulum (ER) of hepatocytes [84]. More specifically, as for other volatile anesthetics, the cytochrome P450 enzyme system represents the major metabolic pathway [85,86]. The 2E1-isoform of cytochrome P450 catalyzes sevoflurane oxidative metabolism to inorganic fluoride (F) and hexafluoroisopropanolol (HFIP, which is then glucuronidated and eliminated via the kidney) in a ratio of 1:1 [87,88]. This metabolic reaction is dose-dependent (MAC-hours) [70,89]. Enzyme induction by pretreatment with phenobarbital [90], phenytoin [91], isoniazid [92], and chronic ethanol administration [93] may enhance sevoflurane defluorination.
Hepatic Effects - Little information is available about direct hepatic effects of sevoflurane. However, splanchnic circulation and with it portal and hepatic arterial blood flow suffers only little from a generalized cardiovascular depression that is assumed to be similar to isoflurane [94]. Despite potential hepatotoxicity of HFIP, fulminant hepatic failure or hepatic necrosis have not been reported with the use of sevoflurane, probably because HFIP is glucuronidated so quickly that it cannot exert toxic action [80,95]. However, hepatic dysfunction as measured, increased serum enzyme levels after sevoflurane occasionally has been suspected in human patients [96,97]. Controlled, prospective studies in humans, on the other hand, did not show significant potential of sevoflurane to produce liver dysfunction [98,99], a result confirmed in one study in horses as well [100].
Renal Effects - Two sevoflurane breakdown products are of potential concern because of their nephrotoxicity: Compound A and inorganic fluoride. The first results mainly from sevoflurane reaction with desiccated, warm (> 40ºC) carbon dioxide absorbers containing strong alkali (baralyme > sodalyme). It has been demonstrated to cause renal tubular necrosis in Fischer 344 rats when at concentrations exceeding 50 ppm for three hours [19], which has led the US Food and Drug
Administration (FDA) to recommend the use in human patients for not more than 2 MAC-hours of low-flow anesthesia (at 1L/min fresh gas flow, as of December 1997). In the countries of the European Union, there is no restriction regarding the applied fresh gas flows for management of human patients. Concentrations necessary to produce severe renal injury are inversely related to duration of anesthesia in rats [101]. However, several studies by Bito and Ikeda [102-104] using sodium- and barium hydroxide lime have shown no toxic effects attributable to compound A even after prolonged low-flow anesthesia (up to 18.6 MAC-hours) and the highest compound A concentrations measured in these studies were 30 ppm, 46.5 ppm, and 60.78 ppm, respectively. Goeters et al., [105] found compound A concentrations of up to 57 ppm after two hours of minimal-flow anesthesia (0.5 L/min), but no detectable changes in renal or hepatic function. Conversely, Eger et al., [106] found compound A concentrations of up to 56 ppm after 10 MAC-hours of sevoflurane anesthesia at 2 L/min fresh gas flow. In this study, sevoflurane administration was associated with transient injury to the glomerulus (albuminuria), the proximal tubule (glucosuria, increased urinary alpha-GST), and the distal tubule (increased urinary gamma-GST). No clinical studies of humans demonstrate significant changes in BUN, creatinine, or the ability to concentrate urine after sevoflurane anesthesia when compared to other inhalant anesthetics. This is true also for a study in horses [100]. Inorganic fluoride is another metabolic breakdown product from sevoflurane (table 5), and serum fluoride levels are increased after sevoflurane anesthesia in humans [11,86,95], horses [82,100], and other species [18]. Its nephrotoxicity has not been shown, even at elevated serum concentrations. Nephrotoxicity of increased serum fluoride concentrations seems to be related only to methoxyflurane and, to a lesser degree, to enflurane. Kharasch et al., [107] hypothesized this to be related to the relative lack of intrarenal cytochrome P450 2E1 production of fluoride ions with sevoflurane when compared to methoxyflurane and enflurane. Nephrotoxicity from sevoflurane increased serum fluoride levels is therefore probably not a clinical problem.
Effects on Skeletal Muscles - Sevoflurane produces skeletal muscle relaxation that is comparable to that of isoflurane and enhances neuromuscular block to a similar degree as isoflurane [108,109]. Sevoflurane, as other inhalant anesthetics, can trigger malignant hyperthermia in animal [110] and human patients [111].
Other Effects - Sevoflurane decreases capillary filtration coefficients in the microvascular bed, thereby decreasing the extravasation of fluids into the interstitial space in human patients [112].
Desflurane History - In the 1960’s, RC Terrell at Ohio Medical Products (later Anaquest, today Ohmeda/BOC) synthesized some 700 compounds in the search for a better inhalant anesthetic [113]. Enflurane, introduced into clinical practice in 1973 was compound number I-347 in this series. Isoflurane, its stereoisomer, released in 1981 (compound number I-469), as well as desflurane (compound I-653) were also synthesized in that series. The latter was released only in 1992 as it was initially produced with a hazardous method of synthesis involving elemental fluorine. Almost twenty years passed before a less explosive method involving hydrogen fluoride and antimony pentachloride was developed for the synthesis of desflurane [114].
Physico-chemical Properties and Recovery Characteristics - Desflurane's vapor pressure is the highest among the volatile anesthetics in clinical use and close to normal atmospheric pressure (Table 1). In fact, desflurane boils at room temperature (22.8ºC) and, hence, confers special technical problems for its vaporization. Currently, the only vaporizer that produces controllable and predictable concentrations of desflurane is electronically controlled and therefore requires electricity. For a detailed technical description the reader is referred to other literature [114,115] or the manufacturers’ website (Datex Ohmeda - Product Portfolio - Tec 6 Plus Vaporizer). The blood/gas partition coefficient for desflurane is very close to that of nitrous oxide (Table 2) and consequently modulation of anesthetic depth should be achieved quickly, and recovery from anesthesia fast [6,114]. Clinical data from human patients seem to confirm this contention [116-118], for example Eger et al., [119] found recovery from desflurane anesthesia to proceed nearly twice as fast than with sevoflurane. The few studies published that mention recovery from desflurane show analogous results for equine use [120]. Horses’ recovery from desflurane anesthesia is fast (for example 15 min to standing after 100 minutes of anesthesia), and subjectively rated good to excellent [121]. Desflurane is stable in sodium hydroxide lime unless the latter is dry and temperatures high (60ºC) [122], when desflurane is broken down to produce significant amounts of CO [51].
Central Nervous Effects - Desflurane is the least potent among the volatile anesthetics in clinical use (Table 4), and only the gaseous anesthetics have higher MAC values. This confers a notable decrease in inspired oxygen concentration, for example: at 2 MAC (a dose commonly used at the beginning of a clinical anesthetic) the delivered concentration of desflurane lies in the range of 16% (for isoflurane circa 2.6%). Consequently, carrier gas (oxygen) concentration cannot be
higher than 84% (97.4% for isoflurane), and inspired oxygen is decreased by 13.4% with respect to isoflurane anesthesia. This may result in a decrease of arterial oxygen partial pressure (PaO2) in the range of 54 - 67 mmHg compared to similar isoflurane anesthesia, a dramatic reduction not always tolerable-particularly in equine clinics. The changes in EEG seen with desflurane anesthesia are similar to those found with isoflurane [123] and are probably the ones associated with anesthesia [80]. Electrical silence is not produced until 1.7 MAC is achieved [123]. Seizuregenicity is not reported with desflurane use [6]. Desflurane, as sevoflurane, can decrease cerebral vascular resistance and cerebral metabolic oxygen requirements and increase intracranial pressure in a dose-dependent fashion [11,80,114]. This has led to the recommendation to use desflurane with caution in patients with decreased intracranial compliance [6,7]. Cerebrovascular autoregulation in response to carbon dioxide is well maintained as with isoflurane [124]. Jones et al. reported desflurane to exert good analgesic effects in horses [125].
Cardiovascular Effects - The circulatory effects of desflurane parallel those of isoflurane [72]. Desflurane decreases blood pressure by decreasing systemic vascular resistance, but tend to preserve cardiac output at clinically used doses [126]. It can, however depress myocardial contractility [126,127]. Desflurane consistently causes increases in heart rate more than the other volatile agents [128]. Studies in horses confirm this effect [129-131]. Chronotropic effects are accentuated by sudden changes in anesthetic delivery, such as induction of anesthesia [114]. Such increases in heart rate may well be caused by sympathetic stimulation and are blunted by administration of opioid or alpha-2 agonist drugs [80]. Desflurane does not cause in itself or predisposes the heart to epinephrine-induced arrythmias [132].
Respiratory Effects - As the other inhaled anesthetics, desflurane causes dose-dependent respiratory depression. The magnitude in horses seems to parallel or exceed that of isoflurane [72,125,129,130], and is expressed in drastic decreases of respiratory rate, but tidal volume decreases as well, once 1.5 - 2 MAC are reached. In humans, desflurane causes airway irritation with resulting coughing, secretions and breath holding [11,133].
Biotransformation - Only very small amounts of desflurane are metabolized (0.02%, Table 5) [134]. Consequently, in humans [134], rats [135], and pigs [136] little or no increases in serum or urine inorganic or organic fluoride levels has been demonstrated and the trifluoroacetate levels found are only 1/5 - 1/10 of those produce by isoflurane metabolism [11].
Hepatic Effects - As predictable by the minimal biodegradation, the sustained cardiac output and the rapid elimination after anesthesia, desflurane affects liver function minimally or not at all [137]. Furthermore, desflurane seems not to worsen pre-existing liver disease [138]. Studies about hepatic blood flow in swine and dogs have not shown significant decreases in total hepatic blood flow (portal and hepatic arterial), and there is evidence of decreases in portal vascular resistance in normotensive and hypotensive pigs under desflurane anesthesia [139,140]. To assess hepatocellular function in desflurane exposed human patients, Schmidt et al., [141], measured the centrilobularly sensitive alpha glutathione S-transferase and found no changes. Conversely, Steffey et al., found mild, transient increases in aspartate aminotransferase and sorbitol dehydrogenase in horses after desflurane anesthesia, but judged these alterations as clinically unremarkable (as for sevoflurane) [100].
Renal Effects - As for hepatic function, desflurane only minimally affects renal function [142]. This has proved true for human patients [137], rats [143], and dogs [144] and the study done by Steffey et al., suggests similar findings in horses [100]. Both, renal function and blood flow seem unaffected. Consequently, even in patients (human) with pre-existing disease, no worsening of renal function could be detected [138].
Effects on Skeletal Muscles - As the other volatile agents, desflurane causes muscle relaxation, enhances action of neuromuscular blocking agents and may trigger malignant hyperthermia [145,146].
Data from: [4-6,49,70,80,113,114,147-151].
Table 1. Some Physicochemical Properties of Inhalational Anesthetics Used in Veterinary Medicine.
Data from: [5,83-85,95,107,159-161] * : Degree of metabolism includes estimates from recovery of metabolites and estimates from recovery of the unchanged drug ¤ : Smaller fonts in italics indicate reductive metabolism
References
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Degree of Metabolism * (%) Mechanism of Metabolism Principal Metabolites
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In: Recent Advances in Anesthetic Management of Large Domestic Animals, SteffeyE.P. (Ed.)Publisher: International Veterinary Information Service (www.ivis.org)
Anesthetic Management of Donkeys and Mules (15 July 2000)
N.S. Matthews and T.S. TaylorDepartment of Veterinary Small Animall Medicine and Surgery, The Texas Veterinary Medical Center, TexasA&M University, College Station, Texas, USA. IntroductionDonkeys and mules are easily recognized as looking different from horses. However, most of us seem to havedifficulty in recognizing the many other differences, which exist between horses, donkeys and mules. A littlebackground may help in identification and appreciation for how they are different from horses. The donkey orburro (Equus asinus) has been associated with mankind throughout recorded history. It is still widely used(recent estimates put their number at 44 million) in many parts of the world for work and transport of all kindsof goods [1]. Donkeys are desert-adapted animals, who survive where horses cannot. Reasons which havebeen advanced for their hardiness and survival include: desert-adaptations to water shortages, the ability torehydrate quickly when water is presented, greater variability in thermoregulation to reduce stress fromvariation in ambient temperature, willingness to eat feeds unpalatable to horses, and perhaps, differences insusceptibility to diseases highly fatal to horses [2,3]. Physiologically, donkeys are known to have differentfluid-balance and partitioning of fluids than does the horse [4]. This affects the way they distribute drugs,including anesthetics. Based on what we know about the pharmacokinetics of some drugs in donkeys, webelieve that they may also metabolize some drugs faster than horses, which affects anesthetic drug duration.Behaviorally, the donkey is very different from the horse [5]. Donkeys do not seem to share the horse’simmediate flight response; they will often face a frightening object and freeze (hence their reputation forstubbornness). The best advice (usually ignored) is to plan to take more time in first introducing the animal tonew experiences. Behavioral differences are also seen in their responses to injections, twitches, leading, andother common procedures related to anesthesia. If you are not familiar with these differences, it is wellworthwhile to enlist the help of an experienced donkey or mule person. If no such person is available, and youhave a choice between an experienced horse person and experienced cattle person, choose the one who hasworked with cattle! Be aware of the fact that both donkeys and mules are extremely intelligent and kickwithout warning with excellent aim! Donkeys and mules are very trainable and have excellent memories forwhat they have learned (both good and bad!). By nature, their disposition is quite sedate; therefore, recoveryfrom anesthesia and surgery is almost always smooth and without excitement.Physiologically, mules (the hybrid of Equus asinus and Equus caballus) seem to be more like the horse,although not identical. Therefore, they may range in appearance and temperament depending on the type ofhorse used in the breeding (e.g., Thoroughbred vs. Percheron). They also look more horse-like and muleowners will be insulted if you call their mule a donkey (as will donkey owners if you call their donkey amule)! Donkeys and mules are differentiated by size and range from miniature (less than 85 - 90 cm) tostandard (90 to 135 cm) to Mammoth (donkeys >135 cm) or Draft (mules). In the United States, theregistering organization for all types is the American Donkey and Mule Society (2901 N. Elm St., Denton, TX76201, USA) which also serves as the parent organization for all local clubs.As with other Equidae species, there is a wide variation in behavior depending on how accustomed to peoplethe animal is (feral versus domesticated), and how much training and handling it has had. In our experience,feral or unhandled animals will require much higher drug doses to achieve the same results (estimated 1.5 to 2times the usual dose), and if drugs must be administered by the intramuscular route (instead of intravenous
route), dosages are higher yet (estimated 3 times the usual dose).
Preanesthetic EvaluationIn performing a preanesthetic evaluation of the patient, it is important to realize that there are differencesbetween donkeys, mules and horses. Donkeys will only appear sick when they are very significantly ill. Theuse of a hematocrit to evaluate the degree of dehydration may be used, however it is important to recognizethat donkeys can dehydrate significantly (12 - 15%) before hematocrit will increase [2]. Normal hematologicand biochemical values are reported for donkeys [6], and there are differences from normal horse values.References for mules are very limited and old enough that they may not represent current diagnostictechniques. Baseline values for heart and respiratory rate are also slightly different from horses. Heart ratesmay range from 35 - 55 bpm (depending on degree of fitness and anxiety), while respiratory rates are higher atrest; 20 - 35 bpm and will vary with ambient temperature (respiratory rate increases to reduce the amount ofwater expended for evaporative cooling). Daily body temperature may vary by 3 degrees C [4].
Restraint and Injection TechniquesGenerally, donkeys and mules don’t appreciate needles at all! Use of a twitch appears to be much lesseffective than in horses. A donkey will usually stand still if it is tied with the rope very short to a stout objectwhich it has already determined it can not budge (so it is advantageous to give them some time to accept beingtied before trying to proceed. Donkeys may also be more safely restrained in a chute or with a squeeze gatethan would a horse (they are more like cattle in their response to restraint; accepting it rather thanunreasonably fighting restraint). Tying up one leg or hobbles may also be effective for restraint; once theanimal has determined that it is restrained: it will usually not continue to fight the rope.The jugular vein lies in the same anatomic location as in the horse, but donkey skin is thicker than horse skin.Therefore, it is necessary to change the angle of needle placement slightly when attempting venipuncture fordrug administration or catheter placement; angle deeper (i.e., more perpendicular to the skin). It also seemsbetter to lay the needle on the skin and insert slowly by increasing the pressure gradually, instead of quickly"slapping" the needle through as you would in the horse. Often, the mule or donkey will lean into the needle.
PreanestheticsAlthough the pharmacokinetics of xylazine have not been researched in mules, they appear to requireapproximately 50% more xylazine (1.6 mg/kg IV) or detomidine (0.03 mg/kg IV) to produce good sedationthan most donkeys or horses. However for unhandled or feral donkeys and miniature donkeys, the higherdoses should also be used. Injectable anesthesia is most satisfactory when butorphanol (0.04 mg/kg IV) ordiazepam (0.03 mg/kg IV) are combined with xylazine or detomidine to increase the sedation produced.Acepromazine (0.04 mg/kg IV) has also been used satisfactorily for tranquilization of donkeys and mules.
Injectable AnesthesiaKetamine (2.0 - 3.0 mg/kg IV) can be used in donkeys and mules for short procedures, following sedation (asabove). The half-life of ketamine is shorter than in the horse, so it may be necessary to administer additionaldoses [7] however, increasing ketamine above 3.3 mg/kg has been associated with rough recoveries [8]. Theuse of a local anesthetic in combination with injectable anesthesia will reduce the need to redose as frequently.Prolonging anesthesia with the combination of guaifenesin/ketamine/xylazine (often called G/K/X or "tripledrip") may be used, but careful monitoring is necessary. Donkeys have a lower tolerance for guaifenesin; theyrequire approx. 40% less to produce recumbency than do horses. However, we have used the combination ofguaifenesin and thiopental for induction of anesthesia in donkeys and mules; the combination is administered"to effect" with careful monitoring to prevent excessive depth of anesthesia.Xylazine premedication followed by Telazol (1.0 mg/kg IV) is effective for producing anesthesia in donkeysand mules. Recumbency time is longer than xylazine-ketamine combinations. Recoveries are satisfactory indonkeys, but we have observed occasional rough recoveries in mules [9].Miniature donkeys appear to be more difficult to anesthetize than standard donkeys and mules. Standard dosesof xylazine, butorphanol and ketamine DO NOT usually produce acceptable surgical anesthesia for longerthan 5 min. However, xylazine (1.1 mg/kg IV) with butorphanol (0.04 mg/kg IV) followed by Telazol(1.1 - 1.5 mg/kg IV) produces sufficient anesthesia to allow short surgical procedures (approx 20 minduration). Propofol also provides good anesthesia in miniature donkeys. Following premedication withxylazine (0.8 mg/kg IV), a bolus of propofol (2.0 mg/kg IV) is administered. For procedures longer than 10
min, additional propofol can be given as intermittent boluses (0.2 mg/kg/min). We are continuing toinvestigate drug combinations and dosages, which produce field anesthesia in donkeys, and mules lasting longenough to facilitate the surgical procedure, without resulting in prolonged recoveries. Differences in drugkinetics as well as behavioral differences between donkeys and horses seem to make it difficult to find theoptimal field anesthetic.
Inhalant Anesthesia, Maintenance and MonitoringInhalant anesthetics produce very satisfactory anesthesia in donkeys and mules. The minimum alveolarconcentrations (MAC-values) for halothane and isoflurane in donkeys are very similar to those reported forhorses and ponies [10]. Studies have not been done in mules, but our clinical experience is that concentrationsneeded are very similar to those used in horses or donkeys. Endotracheal intubation is performed manner as inthe horse. Although not anatomically documented, the size of the trachea may be smaller than for a horse ofsimilar size, so it is wise to have a range of endotracheal tubes available.Donkeys and mules are generally very stoic, so it is easy to inadvertently maintain them at too light a plane ofanesthesia. Eye signs (e.g., nystagmus, palpebral and corneal reflexes) do not seem to be as reliable forjudging depth of anesthesia as in the horse; the eye tends to remain quiet until the animal moves, instead ofseeing nystagmus first. Monitoring blood pressure will fairly reliably indicate depth of anesthesia as it will inthe horse; increases in blood pressure are generally seen as the plane of anesthesia decreases. Blood pressurecan be measured non-invasively, or invasively with a catheter placed into an artery. In some donkeys, it isdifficult to palpate the branch of the transverse facial artery usually found under the zygomatic arch.However, they usually have large palpable auricular arteries which can be catheterized. Respiratory rate andcharacter appears to be different in donkeys; normal respiratory rate is higher than for horses and there is lessthoracic excursion (the character of respiration is similar to that of cattle). Donkeys may hold their breath inresponse to pain, instead of increasing respiratory rate. Other supportive care (e.g., use of IV fluids, treatmentof hypotension) should be as for the horse. Donkeys do not hemoconcentrate until they are extremelydehydrated (more than 15%), so the need for fluid therapy must be evaluated by other means than packed cellreadings. Physical exam and history may be helpful, but donkeys are also fairly stoic about not showing signsof illness. It is wise to assume that the animal is sicker than it may appear. Donkeys are also very susceptibleto hyperlipidemia if they become anorexic for any reason [11].
RecoveryDonkeys rarely get hysterical about anything, so recoveries from anesthesia are almost always quiet andsmooth. It is generally impossible to make a donkey get up before it is ready. A rough recovery would bestrong evidence that the animal was painful, having difficulty breathing, or that there was some otherunderlying problem occurring. Occasionally, young donkeys may need a "boost" on the tail to stand;sometimes they will get up rear-legs first, like a cow. Mules vary more, depending on the influence of thehorse portion; mules bred for racing tend to be "flightier" whereas draft mules are usually quiet. Overall, mostmules are quite sensible if well treated, and can be left to self-recover from anesthesia.Phenylbutazone or flunixin can be used to provide analgesia for donkeys and mules. The half-life forphenylbutazone is much shorter than for the horse, but more of the active metabolite (oxyphenbutazone) isproduced [12]. It appears to be difficult to produce toxicity with phenylbutazone in donkeys even when theyare maintained on high (horse) doses for prolonged periods of time (Tex Taylor, unpublished observations).Flunixin has similar characteristics; in standard donkeys the half-life is about half as long as in the horse [13];the half-life is even shorter in miniature donkeys. Dosing intervals may need to be shorter than in the horse foroptimal analgesia.
References
1. Fielding D. The number and distribution of equines in the world, in Proceedings. First Int Colloquium onWorking Equines 1991; 62-66. 2. Yousef MK, Dill DB and Mays MG. Shifts in body fluids during dehydration in the burro, Equus asinus. JAppl Physiol 1970; 29:345-349.
3. Gupta RB, Yadab MP, Uppal PK, et al. Lower susceptibility of donkeys to equine herpes virus and equineinfectious anaemia virus in comparison to horses. In Proceedings. Third Int Colloquium on Working Equines1998; 112-116. 4. Maloiy GMO. Water economy of the Somali donkey. Am J Physiol 1970; 219:1522-1527. 5. French J. The donkey - a small horse? In Proceedings. The donkey: a unique equid. 2000; 2-8. 6. Brown DG, Cross FH. Hematologic values of burros from birth to maturity: cellular elements of peripheralblood. Am J Vet Res 1969; 30: 1921-1927. 7. Matthews NS, Taylor TS, Hartsfield SM, et al. Pharmacokinetics of ketamine in mules and mammoth assespremedicated with xylazine. Equine vet J 1994; 26:241-243. 8. Trawford A. Anaesthesia in the field (including field castration). In Proceedings. The donkey: a UniqueEquid 2000; 25-30. 9. Matthews NS, Taylor TS, Skrobarcek CL, et al. A comparison of injectable anaesthetic regimens in mules.Equine vet J 1992; 11 (suppl):34-36. - PubMed - 10. Matthews NS, Taylor TS and Hartsfield SM. Anaesthesia of donkeys and mules. Equine vet Educ 1992;9:198-202. 11. Watson T. Metabolic diseases of donkeys, in Proceedings. The Donkey: a Unique Equid 2000; 9-13. 12. Mealey KL, Matthews NS, Peck KE, et al. Comparative pharmacokinetics of phenylbutazone and itsmetabolite oxyphenbutazone in clinically normal horses and donkeys. Am J Vet Res 1997;58:53-55. - PubMed - 13. Coakley M, Peck KE, Taylor TS, et al. Pharmacokinetics of flunixin meglumine in donkeys, mules andhorses. Am J Vet Res 1999; 60:1441-1444. - PubMed -
All rights reserved. This document is available on-line at www.ivis.org. Document No. A0607.0700 .
In: Recent Advances in Anesthetic Management of Large Domestic Animals, SteffeyE.P. (Ed.)Publisher: International Veterinary Information Service (www.ivis.org)
Anesthetic Management of Camelids (4 September 2000)
K.R. MamaDepartment of Clinical Sciences, College of Veterinary Medicine and Biomedical Sciences, Colorado StateUniversity, Fort Collins, Colorado, USA. IntroductionSimilar to other species, general anesthesia in Camelids (e.g., llamas, alpacas, camels) may be induced andmaintained with injectable agents, inhaled agents or a combination of these agents. Previous reports describethe use of many drugs (e.g., xylazine, guaifenesin, ketamine, thiopental, halothane and isoflurane) for sedationand general anesthesia [1-5]. Techniques used to support and monitor these animals in the peri-anestheticperiod have also been described in depth elsewhere [1-5]. Hence, the focus of this manuscript will be to add tothis base of information by reviewing new material pertinent to the anesthetic management of primarily llamasand alpacas. Brief descriptions of anesthetic techniques for the camel are also included; readers are referred toa recent review article for further detail [3].
Sedation, Tranquilization, Chemical RestraintIntramuscular (IM) and intravenous (IV) administration of xylazine or xylazine and ketamine has beenextensively described for use in llamas, alpacas and camels [2,4,6,7] While these drugs alone and incombination do provide effects ranging from sedation to short-term anesthesia, the degree (level of sedation oranesthesia) and duration (may range from 10 to 60 minutes) of response in individual animals is variable.While the variability in the response may be influenced in part by unpredictable drug absorption from the IMadministration site, the IM route is still preferred by many veterinarians due to the difficulties encounteredwhen locating and establishing venous access in Camelids.Barrington et al., describe the use of butorphanol (0.1 mg/kg, IM) in combination with intra-testicularlidocaine (2 %, 2 - 5 ml / testicle) for chemical restraint to facilitate standing castration in over 100 llamas [8].Their impression was that the animals receiving butorphanol appeared less stressed than those receiving onlyintra-testicular lidocaine.Using the dose of butorphanol described by Barrington et al., as a starting point and after preliminarydose-response studies, Mama et al., evaluated the cardiopulmonary and behavioral effects of xylazine (0.03 or0.04 mg/kg, IM), butorphanol (0.3 and 0.4 mg/kg, IM) and ketamine (3 or 4 mg/kg, IM) in 7 male llamas and7 male alpacas [19]. Due to prior investigator experience with this drug regime (indicating the need for higherdrug doses in alpacas), alpacas received the higher dose of each drug. Five out of 7 animals in each groupbecame recumbent in an average of 4.3 (llamas) and 6.7 (alpacas) minutes. Induction quality was good withanimals generally showing some degree of ataxia before assuming a sternal or lateral position. Despitereceiving lower drug doses, llamas appeared more deeply anesthetized and remained recumbent for a longerduration (mean time to standing 63 min) than did alpacas (mean time to standing 22 min). All animalsrecovered without apparent complications.During drug-induced recumbency minor manipulations including ocular centesis and catheter placement wereeasily performed. Direct mean auricular arterial blood pressure was well maintained averaging 131 mm Hg inllamas and 144 mm Hg in alpacas; heart rate ranged from 29 - 37 beats/min in llamas and 37 - 49 beats/min inalpacas. While ventilation was only slightly compromised (average PaCO2 was 46 - 49 mm Hg), the PaO2(average 45 - 55 mm Hg, average barometric pressure 640 mm Hg) decreased to clinically unacceptable levels
in some animals implying the need for inspired oxygen supplementation during recumbency induced withxylazine, butorphanol and ketamine.
Regional Anesthetic TechniquesAs demonstrated in the report by Barrington et al., the regional administration of a local anesthetic canfacilitate surgical intervention and minimize the need for administration of drugs with systemic effects [8].This can greatly change the peri-anesthetic management of the animal and minimize the potential forcomplications (e.g., regurgitation, myopathy) associated with drug-induced recumbency. The use of localanesthetics to facilitate laceration repair and surgical intervention (e.g., inverted L block for paralumbarapproach) have been described in llamas and camels and are largely extrapolated from experiences in otherspecies (predominantly cattle) [4,9,10].In camels, 12 - 15 ml of 2 % lidocaine may be administered in the caudal epidural space with the animal insternal recumbency. This dose is reported to produce analgesia of the perineum, udder or scrotum for 1 - 2hours without influencing motor control [10].Grubb et al., evaluated the use of lidocaine and xylazine for epidural analgesia/anesthesia in llamas [11].Diagnostic and surgical procedures involving the rectum, vagina and perineum may be performed in standinganimals using this technique. Further, this technique may be used to provide regional analgesia. Onset andduration of analgesia was evaluated in 6 mature llamas following sacrococcygeal administration of lidocaine(0.22 mg/kg), xylazine (0.17 mg/kg) and a combination lidocaine and xylazine. Behavioral (e.g., sedation) andphysiologic (e.g., heart rate, respiratory rate) were also evaluated at predetermined intervals following drugadministration.Time to onset of analgesia was similar in the lidocaine (average of 3.2 minutes) and xylazine/lidocaine(average of 3.5 minutes) groups. Onset time in the group receiving xylazine alone was longer averaging 20.7minutes; duration of analgesia in this group was intermediate (187 minutes) between the lidocaine only group(average of 71 minutes) and the xylazine/lidocaine group (average of 326 minutes). Evidence of mild sedationwas seen in only some of the animals receiving xylazine. Ataxia was not observed when the animals werestanding or encouraged to stand from the seated position.
Anesthetic Induction and Maintenance TechniquesWhile the aforementioned regional techniques are suitable for numerous situations, general anesthesia iswarranted in animals scheduled for highly invasive surgical procedures (e.g., celiotomy). The inhalationanesthetics (e.g., halothane, isoflurane) have been used to maintain anesthesia in camels, llamas and alpacas. Recently results of two studies highlight the anesthetic dose requirement, behavioral and cardiopulmonaryeffects of isoflurane in llamas [12,13]. The minimum alveolar concentration (MAC) of isoflurane in eightotherwise unmedicated mature llamas was 1.05 +/- 0.17 % (barometric pressure 760 mm Hg) [12]. Anestheticinduction took an average of 19 minutes from time of first isoflurane breath to orotracheal intubation. Animalswere anesthetized for approximately six hours during the study but regained a sternal posture with the abilityto support their heads an average of 23 minutes after the anesthetic was discontinued. Six of the aforementioned eight animals in whom MAC had been previously determined were anesthetizedwith isoflurane in oxygen at a later date and then administered one of three doses (1.0, 1.5 and 2.0 MAC) ofisoflurane in random order [13]. Cardiopulmonary responses were assessed at each dose and during bothspontaneous and controlled ventilation. As anesthetic dose was increased, a decrease in mean arterial bloodpressure and an increase in heart rate were observed in animals during both spontaneous and controlledventilation. Cardiac output and PaCO2 recorded during spontaneous ventilation were higher than thoserecorded when ventilation was controlled. The PaCO2 was also influenced by anesthetic dose inspontaneously ventilating animals (increasing in value as anesthetic dose increased). The average time from induction to endotracheal intubation was 17 minutes. Recovery to sternal recumbencyand standing averaged 15 and 36 minutes, respectively. During anesthetic induced recumbency, spontaneousbehaviors (e.g., swallowing, limb movement) decreased with increasing anesthetic depth. Jaw tone andpalpebral reflex activity appeared to be most consistently influenced by anesthetic dose; positive responsesdecreased as dose increased. Eyelid aperture also tended to increase in a dose-dependent manner and in five ofsix llamas the globe was centrally positioned at 2 MAC (deep plane of anesthesia). Following anesthetic induction with injectable agents, anesthesia has been successfully maintained in camelsusing halothane [14,15]. White et al., report mean saphenous arterial blood pressure ranging from 76-115 mm
Hg during anesthesia maintained with halothane [15]. Mean carotid arterial pressure reported in another studywas lower in halothane-anesthetized camels when compared to those receiving only thiopental [14].Respiration during halothane anesthesia was characterized by a shallow rapid pattern, but PaCO2 increasedprogressively up to a high value of 57 mm Hg [15]. Both studies describe recovery from anesthesia asuneventful. Sternal recumbency was achieved in an average of 25 to 39 minutes for animals in each of twostudies; time to standing was more variable [14,15].While inhalation anesthetics continue to be used to maintain anesthesia, the need for specialized deliveryequipment generally limits the use of this technique to the hospital environment. The advent of short acting,rapidly cleared IV drugs provides veterinarians with the option of maintaining general anesthesia usingcontinuous infusions (or repeated injections) of injectable agents. Duke et al., evaluated propofol, a drug withthese potentially beneficial characteristics, for anesthetic maintenance in 5 llamas [16]. The cardiopulmonaryeffects of two infusions of propofol (0.2 mg/kg/min and 0.4 mg/kg/min) were assessed followingadministration of 2 mg/kg IV for anesthetic induction. The infusions were maintained for 60 minutes duringwhich time llamas receiving the higher dose appeared adequately anesthetized and generally unresponsive toexternal stimuli. Conversely, llamas receiving the lower dose were noise sensitive and made some weakattempts to raise their head. Animals stood an average of 13 to 22 minutes following termination of the lowand high dose infusion, respectively and showed little to no ataxia. During anesthetic maintenance with both infusions of propofol the heart rate was increased (to approximately90 beats/min) over pre-drug values (of approximately 55 beats/min). Mean carotid arterial pressure wassimilar to pre-drug values and ranged from an average of 103 mm Hg to 147 mm Hg during drug-inducedrecumbency. Although the PaCO2 increased and PaO2 decreased in recumbent animals, the values remainedwithin a clinically acceptable range (mean PaCO2 no greater than 45 mm Hg and mean PaO2 no less than 83mm Hg). Three llamas did however become dyspneic and required placement of a nasopharyngeal tube toensure a patent airway.Propofol (2 mg/kg IV) has also been used in premedicated (with xylazine and diazepam) camels to induce andmaintain short-term anesthesia [17]. Duration of recumbency was longer in animals receiving high doses ofxylazine and diazepam and ranged from an average of 26 minutes to an average of 60 minutes. Heart rateincreased from post sedation values averaging 45 beats per minute to a high value of 88 beats per minute 10minutes after propofol administration. Respiratory rate ranged from 12-18 breaths per minute duringdrug-induced recumbency.
Muscle Relaxation During General AnesthesiaWhen muscle relaxation provided by the anesthetic agents alone is not adequate (e.g., intraocular surgery,reduction/repair of a displaced long bone fracture), drugs that block the neuromuscular junction are used asadjuncts during general anesthesia. Hildebrand et al., evaluated the efficacy of atracurium, administered viaintermittent IV bolus (0.15 mg/kg initial dose, followed by 0.08 mg/kg) or IV infusion (0.15 mg/kg initialdose, followed by 0.4mg/kg/hr) in halothane anesthetized, mechanically ventilated llamas [18]. Both methodswere found to provide adequate relaxation in these animals as monitored by reduction of the evoked hind limbdigital extensor tension (twitch), but authors noted that twitch strength recovery time was variable betweenanimals. Residual neuromuscular blockade was antagonized with edrophonium (0.5 mg/kg IV). Atropine(0.01 mg/kg IV) was given with this reversal agent to avoid its muscarinic side effects.
SummaryWhile none of the recent developments in anesthetic management of Camelids provide flawless technique,they offer additional options and opportunities to provide improved care to llamas, alpacas and camelsneeding sedation or general anesthesia.
References
1. Riebold TW, Kaneps AJ, Schmotzer WB. Anesthesia in the llama. Vet Surg 1989; 18:400-404. - PubMed -2. Gavier D, Kittleson MD, Fowler ME et al. Evaluation of a combination of xylazine, ketamine andhalothane for anesthesia in llamas. Am J Vet Res 1988; 49:2047-2055. - PubMed -3. Alsobayil FA, Mama KR. Anesthetic management of Dromedary Camels. Compend Cont Edu - Food
Anim Med Manage 1999 (suppl); 20:125-139.4. Anesthesia. In: Fowler ME. Medicine and Surgery of South American Camelids. Ames: Iowa StateUniversity Press, 1989; 51-63. Available from amazon.com -5. Heath RB. Llama anesthetic programs. In Vet Clin of North Am Food Anim Pract, 1989;5:71-80. - PubMed -6. White RJ, Bali S, Bark H. Xylazine and ketamine anaesthesia in the dromedary camel under fieldconditions. Vet Rec 1987; 120:110-113. - PubMed -7. Bolbol AE. Clinical use of combined xylazine and ketamine anaesthesia in the dromedary. Assiut Vet MedJ, 1991; 25:186-192.8. Barrington GM, Meyer TF, Parish SM. Standing castration of the llama using butorphanol tartrate andlocal anesthesia. Equine Pract, 1993; 15:35-39.9. Said AH. Some aspects of anaesthesia in the camel. Vet Rec 1964; 76:550-554.10. White RJ. Anaesthetic management of the camel. In Higgins A, ed. The camel in health and disease.Philadelphia: Balliere Tindall, 1986; 136-148.11. Grubb TL, Riebold TW, Huber MJ. Evaluation of lidocaine, xylazine, and a combination of lidocaine andxylazine for epidural analgesia in llamas. J Am Vet Med Assoc 1993; 203:1441-1444. - PubMed -12. Mama KR, Wagner AE, Parker DA et al. Determination of minimum alveolar concentration of isofluranein llamas. Vet Surg 1999; 28:121-125. - PubMed -13. Mama KR, Wagner AE, Steffey EP. Circulatory, respiratory and behavioral responses in isofluraneanesthetized llamas. Vet Anaest Analg 2000; 7:1-7 (in press)14. Singh R, Peshin PK, Patil B et al. Evaluation of halothane as an anesthetic in Camels (Camelusdromedaries). ZBL Feur Vet 1994;41:359-368. - PubMed -15. White RJ, Bark H, Bali S. Halothane anaesthesia in the dromedary camel. Vet Rec1986;119:615-617. - PubMed -16. Duke T, Egger CM, Ferguson JG, et al. Cardiopulmonary effects of propofol infusion in llamas. Am J VetRes 1997; 58:153-156. - PubMed -17. Fahmy LS, Faraq KA, Mostafa MB et al. Propofol anaesthesia with xylazine and diazepam premedicationin camels, J Camel Pract Res 1995; 11-113.18. Hildebrand SV, Hill T. Neuromuscular blockade by use of atracurium in anesthetized llamas. Am J VetRes 1993; 54:429-433. -PubMed -19. Mama KR, Aubin ML, Johnson LW. Experiences with xylazine, butorphanol, and ketamine forshort-term anesthesia in llamas and alpacas. In: Proceedings of the World Congress of Veterinary Anaesthesia2000; (in press).
All rights reserved. This document is available on-line at www.ivis.org. Document No. A0608.0900 .
In: Recent Advances in Anesthetic Management of Large Domestic Animals, SteffeyE.P. (Ed.)Publisher: International Veterinary Information Service (www.ivis.org)
Focused Supportive Care: Blood Pressure and Blood Flow during EquineAnesthesia (9 September 2000)
A.E. WagnerDepartment of Clinical Sciences, Veterinary Teaching Hospital, College of Veterinary Medicine andBiomedical Sciences, Colorado State University, Fort Collins, Colorado, USA. Cardiovascular Function During AnesthesiaThe goals for management of anesthetized animals undergoing surgery include providing sufficient centralnervous system depression and muscle relaxation to facilitate surgical conditions, while maintaining adequateperfusion of vital organs with oxygenated blood. Cardiac output (CO), the quantity of blood pumped by oneside of the heart per minute, is the amount of blood available for perfusion of organs and tissues. Awakehorses have an average CO of approximately 70 ml/kg/min, which decreases by aproximately 1/3 to 1/2during inhalation anesthesia, depending on the agent used, the depth of anesthesia, and the mode ofventilation [1,2]. Cardiac output is related to blood pressure according to the formula:P = F x Rwhere P is mean arterial blood pressure (MAP), F is flow or CO, and R is systemic vascular resistance (SVR).Because CO measurement is generally too complicated for routine clinical applications, anesthetists generallyrely on measurement of arterial blood pressure to assess adequacy of circulatory function.Mean arterial blood pressure of awake horses is generally in the range of 105 to 135 mm of Hg, but decreasesduring inhalation anesthesia [1,2]. In most species, a MAP of 60 to 70 mm of Hg is considered to be theminimum pressure that will result in adequate perfusion of vital organs and tissues such as the brain andkidney [3]. An additional consideration for horses (and other large animals such as cattle) is thatanesthetic-induced hypotension and hypoperfusion may lead to inadequate perfusion of their large musclemass, which may be evidenced in the immediate recovery period as post-anesthetic myopathy. Experimentally,post-anesthetic myopathy has been produced by maintaining horses for 3 1/2 hours at a level of halothaneanesthesia deep enough to result in MAP between 55 and 65 mm of Hg and CO between 23 and 29mL/kg/min [4]. Clinically, it has been noted that the greater the degree of hypotension and the longer theduration of anesthesia, the greater is the incidence of post-anesthetic lameness [5]. In severe cases, muscledamage can prevent the horse from being able to stand after anesthesia, and may even necessitate euthanasia[4].Although anesthetists tend to focus on results of measurement of blood pressure, it is important to recall thatchange in blood pressure is not always accurately reflecting change in blood flow (CO) or regional tissueperfusion. In fact, during some conditions, MAP maybe negatively correlated with CO [6]. This is becausechanges in vascular tone also have important effects on blood pressure. For example, horses with endotoxemiasometimes have very low MAP, presumably from vasodilation, yet may maintain reasonably good CO [7]. Incontrast, during surgical stimulation [8,9] or administration of an alpha-adrenergic agonist [10]vasoconstriction frequently causes MAP to increase, while CO may decrease.Both clinical impression and experimental work indicate that horses are more susceptible toanesthetic-induced cardiovascular depression than are dogs. Mean arterial blood pressure of horses at 1.5MAC halothane (which approximates a surgical plane of anesthesia) is decreased approximately 38%compared to the awake state, while dogs at the same anesthetic depth have only a 19% decrease in MAP.
Cardiac index (CI) of horses at 1.5 MAC halothane may be decreased 64% below the awake value, whereas indogs CI is decreased 30% [11]. Therefore it is not surprising that the majority of horses (even young,apparently healthy horses) anesthetized with inhalation agents require therapeutic intervention to maintain aMAP considered to be necessary for adequate tissue perfusion. In a retrospective clinical report published in1988, 55.4% of halothane-anesthetized horses required treatment for hypotension [12]. A cursory survey ofequine anesthesia records from the Veterinary Teaching Hospital at Colorado State University for the monthof August 1999 revealed that 91% of horses subjected to halothane, isoflurane, or sevoflurane anesthesia forelective surgical procedures were treated for hypotension. It is apparent that appropriate management ofanesthetized horses requires the ability to support the cardiovascular system, by use of fluid therapy, inotropes,and/or vasopressors.
Fluid TherapyIn general, a fluid administration rate of 10 ml/kg/hr is adequate for routine, elective inhalation anesthesiaprocedures. Preexisting dehydration or hypovolemia, and/or anesthetic-induced vasodilation, may contributeto intraoperative hypotension and conditions requiring additional fluid administration. For routine fluidtherapy during anesthesia, or for replacement of large-volume deficits, use of lactated Ringer's or one of thecommercially-produced balanced electrolyte solutions is recommended, in order to maintain relatively normalserum levels of sodium, potassium, calcium, and chloride. Dextrose (5%) in water may occasionally beindicated for treatment of primary water deficits or hypoglycemia, and normal (0.9%) saline may be preferredfor animals with hyperkalemia, hyponatremia, or hypochloremia.Horses administered lactated Ringers solution, 20 ml/kg, IV, before halothane anesthesia, maintainedsignificantly higher MAP and a nonsignificant trend toward higher CO and central venous pressure, comparedto control horses that received no IV fluid therapy [13]. However, because of the depression of myocardialcontractility induced by inhalation anesthetics, IV fluids alone may not be sufficient to maintain acceptableCO and blood pressure.Preanesthetic administration of hypertonic (7.5%) saline, 4 ml/kg, IV, resulted in significantly higher MAPand CO compared to control during halothane anesthesia in horses [13]. Hypertonic saline was associated withimproved myocardial contractility and stroke volume [13]. However, clinical use of hypertonic saline isgenerally reserved for emergency treatment of shock, and requires follow-up treatment with balanced isotonicelectrolyte solutions or blood products, in order to avoid depleting intracellular fluids by its osmotic effect.For horses that are hypoproteinemic (serum protein < 3 to 3.5 g/dl, or serum albumin < 1 to 1.5 g/dl), plasmaoncotic pressure may be insufficient to retain fluid within the vascular space, and pulmonary edema may resultfrom administration of electrolyte solutions. To increase or maintain plasma oncotic pressure and improvevascular volume, colloids such as dextrans or hetastarch, or blood plasma can be used. For horses that areseverely anemic (PCV < 20%), whole blood or packed erythrocytes may be required to restore adequateoxygen-carrying capacity.
InotropesPositive inotropes are drugs that strengthen the force of myocardial contractions. Vasopressors are drugs thatstimulate contraction of the muscular tissue of capillaries, arteries, and/or veins, causing vasoconstriction. Thedrugs listed in this section are positive inotropes, but some also have vasopressor effects. For a given drug, therelative inotropic and vasopressor effects often vary with dose.Calcium - calcium gluconate administered to awake horses at 0.1, 0.2, and 0.4 mg/kg/min resulted inincreased CO, stroke index, and contractility, while MAP was unchanged and HR decreased [14]. Bothhalothane and isoflurane cause significant decreases in serum ionized and total calcium concentrations inhorses [15]. In halothane-anesthetized horses, infusion of calcium gluconate (0.1, 0.2, and 0.4 mg/kg/min)resulted in increased MAP, but HR decreased, and contractility and cardiac index did not improve [15]. Inisoflurane-anesthetized horses, calcium gluconate increased contractility and cardiac index as well as MAP,and HR remained decreased only until termination of the infusion [15]. Cardiac arrhythmias associated withcalcium infusion were not detected [15]. However, arrhythmias accompanying calcium administration arepossible occurrences in some clinical circumstances and vigilance is warranted. The authors concluded thatfor isoflurane-anesthetized horses, calcium gluconate at the lowest dosage (0.1 mg/kg/min) was effective ataugmenting cardiac function, but that the highest dosage (0.4 mg/kg/min) would be required forhalothane-anesthetized horses [15]. The effective half-life of calcium solutions is very short; therefore,constant infusion is required to achieve sustained effects.
Dobutamine - dobutamine is a synthetic catecholamine with direct agonist activity at β-1, β-2, and α-1adrenergic receptors [16]. The hemodynamic effects of dobutamine infusion at 3 or 5 µg/kg/min inhalothane-anesthetized horses include increases in systolic, mean, and diastolic blood pressures, CO, and leftventricular dP/dt (an index of contractility), whereas SVR remained unchanged and HR decreased [17].Clinically, an infusion rate of approximately 2 (range, 1.5 to 3.2) µg/kg/min in anesthetized horses is generallyeffective in restoring MAP > 70 mm of Hg [12]. Bradyarrhythmias, such as sinus bradycardia and/or 2nddegree atrioventricular (A-V) block, are potential sequellae of dobutamine therapy, occurring in about 26% ofanesthetized horses [12,17]. A recent study, in which halothane-anesthetized horses were given dobutamine at4 µg/kg/min, suggested that the effective half-life of dobutamine in anesthetized horses may be longer thantraditionally assumed, as peak hemodynamic effects were not achieved within 40 minutes of infusion, andeffects of a 60-minute infusion persisted more than 30 minutes after it was discontinued [18]. The clinicalsignificance of these findings is unclear, as dobutamine is generally very reliable in the treatment of low COand low blood pressure in anesthetized horses, and serious side effects are rare. In addition, dobutamine hasbeen shown to be superior to dopamine, dopexamine, phenylephrine, and saline solution in improving MAP,CO, and intramuscular blood flow in anesthetized ponies [10]. Dopamine - dopamine is a naturally-occurring catecholamine with direct agonist activity at β-1, α-1, and α-2adrenergic receptors, as well as at dopaminergic receptors [17]. Dopamine is also reported to have indirectadrenergic activity through release of endogenous norepinephrine [17]. In one study of anesthetized horses,dopamine infusion at 2.5 or 5 µg/kg/min significantly increased CO, but because of decreased SVR, MAP didnot change [19]. In another study, although dopamine infusion at 3 µg/kg/min did not alter any hemodynamicvalues, an infusion at 5 µg/kg/min increased CO and left ventricular dP/dt. Again, because of decreased SVR,there was no significant change in MAP at that dosage [17]. At an infusion rate of 10 µg/kg/min, both CO andMAP significantly increased, with SVR returning (increasing) to baseline values [17]. Increased HR and 2nddegree A-V block occurred in some horses given the 2 higher infusion rates of dopamine [17]. A more recentstudy reported the occurrence of premature atrial and ventricular contractions, ventricular tachycardia, andventricular fibrillation in halothane-anesthetized horses administered a one-hour infusion of dopamine at 10µg/kg/min [20]. Therefore it appears that, in anesthetized horses, dopamine is a less potent inotrope thandobutamine, is less efficacious at increasing MAP, and is more likely to induce serious cardiac dysrhythmias[17].Dopexamine - dopexamine, a structural analogue of dopamine, is reported to be a potent β-2 adrenergicagonist, a weak dopaminergic (DA-1) agonist, an inhibitor of reuptake of norepinephrine at sympathetic nerveterminals, and has no α- and minimal β-1 adrenergic activity [21]. At stepwise infusion rates of 5, 10, and 15µg/kg/min in halothane-anesthetized horses, HR, cardiac output, and MAP increased, while SVR decreased,in a dose-dependent manner [21]. However, the side effects of dopexamine, which include tachycardia,tachyarrhythmias, profuse sweating, muscle twitching, and a noticeable lightening of anesthesia depth, limitits usefulness in clinical cases [10].Ephedrine - ephedrine is a non-catecholamine sympathomimetic with both direct and indirect actions at α andβ adrenergic receptors. In both lightly and deeply halothane-anesthetized horses, ephedrine (0.06 mg/kg) hasbeen shown to increase CO, stroke volume, and arterial blood pressure. These effects were more pronounced,and SVR was decreased, at the deeper plane of anesthesia [22]. Ephedrine has the advantages of beinginexpensive and simple to administer; it can be given as an IV bolus, rather than as a continuous infusion as isrequired with many inotropic drugs. The same dose of ephedrine can be repeated several times, if hypotensionpersists or recurs a few minutes after the initial dose. Heart rate may increase or decrease slightly, but changesare usually transient and dysrhythmias are rare. Although experimentally ephedrine has been shown toincrease cardiac output in anesthetized horses, clinical impression suggests that ephedrine is not asconsistently reliable as dobutamine at increasing blood pressure. Ephedrine may increase the requirement foradditional anesthetic agent [23].Epinephrine - epinephrine is a potent inotrope, but its clinical usefulness is limited by its arrhythmogenicity.In halothane-anesthetized horses, infusion of epinephrine produces premature ventriuclar depolarizations and,in some cases, ventricular fibrillation and death [24]. Hypercapnia, which is common inspontaneously-breathing anesthetized horses, may exacerbate the risk of epinephrine-induced ventriculararrhythmias [25]. Epinephrine is not recommended for routine use in anesthetized horses, but remains acomponent of cardiopulmonary resuscitation in response to cardiac arrest.
VasopressorsThe following drugs are used primarily for their vasopressor effects, although they may also affect CO.Norepinephrine - norepinephrine is a naturally occurring endogenous neurotransmitter approximately equal(or slightly less) in potency to epinephrine for stimulation of β-1 (cardiac) receptors. It is a potent α-agonistand produces intense arterial and venous vasoconstriction. A continuous infusion of norepinephrine (0.1-0.2µg/kg/min, IV) is used in horses to provide short term (e.g., 15 - 30 minutes) treatment of refractivehypotension (Steffey, EP, personal communication). Like dobutamine, but unlike phenylephrine the actions ofnorepinephrine are short-lived; usually subsiding within 2 - 5 minutes of discontinuing IV infusion. Adecrease in HR (due to a pressor induced reflex in vagal tone) commonly accompanies use of norepinephrine.Clinical experience suggests cardiac arrhythmias in horses are less frequent with use of norepinephrinecompared to epinephrine. Despite infrequent occurrence of life-threatening ventricular arrhythmias with use ofnorepinephrine, vigilance is necessary.Phenylephrine - phenylephrine is an α-1 adrenergic agonist which in conscious horses causes vasoconstriction(increased SVR), resulting in an increase in MAP, while cardiac output is decreased [26]. There is minimalpublished documentation of its effects in anesthetized horses. In a study of 8 halothane-anesthetized ponies,phenylephrine infusion (0.25 to 2 µg/kg/min) not only failed to improve intramuscular blood flow, but wasassociated with clinical signs of post-anesthetic myopathy in 2 of the ponies.10 The authors suggested thatphenylephrine should be reserved only for situations in which hypotension is refractory to other medications,and not used for routine treatment of halothane-induced hypotension in horses [10]. The use of phenylephrinemight be considered an appropriate adjunct to treatment of hypotension associated with large doses ofacepromazine, with endotoxemia, or other situations in which vasodilation is profound. Clinical experiencesuggests that a phenylephrine bolus of 0.002 mg/kg, IV, may be effective at increasing MAP. This dosage maybe repeated if needed, or a constant rate infusion can be administered for prolonged effect. Heart rate shouldalso be monitored carefully, since phenylephrine can contribute to bradycardia [26].
AnticholinergicsBecause blood pressure is directly related to CO, and CO is directly related to HR, prevention of bradycardiaby administration of anticholinergics appears to be a logical approach to alleviating intraoperativehypotension. However, if bradycardia in horses is arbitrarily defined as HR < 25 beats/min, only a smallminority of anesthetized horses are actually bradycardic. Anesthetic-induced hypotension is more rationallyand effectively treated by use of drugs that improve contractility, such as dobutamine or ephedrine. Inaddition, administration of anticholinergics to horses has been shown to depress gastrointestinal motility andincrease the risk of abdominal discomfort or colic [27,28]. For these reasons, it is recommended that the use ofanticholinergics be limited to horses that are truly bradycardic as well as hypotensive, that only low dosages beused, and only in horses without predisposition to gastrointestinal problems.Atropine - as mentioned previously, atropine is not routinely administered to horses because of concerns aboutpossible detrimental effects on gastrointestinal motility. Clinical signs of abdominal discomfort have beenobserved following atropine dosages of 0.044 and 0.176 mg/kg, IV, in ponies [27]. However, a smaller doseof atropine (0.006 mg/kg, IV) given at anesthesia induction, 1 hour after detomidine administration, reverseddetomidine-induced bradycardia and was associated with higher MAP and reduced need for inotropic supportduring halothane anesthesia [29]. At that dose of atropine, none of the horses developed cardiac dysrhythmiasor signs of colic [29]. Clinical experience suggests that even smaller doses of atropine (0.002 to 0.004 mg/kg,IV) are often effective at correcting intraoperative bradycardia or bradyarrhythmias associated with α-2agonists and/or dobutamine administration. However, atropine has been shown to reduce the arrhythmogenicdose of dobutamine in halothane-anesthetized horses [30]. Therefore, if atropine is used to treat intraoperativebradycardia, it is recommended that dobutamine or other inotrope infusions be terminated a few minutesbefore atropine is given.Glycopyrrolate - in awake horses, glycopyrrolate, 0.005 mg/kg, IV, increased HR but caused some depressionin gastrointestinal motility; a dosage of 0.01 mg/kg produced signs of colic [28]. When glycopyrrolate, 0.0025mg/kg, IV, was used as a premedication for xylazine and ketamine anesthesia, HR, CO, and blood pressureincreased for approximately 30 minutes, but gastrointestinal motility was reduced for up to 9 hr, and one horseshowed mild signs of colic [31]. In halothane-anesthetized horses, glycopyrrolate at 0.0025 to 0.005 mg/kg,IV, resulted in increased HR and improved blood pressure [32]. One horse out of 17 in the latter study did
develop clinical signs of colic, although it was not clear that glycopyrrolate was a contributing cause [32].Therefore glycopyrrolate, like atropine, should be used with caution in horses.
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All rights reserved. This document is available on-line at www.ivis.org. Document No. A0612.0900.
In: Recent Advances in Anesthetic Management of Large Domestic Animals, E.P. Steffey (Ed.) Publisher: International Veterinary Information Service (www.ivis.org), Ithaca, New York, USA. Hazards Associated with Laser Surgery in the Airway of the Horse: Implications for the Anesthetic Management (18-Apr-2003) B. Driessen, L. Zarucco, L. E. Nann and L. Klein
Department of Clinical Studies - New Bolton Center, School of Veterinary Medicine, University of Pennsylvania, PA, USA.
Summary Over the past two decades, the use of lasers has become an integral part of upper airway surgery in the horse as it allows for more precise tissue dissection, produces less bleeding and tissue trauma, and reduces the incidence of postoperative complications. Nevertheless, laser surgery in the anesthetized horse also holds novel hazards for patient and operating room personnel alike. Certainly laser surgery constitutes a challenge as operation of a high-energy temperature source in the airway, often in vicinity of the endotracheal tube, bears the risk of accidental tube ignition and subsequent airway fire. This danger is of particular concern during inhalation anesthesia, when patients breathe oxygen-rich gas mixtures that readily support tube combustion. Therefore laser surgery in the respiratory tract requires a detailed pre- and intraoperative communication and cooperation between surgeon and anesthesiologist, and a specific anesthetic management tailored to the individual surgical procedure and laser instrument being used. By following prescribed safety procedures and precautions, and employing appropriate measures in an emergency situation, both surgeon and anesthesiologist can markedly reduce the risk of potentially life threatening complications.
Introduction The term laser is an acronym for light amplification by stimulated emission of radiation. Light emitted by a laser differs from natural light in that it is coherent (waves are all in phase with each other in space and time), collimated (highly directional), monochromatic light (light of one wavelength), and of high energy [1-3]. Since the introduction of lasers into medicine in the early 1960s, laser technology has progressed rapidly and is now widely used in all surgical disciplines. A variety of surgical lasers are available today with different physical properties (Table 1). The destructive effect of a given laser output is dependent upon the wavelength emitted, the power density projected, the time the tissue is exposed to the high-energy source, and the blood supply to the tissue. Laser light might be accurately focused on small areas evaporating tissue and cauterizing small blood vessels. Long wavelength lasers, such as the carbon dioxide (CO2) laser, transfer energy to tissue and water, therefore tissue penetration is shallow (approximately 0.01 mm). The neodymium:yttrium aluminum garnate (Nd:YAG) laser, on the other hand, operating with a shorter wavelength than the CO2 laser readily penetrates tissue up to a depth of 5 mm.
Table 1: Properties of surgical lasers [12,28,35].
Laser type Wavelength Light spectrum (nm) Fiberoptic transmission
Use of lasers in the airway of the horse requires technology that allows transmission of the laser beam through fiberoptic delivery systems, small and flexible enough to fit the biopsy channel of an endoscope in order to reach the surgical field in the respiratory tract [4]. The Nd:YAG laser became the first commercially available device that fulfilled this condition. More recently the gallium aluminum arsenide (GaAlAs) diode laser has entered the veterinary market as well and is now widely available for transendoscopic laser surgery in the airway [5]. As a result of this technological progress, the laser is today an established instrument for upper airway surgery in the horse and offers the advantages of less traumatic tissue dissection with reduced risk for bleeding and postoperative complications (e.g., laryngeal edema) as well as faster return to preoperative training activity. The close proximity of laser beam and endotracheal tube (ETT) in the surgical field, however, is of great concern to the anesthesiologist as it inflicts significant risk upon the equine patient during general anesthesia. This chapter will describe common problems associated with the use of lasers in airway surgery, lists precautions to be considered to minimize the risk of laser-related complications as well as rapid corrective measures to be taken in the event of complications.
Common Laser Applications in Equine Airway Surgery Over the past years, the use of lasers in equine upper airway surgery has become increasingly popular [6,7]. Among frequently performed procedures that involve transendoscopic laser surgery are correction of epiglottal entrapment, vocal cordectomy and laryngeal sacculectomy, partial soft palate resection (staphylectomy), excision of arytenoid cartilage granulomas (resulting from arytenoid chondritis), ablation of pharyngeal lymphoid hyperplasias and pharyngeal masses, and removal of sub- or dorsal epiglottic cysts or granulomas. Lasers have also been effectively applied in tissue biopsies, creation of a salpingopharyngeal fistula, treatment of conditions such as progressive ethmoid hematoma, guttural pouch tympanitis, choanal atresia, axial deviation of the aryepiglottic fold, tracheal ulcers and pyogranulomas, and debridement of dorsal epiglottic abscesses. A detailed description of the surgical techniques and instruments used for those procedures is beyond the scope of this chapter and the interested reader is referred to recent review papers and equine surgery textbooks [6-11].
Hazards Associated with Laser Surgery in the Airway The American National Standards Institute (ANSI) has classified most medical lasers, including the CO2, Nd:YAG, and GaAlAs diode lasers, as Class IV, or most hazardous lasers on the basis of their optical emissions [12]. Direct intrabeam viewing or contact with the laser beam are considered the most dangerous, but also specular or diffuse reflection of laser light may damage skin or eyes of surgeons, other operating room personnel and the patient alike, unless appropriate safety measures are taken (see Table 2). While light emitted from lasers in the far infrared portion of the spectrum, such as the CO2 laser, is highly absorbed by all surfaces and may damage only the outer layers of the eye, causing corneal ulceration and opacification, infrared light from the Nd:YAG or GaAlAs diode lasers is transmitted through the cornea and lens and, thus, may damage the retina [12]. These risks are minimal during transendoscopic laser surgery in the airway, where the threat of direct exposure is reduced.
In general, airway surgery has a disproportionately larger potential for complications than surgery in other parts of the body because surgical manipulation in the airway may lead to serious impairment of respiratory function and complications in the recovery period. This applies regardless of whether or not a laser instrument is used or the patient is awake or anesthetized. Hence, general complications associated with surgery in the respiratory tract, such as airway obstruction, aspiration (blood, tissue particles, etc.), hypoventilation, hemorrhage, cardiac arrhythmias (e.g., vagal or sympathetic nerve stimulation), or postoperative laryngeal edema, may also occur during or after laser surgery in the airway.
The main problem associated with use of lasers in airway surgery is the introduction and operation of a high-energy temperature source in the patient’s airway [13]. Though reported to be rare in the horse, inappropriate handling of the laser fiber tip and/or endoscope may cause inadvertently deep tissue trauma, particularly when exposure time is long and power density is high or the instrument is operated in the continuous (opposed to pulsed) mode [6,7]. Dependent on location, those lesions may cause extensive damage to mucosal and submucosal tissue, blood vessels, nerves, and laryngeal or tracheal cartilage. In addition, in human patients transesophageal or bronchopleural fistulas and pneumothorax are reported complications of laser-induced tissue trauma [14,15].
In intubated anesthetized patients breathing O2-enriched gas mixtures the use of lasers in the airway bears a significantly higher risk of an airway fire. Any inadvertent misdirecting of the laser beam may cause heat damage to the ETT, considering that the tube is typically in close proximity to the laser target in the surgical field. In the best of circumstances, the laser may
hit the tube for only a very brief moment causing no more damage than a small, partial, or perhaps complete, puncture of the tube. Significant leakage of inhalant gases into the surgical field can occur if the perforation of the tube wall is complete. Any puncture of the cuff, the most vulnerable part of the ETT, will immediately cause it to collapse, resulting in a major leak that mandates immediate discontinuation of positive pressure ventilation (PPV) and significantly increases the risk of aspiration. Dependent on contact time, the intense heat generated by lasers can ignite all ETT materials commonly used in anesthesia [16]. This risk is particularly high if the laser beam hits any dark colored labels or marks on the tube’s surface, as laser light absorption is increased in those spots. Once ignited, the anesthetic gases that pass through the ETT may readily support combustion, leading to a "blowtorch effect" (Fig. 1), best described as flames streaming from the distal (tracheal) end of the burning ETT and causing severe burn injuries that may reach far into the bronchial tree [17]. Due to a Venturi effect, flames of the laser combustion may sweep a jet-like stream of hot gases and soot particles into the lower airways worsening tissue damage in trachea and lungs (Fig. 2). Extensive thermal injuries associated with fire and gas explosion in the airway may affect pharynx, larynx, trachea and lower airways. Most dangerous are thermal lesions of epiglottis and tracheobronchial tree, which may cause severe posttraumatic edema (leading to partial or complete airway obstruction) and impairment of pulmonary gas exchange [18].
Figure 1. A blowtorch effect may result from a laser impingement and subsequent ignition of the proximal part of the endotracheal tube. Once ignited, the oxygen-rich anesthetic gases passing through
the endotracheal tube may support combustion, leading to a blowtorch fire and immediate heat damage to the cuff that then rapidly collapses. - To view this image in full size go to the IVIS website at www.ivis.org . -
Figure 2. This diagram depicts the Venturi effect that may occur during positive pressure ventilation when the cuff of the endotracheal tube suddenly collapses due to a blowtorch fire in the airway. In the moment of cuff deflation, gas under high pressure (jet stream) exiting the distal end of the endotracheal
tube entrains ambient air because the pressure around and behind the distal end of the tube becomes lower (relative negative pressure area N) with respect to the pressure in the mainstream gas flow (positive pressure area P). This further enhances the propulsive force of the blowtorch flame. - To view this image in full size go to the IVIS website at www.ivis.org . -
Besides the laser source itself, "laser smog", a fume formed during tissue coagulation by the laser, is another source of danger [17,19]. It contains organic materials (e.g., xylene or toluene) that are noxious and mutagenic. Particularly in the non-intubated horse smoke particles are easily inhaled by the patient and may cause an inflammatory response in the lower respiratory tract with or without clinical symptoms such as coughing and forceful breathing [20]. While of less concern to the patient when intubated and connected to an anesthetic circuit, laser smog can be hazardous to surgeons and other operating room personnel as well. They may develop bronchial inflammation with bronchospasm, alveolar edema and potentially diffuse pulmonary atelectasis when inhaling the smoke [19]. Mixing of laser smog with O2 may occur where there is an insufficient seal of the ETT cuff, or a cuff deflation, and may rapidly produce a highly explosive gas mixture that further supports or enhances combustion [19].
The ease with which tube combustion occurs may depend on the tube material, inhaled gas composition, duration of laser exposure and the power density [18]. Silicon tubes burn more easily than red rubber and polyvinyl chloride (PVC) ETTs [21]. Studies comparing the combustibility of red rubber versus PVC have produced conflicting results. While Patel and Hicks [22] reported a higher heat resistance of red rubber tubes as compared to PVC tubes upon direct exposure to a laser, Wolfe and Simpson [16] found opposite results under similar experimental conditions. The risk of ETT combustion is significantly increased with inspired O2 concentrations exceeding 30% (FiO2 > 0.3) [23-25]. As reported by Wolfe and Simpson [16], red rubber tubes combust at a significantly lower O2 concentration (18%) than PVC tubes (26%). Silicone, much like red rubber, can ignite in room air [23]. In addition, the incidence of "flaring" of carbonized tissue increases as the O2 concentration at the surgery site increases [19].
Besides direct heat injury to the airways, patients may demonstrate severe respiratory symptoms resulting from the toxic effects of waste products of tube combustion. Burning of PVC produces hydrochloric acid and vinyl chloride, both of which can cause severe airway irritation and bronchoconstriction [26]. Red rubber tubes are stiffened with compounds that reduce their flammability, but when ignited produce thick black smoke which then may be inhaled by the patient; fortunately, the smoke does not seem to contain irritating substances [27]. Silicone, if ignited, rapidly becomes a brittle ash that crumbles easily and, hence, tends to quickly accumulate in lower airways and lung [22].
Safety Precautions for use of Lasers General Precautions - Although the use of lasers for minimally invasive airway surgery of the horse has proven to be relatively safe, important safety procedures must be followed in order to protect both patient and operative personnel from the hazards of lasers [6]. Laser safety training is available for veterinarians and veterinary technicians with certification courses administered by the American College of Veterinary Surgeons (4401 East West Highway, Suite 205, Bethesda, MD 20814-4523, USA) and the American Society for Laser Medicine and Surgery (2404 Stewart Square, Wausau, WI 54401, USA). It is recommended that all operating room personnel become familiar with a Laser Safety Protocol, which lists the main safety precautions for use of lasers in surgery (Table 2). Also all new equipment and procedures should be carefully reviewed before being allowed to operate in the hospital on patients.
The most efficient way to protect the eye from the laser is to avoid any contact with its emitted light and to use instruments with matte surfaces that diffuse reflected beams. Whenever a surgical laser is used, doors to the operating room should be closed and access restricted. A conspicuous warning sign (e.g., "Danger - Laser radiation: Do not enter without appropriate eye protection") should be displayed on the outside of all doors of the surgery suite, indicating the type of laser used and the form of eye protection required [1,6,12]. This will prevent any accidental exposure of animal health care personnel to the laser beam when entering the room. Surgeon, anesthesiologist and all other personnel in the operating room must wear goggles of specific colors in order for the laser light to be absorbed. There is only one exception to this rule. Eyes can be protected from CO2 laser radiation by any glasses or goggles because far infrared light emitted by this laser is absorbed by almost all surfaces [12]. It is important to note that tinted goggles may impair the anesthesiologist’s ability to evaluate changes in the horse’s mucous membrane color and to read screens of certain ECG or other physiological data monitors, particularly those using color-coding for display of various traces or data. The eyelids of the horse may be closed and covered with moist gauze patches in order to avoid any remotely possible exposure to the laser beam.
Safe transendoscopic laser surgery in horses requires a laser-compatible endoscope [6,7]. This includes an eyepiece that is equipped with an interchangeable filter, which filters out the wavelength emitted by the specific laser being used. Otherwise the surgeon is at risk of eye injury when viewing the operative field during laser operation without the obligatory protective eyewear that is specific to the wavelength of the laser being used. Use of a video recording/monitor system attached to the endoscope does not only offer the surgeon the advantage of a magnified display of the surgical field, but also allows the anesthesiologist to carefully monitor the progress of the surgical procedure and the position of the laser tip in relation to the airway and ETT at any given time. In this case, the filter mentioned before will prevent an optical flare, which may occur upon activation of the laser, and may obscure visibility for the surgeon or cause complete "whiteout" of the video screen. Activating the laser only when the tip of the laser fiber is located within the body cavity and the surgical image is viewed on a television monitor further minimizes the risk of ocular injury to surgeon, patient and operating room personnel.
The tip of laser-compatible endoscopes usually contains a ceramic element to protect the tip of the endoscope from heat generated by the laser beam [6]. Keeping the tip of the laser fiber at least 1 cm beyond the tip of the endoscope when performing surgery will reduce the amount of heat applied to the surface of the endoscope and minimize the accumulation of debris on the surface of the lens and thus decreases the risk of ignition of this material and/or the distal end of the endoscope.
Of less concern than eye injury is skin damage associated with lasers. This may range from a minor erythema to a full-thickness burn, and may affect primarily the patient’s skin close to the surgical site and, much less commonly, the hands of the surgeon or his assistant [12]. More severe damage can occur when drapes or other flammable material in close vicinity to the operating field ignite. In order to prevent burns and accidental exposure during laser use, the patient’s skin around the operative field (e.g., a tracheostomy site) should be covered with wet towels.
As mentioned before, evaporation of tissue by laser energy produces smoke known as "laser smog" that may cause bronchial inflammation and bronchospasm, nausea, vomiting and lacrimation in susceptible individuals [1,12,19]. Adequate suction applied close to the site of smoke production or continuous gas evacuation from the suction channel of the endoscope will reduce environmental pollution and thus protect both surgeon and operating room personnel from the effects of smoke inhalation, and at the same time facilitate the operator’s view of the surgical field. Wearing of special laser masks that prevent inhalation of smoke particles can further help reduce the risk of adverse effects of laser plume in susceptible individuals.
Specific Anesthetic Considerations during Laser Surgery in the Airway of Horses - Because of the described hazards, especially that of tracheal tube fire, any laser procedure in the airway of the horse under general anesthesia requires thorough pre- and intraoperative communication between anesthesiologist and surgeon and mutual preparedness to solve unexpected
complications as a team.
Before induction of anesthesia, a plan should be formulated for the various steps in both administering anesthesia via the airway and performing surgery as well. A detailed knowledge of the location and type of surgical procedure to be performed is essential for the anesthesiologist, who has to develop a strategy that fulfills the goal of satisfactory surgical access to the airway while maintaining a safe ventilatory pathway. Some laser procedures in the upper airway of horses (e.g., arytenoidectomy, excision of large subepiglottic granulomas) are best performed through a ventral laryngotomy incision and hence require a tracheostomy for placement of an ETT, thereby providing an optimally secure airway with minimal risk of an airway fire during laser operation. Similarly, nasal intubation for procedures in more rostral areas of the larynx often achieves both a safe airway and relatively unhindered access to the surgical site with little chance for the laser beam getting into contact with the ETT and eliciting an airway fire. Whenever circumstances allow for a significant spatial separation of surgical field and airway (or ETT), a routine inhalation anesthetic protocol might be used safely. In all other situations precautions need to be taken to minimize the fire hazard associated with laser surgery in the airway.
Since all ETTs used in large animal anesthesia are made of inflammable material (silicone rubber, red rubber or PVC), protection of these tubes from laser damage can be achieved by carefully wrapping them with self-adhesive, non-reflective aluminum tape in a spiral fashion with overlapping edges, beginning just above the ETT’s cuff and ending at the Y-piece adapter [19]. However, laser beams may still penetrate these metallic foils and/or may be reflected off the metallic surface into surrounding tissues. In addition, the tape may not always adhere adequately to the tube and may loosen or break off during intubation or extubation, resulting in aspiration of or airway obstruction with tape particles. As mentioned before the cuff is the most vulnerable part of any ETT and, when ruptured, allows a massive leak of anesthetic gases, leading to hypoventilation of a ventilated patient as well as providing an O2-rich environment for ignition of the tube [19]. Filling the ETT cuff with water or saline (possibly mixed with methylene blue as an indicator of rupture) may reduce fire by dispersing the heat energy [29], but might also result in excess pressure on the tracheal mucosa. When the cuff is punctured by the laser beam the fluid can act as an immediate fire extinguisher [30]. Laser resistant ETTs that use opaque or foam coverings, even metal inserts, are commercially not available in sizes appropriate for use in horses, and have met with only limited success in retarding airway fires [1,12,23].
Both total intravenous (TIVA) and inhalant anesthetic techniques are appropriate for horses undergoing general anesthesia for laser surgery in the upper respiratory tract. This applies particularly when lasers are used for only a limited period of time during the initial or final phase of the surgical procedure. In horses inhalation anesthesia offers generally a greater advantage for intermediate and long-term procedures (> 60 - 90 minutes), though progress has been made recently to improve techniques using TIVA in horses for prolonged general anesthesia [31]. In general, horses undergoing airway laser surgery, except may be for very short procedures, should be intubated to avoid inhalation of "laser smog" and aspiration of tissue debris and blood. In most anesthetized horses breathing room air will result in hypoxemia, thus necessitating O2 supplementation, which however, will negate the possible advantages of TIVA in terms of reducing the fire hazard. Anesthetic vapors currently in common use in equine anesthesia (halothane, isoflurane, and sevoflurane) are not inflammable in clinically used concentrations [1]. Regardless of the technique used, it is recommended to use the lowest concentration of inspired O2 (FiO2) that is compatible with adequate oxygenation of the patient [32], because, as mentioned before, the higher the O2 concentration the greater the risk that combustible material hit by the laser beam will ignite [23-25]. Based on the authors’ experiences concentrations of 25 - 40% (FiO2 < 0.4 with PPV) are usually adequate, except perhaps in some larger animals such as unusually heavy Warmblood or draft horses. Though not commonly used in horses, it is important to stress that also nitrous oxide (N2O) supports combustion, when it disintegrates into nitrogen (N2) and O2 at temperatures above 450ºC (N2O --> N2 + 1/2 O2 + heat) [25]. Thus, N2O should be strictly avoided during laser surgery. Room air, nitrogen (N2), or helium (He) should be used to reduce FiO2, which effectively decreases the risk of igniting the ETT and minimizes the chance of producing a highly explosive gas mixture in the event of an ETT cuff failure (leakage or deflation) that would allow laser smog to mix with the oxygen of the inspired gas.
Regardless of the anesthetic protocol and ventilatory technique (spontaneous or mechanical ventilation) used, administration of an inspired gas mixture low in FiO2 requires careful monitoring of the patient’s oxygenation status. Continuous recording of the inspired O2 concentration combined with pulse oximetry (SPO2) and/or repeated arterial blood analysis (SaO2, PaO2) is mandatory to detect signs of oxygen desaturation accurately, allowing for rapid corrective measures. Though PPV might often not be absolutely necessary, it is the authors’ experience that anesthetized horses breathing spontaneously gas mixtures low in FiO2 (< 0.4) tend to desaturate significantly more frequently than mechanically ventilated horses. However, it is important to recognize that if the ETT is ignited by a laser burst, PPV will further support the development of the above described
"blowtorch" effect and must be immediately discontinued [17,30].
* Adopted and modified from [32].
Table 2: Laser safety protocol for surgery in the equine airway.*
Precautions
I. Prevention of unintentional exposure to laser radiation.
1. Limit access into the operating room (locked doors) when laser is in use. 2. Display warning signs on the outside of all entrances into the surgery room. 3. Avoid direct eye contact with laser beam. 4. Wear laser-specific, protective glasses with side protectors. 5. Cover the patient’s eyes with moist gauze patches if laser exposure is possible. 6. Cover patient’s skin surrounding the operative field if exposure is possible.
II. Smoke evacuation.
1. Aspirate "laser smog" from the surgical field with separate metal suction tip. 2. Evacuate smoke from suction channel of the endoscope. 3. Use laser masks to protect susceptible individuals from smoke inhalation.
III. Transendoscopic laser application.
1. Use laser-compatible endoscope. 2. Attach filter specific for the laser’s wavelength to eyepiece.
IV. Instrument selection.
1. Use instruments with matte surfaces that diffuse reflected laser beams. 2. Test all new laser equipment and instruments prior to use in the patient.
V. Specific anesthetic considerations.
1. Allow satisfactory surgical access to the airway while maintaining safe airway. 2. Shield endotracheal tubes with self-adhesive, non-reflective aluminum tape, if deemed
necessary. 3. Inflate endotracheal tube cuff with saline (+/- indicator dye) to reduce puncture or fire
a. Mix oxygen with helium (alternatively nitrogen or air). b. Avoid nitrous oxide because of its combustibility. c. Use lowest FiO2 compatible with adequate arterial blood oxygenation.
5. Carefully monitor FiO2 and for evidence of hypoxemia (SPO2, PaO2). 6. Employ positive pressure ventilation in order to decrease risk of hypoxemia.
VI. Prevention of postoperative hemorrhage and tissue swelling.
1. Presurgical irrigation of mucosal surface areas with vasoconstrictor containing solutions (e.g., epinephrine, norepinephrine).
Postoperative edema of and bleeding from operated tissues are still a common complication following laser surgery in the upper airway of the horse [6,7]. Pre- or intraoperative administration of non-steroidal anti-inflammatory drugs (phenylbutazone) and eventually glucocorticoids (prednisolone, dexamethasone) helps minimize inflammatory reactions that otherwise may lead to potentially life threatening airway obstruction during the recovery period. If it is anticipated that edema formation will be more severe or may persist despite anti-inflammatory medication, a tracheostomy with subsequent tracheal tube placement distal to the surgical site may be indicated to maintain an open airway. Hemorrhage from the surgical site can be reduced by preoperative irrigation of the mucosal surface area with solutions containing an adrenergic vasoconstrictor (e.g., epinephrine, norepinephrine) [6].
Prior to moving the patient to the recovery stall at the end of the anesthetic, the head should be lowered to allow drainage of blood clots, cell debris and remaining flush solution. Subsequently, the (most commonly nasally placed) ETT should be pulled temporarily up to the level of the surgical site with its cuff inflated, as this further facilitates clearing of remaining blood clots from the airway. The cuff of the ETT should remain inflated throughout the recovery period till the horse is standing to prevent aspiration of blood from persisting hemorrhage. At that time, the ETT can be withdrawn with the cuff still partially inflated. If the ETT is rather small in diameter for the size of the horse, thus significantly increasing the patient’s work of breathing, the cuff might be deflated already earlier, however not before an adequate cough reflex has returned.
Management of Airway Fire and other Complications - Serious complications associated with laser surgery in the airway of the horse are very rare and then often due to unfamiliarity with the specific laser being employed or due to disregard of appropriate safety precautions described above [3,6,7]. Though the incidence of airway fires during laser surgery in the horse is unknown, earlier studies in human medicine, conducted at times when laser-resistant ETTs were not yet available, report an incidence of 0.4 - 1.5% [17,33]. Since it is impossible to totally eliminate the risk of fire when a laser is used in the airway, the entire operating room team must be familiar with all steps to be taken in the event of an airway fire, prepare for them prior to surgery and be constantly on the alert during the procedure [12,28]. If immediate steps are followed to prevent fire from extending down the tracheobronchial tree (Table 3), and if appropriate secondary and postoperative steps are taken in
Table 3: Management of airway fire during laser surgery (modified from [12,30,34,35]).
ThirdIrrigate surgical site with saline if smoldering persists and extinguish remaining flames of removed endotracheal tube with aqueous fluid. Apply suction to clean airway.
Fourth Reintubate trachea and ventilate with as low a FiO2 as possible if patient is apneic, severely hypoventilating, or hypoxemic.
Secondary
Fifth Evaluate extent of burn injury by endoscopy (larynx, trachea, bronchi).
SixthReintubate the trachea (if not already done) or perform a tracheostomy with tracheal tube placement if necessary. Reinstitute positive pressure ventilation if required. Administer steroids and antibiotics as needed.
Seventh Monitor oxygen status of patient with pulse oximetry and arterial blood gas analysis throughout the remainder of the surgery and the postoperative recovery period.
Postoperative
EighthThoracic radiographs if indicated. Symptomatic treatment as needed including O2 supplementation, fluid therapy, anti-inflammatory, pain and antimicrobial medication.
evaluating and treating the injuries that occurred, the morbidity and mortality from a laser fire in the airway can be minimized [12,30,34,35].
In the event of an airway fire, immediate disconnecting of the anesthetic breathing circuit from the tracheal tube is essential to stop any gas flow which otherwise would enhance and maintain the airway fire. Subsequently, removal of the burning ETT must be accomplished as quickly as possible to avoid further thermal and chemical damage to the airway. Remaining flames on the removed ETT or on tube fragments that have fallen off during extubation must be extinguished immediately to prevent ignition of surgical drapes or similar inflammable material. Equally important, the surgeon should instantaneously withdraw the laser’s fiberoptic delivery system and the endoscope (if used for transendoscopic laser application) and irrigate the surgical site with sterile saline or another isotonic aqueous solution to prevent any further smoldering of tissue or tube fragments left in the airway. For these reasons, aqueous solutions must be immediately available to anesthesiologist and surgeon alike during laser surgery in the airway. If there are no obvious tube fragments left in the airway or other material obstructing the airway, and following appropriate suctioning of the airway, the patient may be reintubated and ventilated with a gas mixture containing an O2 concentration not higher than necessary to ensure adequate oxygenation of the patient. Following these immediate steps, larynx and tracheobronchial tree should be thoroughly inspected using an endoscope to evaluate the damage that occurred during the airway fire. The findings obtained during this examination will determine how to proceed with secondary measures and subsequent postoperative management of the patient (Table 3).
References
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Williams & Wilkins, 1999; 557-675. 24. Wainwright AC, Moody RA, Carruth JA. Anaesthetic safety with the carbon dioxide laser. Anaesthesia 1981; 36:411-414. 25. Sosis M. Anesthesia for laser surgery. Adv Anesth 1989; 6:175-194. 26. Dyer RF, Esch VH. Polyvinyl chloride toxicity in fires. Hydrogen chloride toxicity in fire fighters. JAMA 1976; 235:393-397. 27. Boyd CH. A fire in the mouth. A hazard of the use of antistatic endotracheal tubes.Anaesthesia 1969; 24:441-446. 28. Keon TP. Anesthetic considerations for laser surgery. Int Anesthesiol Clin 1988; 26:50-53. 29. LeJeune FE, Guice C, LeTard F, et al. Heat sink protection against lasering endotracheal cuffs. Ann Otol Rhinol Laryngol 1982; 91:606-607. 30. Spiess BD, Ivankovich AD. Anesthetic management of laser airway surgery. Seminars in Surgical Onchology 1990; 6:189-193. 31. Mama KR. Anesthetic Management of the Horse: Intravenous Anesthesia. In: Steffey EP, ed. Recent advances in anesthetic management of large domestic animals. 32. Ossoff RH. Laser safety in otolaryngology - head and neck surgery: anesthetic and educational considerations for laryngeal surgery. Laryngoscope 1989; 99:1-26. 33. Hermens JM, Bennett MJ, Hirshman CA. Anesthesia for laser surgery. Anesth Analg 1983; 62:218-229. 34. Padfield A, Stampi JM. Anaesthesia for laser surgery. Eur J Anaesthesiol 1992; 9:353-366. 35. Heine P, Axhausen M. Anaesthesie und Laserchirurgie im Hals-Nasen-Ohrenbereich. Anaesthesist 1988; 37:10-18.
All rights reserved. This document is available on-line at www.ivis.org. Document No. A0617.0403.
In: Recent Advances in Anesthetic Management of Large Domestic Animals, E.P. Steffey (Ed.) Publisher: International Veterinary Information Service (www.ivis.org), Ithaca, New York, USA. Use of a Helium/Oxygen Carrier Gas Mixture for Inhalation Anesthesia during Laser Surgery in the Airway of the Horse (23-Apr-2003) B. Driessen, L. E. Nann and L. Klein
Department of Clinical Studies-New Bolton Center, School of Veterinary Medicine, University of Pennsylvania, Kennett Square, PA, USA.
Summary The use of lasers during surgery in the airway of the horse bears various hazards of which airway combustion is the most severe. This risk is of particular concern in anesthetized animals breathing an oxygen-rich gas mixture. Though total intravenous anesthesia has been commonly advocated for laser procedures, laser surgery in the airway of the horse can be performed more safely under inhalation anesthesia, if the inspired oxygen concentration (FiO2) is kept low. Use of a helium-oxygen (He/O2) gas mixture and low FiO2 (< 0.4) offers the advantage of significantly decreasing the risk of airway combustion while maintaining adequate arterial oxygenation in the majority of equine patients.
Introduction Over the past decade the laser has become a standard tool for upper airway surgery in the horse [1,2]. While offering numerous advantages, the use of lasers in the airway is not without any risk. The potential hazards, particularly that of airway combustion, during laser surgery in the airway of the anesthetized horse have been reviewed in detail and evidence has been provided that the carrier gas used for inhalation anesthesia is important [3]. It is generally recommended to use the lowest concentration of inspired oxygen (FiO2) that is compatible with adequate arterial blood oxygenation [4-6]. Usually a fraction of inspired O2 of 40% or less (FiO2 < 0.4) will achieve this goal, also in horses. Either nitrogen (N2) or helium (He) may be used to dilute the oxygen [5]. A description of an inhalation anesthetic technique using a helium/oxygen carrier gas mixture (He/O2) for horses undergoing airway laser surgery follows.
Rationale for Use of Helium versus Nitrogen or Air Helium is an odorless, tasteless and inert gas, and offers some important advantages over N2. Not only does He not support combustion, but due to its lower density, as compared to N2, it also produces less turbulent gas flow, which may prove advantageous in a partially obstructed airway [7,8]. Furthermore, He compared to N2, is characterized by a greater thermal conductivity and may thus act as a "heat sink" in the event of an airway fire [9]. In 1985, Pashayan and Gravenstein [10] demonstrated that 60% He retards polyvinyl chloride (PVC) tube ignition by CO2 lasers significantly longer than 60% N2. At the same time the authors confirmed an earlier report showing that an inspired O2 concentration below 40% prevents tube fires [11]. Based on these results, an anesthetic protocol for laryngotracheal operations with CO2 lasers was developed using an inspired He concentration of 60% or more and FiO2 < 0.4 [12]. Pashayan et al., [12] did not report any incidence of airway fire or tracheal tube burns in 523 clinical patients treated with this "helium protocol". This finding coincided with an earlier report suggesting an inspired gas mixture of 30% O2 in He as the safest for use in laser surgery of the airway [13]. Similarly, at the authors’ institution at the large animal hospital there have been no incidents of laser combustion of the endotracheal tube (ETT) or airway fire in more than 300 horses that underwent inhalation anesthesia with a He/O2 gas mixture (FiO2 < 0.4) for laser surgery in the airway. Thus, it is fair to conclude that use of He/O2 avoids the necessity for employing shielded ETTs in anesthesia for airway laser surgery [6].
Technical Requirements for use of He/O2 with Common Large Animal Anesthesia Machines Helium/oxygen gas mixtures can be ordered from standard medical gas suppliers. At the authors’ institution, a gas mixture of 30% O2 and balanced helium 70% (Heliox) is used. The Heliox gas mixture is supplied in "H" size tanks (about 7000 L gas volume) at a cost of approximately $70.00, with each tank lasting for about 15 - 25 surgeries dependent on the duration of the laser procedure and the Heliox gas flow. At the start of inhalant anesthesia, the authors usually choose a Heliox gas flow of about 5 - 6 L/min and an O2 flow of about 1 - 2 L/min. If the inspired O2 concentration exceeds 40%, the O2 flow is gradually
reduced till a FiO2 < 0.4 is reached. The Heliox gas flow may later be reduced to lower flow rates of 3 - 4 L/min with appropriate reduction of the O2 flow to meet the goal of the lowest inspired O2 concentration necessary to ensure adequate oxygenation of the patient. Use of the Heliox gas mixture requires only minor technical modifications that can be carried out without great difficulty on most large animal anesthesia machines. Next, we briefly describe technical modifications made on a Dräger Large Animal Control Center (North American Dräger, Telford, PA) and a Mallard Model 2800 Large Animal Anesthesia Machine (Mallard Medical Inc., Redding, CA).
Large animal anesthesia machines can be fitted with a separate Porter style flow meter (Fig. 1b) for controlling the flow of the He/O2 gas mixture. In the simplest case, such as the Dräger unit (Fig. 1a), one might only replace the flow tube from an already existing nitrous oxide flow meter with a flow tube calibrated for Heliox gas mixture (30% O2, balanced helium; Fig. 1b). The flow tube is commercially available from Dräger Medical, Inc. (formerly North American Dräger). Some large animal anesthesia machines like the Mallard unit (Fig. 1c) are not equipped with two flow meters and thus require the installation (potentially by the manufacturer) of a second flow meter with a flow tube calibrated for the He/O2 (70/30%) gas mixture. The tubing that originates at the Heliox flow meter joins tubing from the oxygen flow meter via a Y-piece connector (see Fig. 1d for gas tube assembly). This allows both the blending of the Heliox gas with 100% O2 to achieve in the breathing circuit a higher FiO2 to ensure adequate oxygenation of the individual patient. It also allows rapid return to 100% O2 as the sole carrier gas once the laser is not anymore in use. To supply the Heliox flow meter, an "H" tank of Heliox gas mixture is fitted with a pressure-reducing regulator (2000 psig [13.789MPa] reduced to 50 psig [345 kPa]) and then connected to the back of the anesthesia machine using a high pressure hose (100 psig [680 kPa] rating) and DISS (diameter index safety system) fitting designated for "special" mixed breathing gases.
Figure 1. Technical adaptations for use of a gas mixture of 30% O2 and balanced helium 70% (Heliox) with common large animal anesthesia machines. The Dräger Large Animal Control Center (A) can be adapted by replacing the Thorpe tube of an existing N2O flow meter with a He/O2 (70/30%) flow tube for controlling the flow of the He/O2 gas mixture. In contrast, the Mallard Model 2800 Large Animal Anesthesia Machine (C) in its standard version is not equipped with two flow meters, thus requiring mounting of a second Porter style flow meter (B) calibrated for use with He/O2 (70/30%). On the back of the Mallard unit a y-piece adapter connects tubing of the Heliox flowmeter with the oxygen fresh gas flow line (D). - To view this image in full size go to the IVIS website at www.ivis.org . -
In order to recognize delivery of a hypoxic gas mixture to the patient, a commercially available galvanic oxygen sensor (Fig. 2) should be mounted on the anesthesia machine, unless other multigas and anesthetic agent monitors with oxygen sensor function are available. These oxygen cells (also called FiO2 sensors) will allow continuous monitoring of the inspired O2 concentration. Since standard FiO2 sensors do not easily adapt to large animal anesthesia circuits, custom adaptations must be made to allow for accurate FiO2 measurement. The simplest solution is to drill a hole in the top of the inspiratory dome valve and glue on an appropriately sized PVC tube that matches the oxygen sensor to be used (Fig. 2 A-C). As a part of the anesthesia machine checkout, the FiO2 analyzer is calibrated first with room air (21% O2; Fig. 2D) and then with 100% O2. Once calibrated, the Heliox gas mixture is checked to ensure a fraction of 30% O2. When properly calibrated, these standard oxygen sensors are quite accurate, although a modest drift over time can occur in high humidity conditions.
Figure 2. Measuring the oxygen fraction in the inspired gas mixture (FiO2) during inhalation anesthesia with helium/oxygen gas mixtures. To use standard galvanic oxygen cells in large animal anesthesia circuits, modifications may be necessary. For example, a spare acrylic glass dome (A) may be equipped with a PVC tube adaptor (a). An oxygen sensor probe (FiO2 sensor) can then be attached to the inspiratory limb of the circuit (B-C) for continuous measurement of the inspired oxygen concentration. When properly calibrated, standard oxygen sensors are quite accurate (21% O2 at room air; see D). - To
view this image in full size go to the IVIS website at www.ivis.org . -
Additional Monitoring Required for Safe use of Low O2 Gas Mixtures During periods of low FiO2 inhalation anesthesia careful monitoring of the horse’s hemodynamic and respiratory functions is critical in order to detect signs of hypoxemia as early as possible and to respond appropriately. Besides FiO2 monitoring, continuous pulse oximetry and intermittent arterial blood gas analysis are important means to accurately evaluate the patient’s respiratory function and oxygenation status. A portable blood gas analyzer (e.g., AVL OPTI CCA, Roche Diagnostics, Indianapolis, IN ; i-STAT system, Sensor Devices Inc., Waukesha, WI) stationed in or in close proximity to the operating
room facilitates immediate blood gas analysis if needed.
Experiences with a Helium/oxygen Carrier Gas Mixture (He/O2) in Inhalation Anesthesia for Horses Undergoing Airway Laser Surgery Based on extensive experience with He as the primary carrier gas in inhalation anesthesia for horses, the authors believe that an inhalant anesthetic protocol employing a He/O2 gas mixture with low FiO2 (< 0.4) offers a safe and potentially advantageous alternative to TIVA, which is generally more frequently advocated for upper airway laser surgery. Arterial blood gas results as well as hemodynamic data (Table 1 and 3) from more than 100 clinical patients anesthetized with volatile anesthetics (isoflurane; sevoflurane), under those conditions for various laser procedures in the upper airway, support this conclusion. Furthermore, in more than 10 years of routine practice of laser surgery in the airway of horses at the University of Pennsylvania’s Large Animal Hospital, no case of endotracheal tube combustion or any other incidence of airway fire has been reported, though incomplete punctures of endotracheal tubes by misdirected laser beams have been occasionally encountered.
Anesthesia was performed in all horses, whose data are presented in Table 1 and Table 3, following a routine inhalant anesthetic protocol with only minor adaptations mentioned below. Necessary technical modifications of the anesthesia machine that allows use of Heliox as the primary carrier gas have been already described. After premedication with either one or a combination of acepromazine, butorphanol, xylazine, and detomidine, anesthesia was induced with clinically common drug combinations of guaifenesin and/or diazepam plus ketamine or thiopental. Immediately following nasotracheal (most commonly) or orotracheal intubation, horses were placed in lateral (most frequently) or dorsal recumbency. The endotracheal tube was connected to an anesthetic rebreathing circuit, from which the animals were immediately breathing a He/O2 gas mixture low in FiO2. Anesthesia was maintained with either isoflurane or sevoflurane in helium and oxygen, using an initial total fresh gas flow (i.e., Heliox plus O2) of 6 - 8 L/min that was changed to a lower total fresh gas flow rate of 4 - 6 L/min if the surgical procedure lasted longer than 30 - 45 min. Blending of Heliox with 100% O2 was aimed to yield an initial FiO2 of 0.3 - 0.4, as measured with the oxygen sensor built in the inspiratory limb of the anesthetic circuit. Standard instrumentation for monitoring of hemodynamic, respiratory and oxygenation parameters was applied to all patients and included ECG, pulse oximetry, FiO2 and peak airway pressure recording, side-stream capnography, and invasive blood pressure and body temperature recording. Provided hemodynamic parameters were within acceptable range, intermittent positive pressure ventilation was instituted soon after completion of instrumentation or as soon as hemodynamic support therapy with IV fluids, inotropes (dobutamine or ephedrine), and occasionally vasopressors (phenylephrine) took effect. Previous experience indicated that mechanically ventilated horses were less likely to be hypoxemic than those breathing spontaneously. Arterial blood samples for blood gas analysis were drawn after catheterization of an artery, i.e., usually before the first activation of the laser instrument. This allowed fine adjustment of both FiO2 and mechanical ventilation prior to beginning the laser procedure, particularly in those cases in which blood gas data revealed hypoxemia and/or hypoventilation. Besides careful monitoring of the pulse oximeter signal, repeated blood gas analysis during the period of laser use was employed to detect changes in respiratory function, allowing for rapid corrective measures if needed. Following conclusion of the laser procedure, which lasted usually between 30 and 60 minutes, surgery proceeded with conventional techniques. At this time, the Heliox gas was turned off and inhalation anesthesia continued under low flow conditions (2.5 - 4 L/min) with 100% O2 as the sole fresh gas source.
The anesthetic records of 108 normal size and 25 draft horses anesthetized between 1999 and 2003 for laser surgery in the upper airway were reviewed (Table 1 and Table 2). Results from arterial blood gas analyses in these isoflurane anesthetized and mechanically ventilated horses revealed that use of a He/O2 gas mixture low in FiO2 (< 0.4) allows adequate oxygenation in the majority of equine patients, at least for the limited time of Heliox application (usually no longer than 60 - 90 min). Nevertheless, arterial oxygen saturation (SaO2) decreased in 2 out of 108 normal sized and 1 out of 25 draft horses below 90%, stressing the necessity for continuous monitoring for signs of oxygen desaturation.
As was to be expected, arterial partial pressures of oxygen (PaO2) were on average lower in draft as compared to normal sized horses, indicating that horses of the giant breeds are at an appreciably higher risk of developing hypoxemia during low FiO2 anesthesia (Table 1 and Table 2).
Minute ventilation (Vmin); inspired oxygen fraction (FiO2); arterial partial pressures of oxygen (PaO2) and carbon dioxide (PaCO2); arterial oxygen saturation (SaO2); arterial standard base excess (SBE). Data recorded during the laser procedure are from 91 male and 17 female horses of various non-draft breeds.
Data recorded during the laser procedure are from 21 male and 4 female draft horses of various breeds
Table 1. Respiratory and arterial blood gas variables in normal size horses anesthetized with isoflurane in He/O2 for upper airway laser surgery.
Parameter FiO2 range n Mean +/- SD Highest Lowest
Age (years) 108 6 +/- 3 15 1
Weight (kg) 108 562 +/- 78 826 332
Vmin (mL/kg/min) 108 81 +/- 10 104 48
FiO2 108 0.37 +/- 0.04 0.49 0.22
pHa0.21 - 0.30 0.31 - 0.40 0.41 - 0.50
3 88 17
7.45 +/- 0. 02 7.42 +/- 0. 05 7.43 +/- 0. 05
7.46 7.51 7.50
7.43 7.28 7.34
PaCO2 (mmHg)0.21 - 0.30 0.31 - 0.40 0.41 - 0.50
3 88 17
46 +/- 6 49 +/- 7 48 +/- 6
52 74 63
41 37 37
PaO2 (mmHg)0.21 - 0.30 0.31 - 0.40 0.41 - 0.50
3 88 17
90 +/- 24 130 +/- 37 139 +/- 53
115 213 246
69 51 61
SaO2 (%)0.21 - 0.30 0.31 - 0.40 0.41 - 0.50
3 88 17
97 +/- 3 98 +/- 2 98 +/- 3
99 100 100
94 83 90
SBE (mmol/L)0.21 - 0.30 0.31 - 0.40 0.41 - 0.50
3 88 17
7.8 +/- 2.4 6.5 +/- 2.3 6.5 +/- 2.3
10.5 11.7 10.2
6.1 2.4 2.7
Table 2. Respiratory and arterial blood gas variables in draft horses anesthetized with isoflurane in He/O2 for upper airway laser surgery.
Parameter FiO2 range n Mean +/- SD Highest Lowest
Age (years) 25 7 +/- 2 14 3
Weight (kg) 25 885 +/- 78 1033 693
Vmin (mL/kg/min) 25 71 +/- 9 87 54
FiO2 25 0.38 +/- 0.03 0.46 0.31
pHa 0.31 - 0.40 0.41 - 0.50
22 3
7.43 +/- 0. 04 7.41 +/- 0. 04
7.49 7.43
7.34 7.37
PaCO2 (mmHg) 0.31 - 0.40 0.41 - 0.50
22 3
45 +/- 5 44 +/- 1
55 44
37 43
PaO2 (mmHg) 0.31 - 0.40 0.41 - 0.50
22 3
83 +/- 31 88 +/- 13
204 101
39 75
SaO2 (%) 0.31 - 0.40 0.41 - 0.50
22 3
95 +/- 4 96 +/- 2
100 98
79 95
SBE (mmol/L) 0.31 - 0.40 0.41 - 0.50
22 3
5.4 +/- 2.1 4.1 +/- 1.7
9.6 5.1
-0.1 2.1
Heart rate (HR); systolic (SAP), diastolic (DAP), and mean arterial pressure (MAP); cardiac output (CO); cardiac index (CI); He/O2 gas flow representing combined flow from Heliox and O2 flow meters; inspired oxygen fraction (FiO2); end-tidal concentration of sevoflurane (SEVOET) and carbon dioxide (PETCO2); minute ventilation (Vmin); peak airway pressure (PAW); arterial partial pressures of oxygen (PaO2) and carbon dioxide (PaCO2); arterial bicarbonate concentration (HCO3-); standard base excess (SBE); and arterial oxygen saturation (SaO2). Values are means +/- SD of 6 Thoroughbred horses (4+/-2 years; 517+/-56 kg), premedicated with IV acepromazine 0.02 mg/kg, butorphanol 0.03 mg/kg, and xylazine 0.3 mg/kg, and then anesthesia induced with IV guaifenesin 0.04 mg/kg, diazepam 0.1 mg/kg, and ketamine 2.0 mg/kg.
Anesthesia in the horse is known to be accompanied by derangements of pulmonary gas exchange leading to impaired arterial blood oxygenation, despite animals breathing oxygen-rich gas mixtures (FiO2 > 0.8) [14-16]. The most important change regarding arterial oxygenation is the development of a large right-to-left intrapulmonary vascular shunt, with blood perfusing unventilated areas of the lung [17]. Therefore significant concern existed whether horses breathing a He/O2 gas mixture low in FiO2 would be able to maintain adequate arterial blood oxygenation. However, the retrospective analysis of blood gas data
Table 3. Hemodynamic, respiratory, and arterial blood gas and acid base variables in horses anesthetized with sevoflurane in He/O2 for airway laser surgery.
Physiological parameter Post induction During laser procedure Post laser procedure
obtained in this large population of clinical patients anesthetized with a He/O2 gas mixture does not seem to support this concern. In fact, previous studies in anesthetized human patients demonstrated that if the use of 100% O2 is avoided during inhalation anesthesia and the FiO2 is maintained at moderately low levels (e.g., 0.4), no or very little atelectasis is produced [18]. Furthermore, changing the inspired gas mixture from room air to 100% O2 in patients showing significant regional ventilation-perfusion (V/Q) mismatching has been shown not to improve blood oxygenation but instead to worsen it due to aggravation of right-to-left shunting [19]. It is thought that the increase in shunt fraction during anesthesia is due to a phenomenon described as absorption atelectasis, which progressively converts poorly ventilated areas of the lung (i.e., units with low V/Q ratio) into atelectatic foci by rapid and complete O2 uptake from those units, which are no longer being ventilated [19]. In humans, atelectatic shunting is effectively reduced during breathing of gas mixtures composed largely of inert gases (e.g., N2 or He), which are minimally absorbed and therefore keep small airways and alveoli open even if underventilated. The same reasons may explain why horses can maintain a reasonably good blood oxygenation while breathing a He/O2 gas mixture with low FiO2. To further substantiate our conclusion that inhalant anesthesia using a He/O2 gas mixture with low FiO2 (< 0.4) can be safely administered in the horse for at least a limited period of time, we conducted also a small prospective study, in which the inspired O2 concentration during sevoflurane in He/O2 anesthesia was kept below 30% (Table 3).
To further substantiate our conclusion that inhalant anesthesia using a He/O2 gas mixture with low FiO2 (< 0.4) can be safely administered in the horse for at least a limited period of time, we conducted also a small prospective study, in which the inspired O2 concentration during sevoflurane in He/O2 anesthesia was kept below 30 % (Table 3). Arterial blood gas and acid base data obtained in this group of horses support the idea that it might be feasible to reduce FiO2 even further below the threshold of 30% of O2 (FiO2 > 0.3), above which the risk of ETT combustion significantly rises [20-22]. Hemodynamic data, including cardiac output (as measured by the lithium dilution technique [23]), showed no clinically significant difference whether measured during periods of low or high FiO2, thus indicating that a lower inspired oxygen concentration is not associated with compromised cardiovascular function.
Admittedly, horses undergoing laser surgery in their upper airway are commonly relatively young and systemically healthy animals. Hence, conclusions as to the safety of an inhalant anesthetic protocol employing a He/O2 gas mixture with low FiO2 (< 0.4) may not apply to a much older or otherwise compromised patient population.
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