Anesthesia and Euthanasia in Zebrafish - PDF

Please download to get full document.

View again

of 9
All materials on our website are shared by users. If you have any questions about copyright issues, please report us to resolve them. We are always happy to assist you.
Information Report
Category:

Novels

Published:

Views: 0 | Pages: 9

Extension: PDF | Download: 0

Share
Related documents
Description
Anesthesia and Euthanasia in Zebrafish Monte Matthews and Zoltán M. Varga Abstract Because of the relative ease of embryonic manipulation and observation, the ability to produce a great number of genetic
Transcript
Anesthesia and Euthanasia in Zebrafish Monte Matthews and Zoltán M. Varga Abstract Because of the relative ease of embryonic manipulation and observation, the ability to produce a great number of genetic mutations, efficient screening methods, and the continued advance of molecular genetic tools, such as the progress in sequencing and mapping of the zebrafish genome, the use of zebrafish (Danio rerio) as a biomedical model organism continues to expand. However, studies involving zebrafish husbandry and veterinary care struggle to keep pace with scientific progress. This article outlines some of the current, acceptable methods for providing anesthesia and euthanasia and provides some examples of how performance-based approaches can be used to advance the relatively limited number of anesthetic and euthanizing techniques available for zebrafish. Introduction Many of the institutions using zebrafish (Danio rerio) for research, testing, or teaching are funded by the Public Health Service (PHS) and accredited by the Association for Assessment and Accreditation of Laboratory Animal Care International. Therefore, these institutions use the Guide for the Care and Use of Laboratory Animals (the Guide; NRC 2011) as a basis for designing, implementing, and evaluating the program for zebrafish care and use, including anesthesia and euthanasia. All institutions that receive PHS funds or support must have a defined policy or PHS Assurance that describes the institution s compliance with the PHS Policy on the Humane Care and Use of Laboratory Animals (PHS 2002) and the Guide. PHS-funded and Association for Assessment and Accreditation of Laboratory Animal Care International accredited zebrafish facilities must also have an institutional animal care and use committee (IACUC) to oversee the animal program, facilities, and animal procedures and to ensure that the institution s program is based on the Guide and PHS Policy. The US Government Principles for the Utilization and Care Monte Matthews is Director of Animal Care Services, and Zoltán M. Varga, PhD, is Director of the Zebrafish International Resource Center at the University of Oregon, Eugene. Address correspondence and reprint requests to Zoltán M. Varga, Zebrafish International Resource Center, 5274 University of Oregon, 1307 Franklin Boulevard, Eugene, OR or of Vertebrate Animals Used in Testing, Research, and Training (IRAC 1985) form the basis of the Guide and can be used by IACUCs to evaluate their program and individual animal use protocols. Consistent with the difficulty of distinguishing between nociception and pain in fish, principle IV of the US Government Principles, which addresses the minimization of discomfort, distress, and pain, states: Unless the contrary is established, investigators should consider that procedures that cause pain or distress in human beings may cause pain or distress in other animals (IRAC 1985, 1). In addition, principle V states: Procedures with animals that may cause more than momentary or slight pain or distress should be performed with appropriate sedation, analgesia, or anesthesia. Surgical or other painful procedures should not be performed on unanesthetized animals paralyzed by chemical agents (IRAC 1985, 1). Because little is known about zebrafish pain, distress, and discomfort, this principle should obviously also be applied when evaluating potentially painful or distressing procedures in zebrafish. The 2011 Guide provides general guidance on determining the appropriate method of anesthesia and euthanasia. According to the Guide: The selection of appropriate analgesics and anesthetics should reflect professional veterinary judgment as to which best meets clinical and humane requirements as well as the needs of the research protocol. The selection depends on many factors, such as the species, age, and strain or stock of the animal, the type and degree of pain, the likely effects of particular agents on specific organ systems, the nature and length of the surgical or pain-inducing procedure, and the safety of the agent (NRC 2011, 121). For evaluating appropriate euthanasia methods, some of the criteria that should be considered are the ability to induce loss of consciousness and death with no or only momentary pain, distress, or anxiety; reliability; irreversibility; time required to induce unconsciousness; appropriateness for the species and age of the animal; compatibility with research objectives; and the safety of and emotional effect on personnel (NRC 2011, 123). These criteria outlined in the Guide are based on those described in the American Veterinary Medical Association (AVMA) (2007) Guidelines on Euthanasia and are specified in the next paragraph. The specific agents and methods chosen for zebrafish euthanasia will depend upon age and the scientific objectives described in the IACUC animal care and use protocol (NRC 2011). Both the PHS Policy and the Guide require the methods of euthanasia to be consistent with the AVMA Guidelines on Euthanasia. These guidelines were extensively revised in 192 ILAR Journal 2011, approved by the AVMA Executive Board in September 2011, and are undergoing final edits (Nolen and AVMA Executive Board 2011). In evaluating various methods of euthanasia, the 2007 AVMA guidelines use the following criteria: (1) ability to induce loss of consciousness and death without causing pain, distress, anxiety, or apprehension; (2) time required to induce loss of consciousness; (3) reliability; (4) safety of personnel; (5) irreversibility; (6) compatibility with requirement and purpose; (7) emotional effect on observers or operators; (8) compatibility with subsequent evaluation, examination, or use of tissue; (9) drug availability and human abuse potential; (10) compatibility with species, age, and health status; (11) ability to maintain equipment in proper working order; and (12) safety for predators/scavengers should the carcass be consumed (AVMA 2007, 3). Whatever method is chosen for zebrafish euthanasia, death must be confirmed by examining the animal for cessation of vital signs (AVMA 2007, 4). Indicators of Discomfort, Distress, and Pain in Fish Humans are able to communicate feelings, subjective states of (dis)comfort, distress, or pain, including the level of intensity of these states. In contrast, animals cannot communicate these states directly and subjectively, so it is necessary to rely on interpretation and knowledge of the animals species-specific behavior to assess their condition. Specifically, techniques for remote biosensation of a fish s state of well-being are limited at best and depend on an interpretation of behavior or a measuring of its physiological responses (Ross and Ross 1999). Thus, understanding of stress or pain in aquatic animals is complicated and indirect. More direct pain and/or stress assessments include netting or other handling of fish to measure, for example, plasma or whole-body cortisol levels (Ramsay et al. 2009a; Schreck 2000). This, however, introduces additional stressors and variables and complicates the differentiation between the specific source of discomfort or pain and the direct netting and/or handling. In spite of these difficulties, there are several physiological and behavioral assays available to establish a normal state versus a state of distress or pain in fish (Ross and Ross 1999; Sneddon 2009). A stressful event can be defined as an abnormal environmental stimulus or threat that jeopardizes survival or negatively interferes with normal activities or physiological balance (homeostasis). For reasons of simplicity, we distinguish short-term (acute) and long-term (chronic) stressors even though stress in fish is a far more complex phenomenon and the term stress is neither well defined nor consistently used (Harper and Wolf 2009; Ramsay et al. 2006, 2009a). The stress response in fish can be generally categorized into three physiological stages: primary, secondary, and tertiary (Casebolt et al. 1998; Iwama et al. 2004; Wedemeyer 1996; Wedemeyer and McLeay 1981; Wedemeyer et al. 1990). In addition, a fish s behavioral response can sometimes be immediately observed when a stressor is introduced. The behavioral response may be to avoid or mitigate the stressor (Schreck et al. 1997). Both behavioral and physiological responses are intimately related (Iwama et al. 2004). The primary physiological response involves the release of stress hormones into the blood stream. Activation of the hypothalamic-pituitary-interrenal axis involves release of cortisol from interrenal cells located in the head kidney. These cells are activated by adrenocorticotropic hormone, which is released from the pituitary gland. The pituitary gland is stimulated by corticotrophin-releasing factor from the hypothalamus (Donaldson 1981; Wedemeyer et al. 1990). Activation of the sympathetico-chromaffin system (Casebolt et al. 1998) involves release of catecholamines (e.g., adrenaline, noradrenaline) from chromaffin tissue of the head kidney after direct stimulation by the sympathetic nervous system (Mazeaud and Mazeaud 1981; Wedemeyer et al. 1990). The secondary response is composed of various changes to blood and tissue, such as elevated blood sugar levels, changes in electrolyte concentrations, such as hypochloremia, changes in hematology, such as reduced clotting times, and changes in the differential leucocyte count (Wedemeyer et al. 1990, 453). In addition, there may be changes in the tissues, including depletion of liver glycogen and interrenal vitamin C, hemorrhage of the thymus, and hypertrophy of the interrenal body (Wedemeyer et al. 1990, 453). Tertiary responses, for individuals and populations, include reduction in growth, reproduction, resistance to disease, and survival (Casebolt et al. 1998; Iwama et al. 2004; Wedemeyer et al. 1990). Physiological indicators of stress resulting from stressful stimuli are, for example, the skipping of heartbeats and hematological effects such as changes in blood plasma ion composition (osmoregulation), swelling of erythrocytes, and dilution or concentration of heme (Ross and Ross 1999). Cortisol is a useful readout of the overall hormonal response of the hypothalamic-pituitary-interrenal axis. The increase of whole-body cortisol in response to acute and chronic stressors has been shown to impact the zebrafish immune system, leading to increased susceptibility to mycobacteria infections (Ramsay et al. 2006; Ramsay et al. 2009a,b). In larger species, blood plasma cortisol levels can be used instead. However, the analysis of such indicators depends on the establishment of a consistent and reproducible baseline that characterizes the normal state of an indicator. This normal state can fluctuate based on species, circadian rhythm, season, water quality, and other environmental factors (Barton et al. 2002; Iwama et al. 2004) and a stressed state may therefore exist or not depending on whether or not the stress indicator readout represents a true departure from the baseline at the time of testing. A previous special issue of the ILAR Journal (2009, vol. 50, issue 4) was devoted to the practical, philosophical, and scientific aspects of whether or not fish can sense or feel pain, as well as their use as biomedical research organisms (Posner 2009). Pain, in addition to being accompanied by stress, can be defined as a noxious stimulus associated with Volume 53, Number an evasive behavioral response and potential or actual injury and tissue damage (Harper and Wolf 2009; Sneddon 2009). Several internal or physiological criteria support the fact that fish are capable of perceiving noxious stimuli or pain; for example, the existence of opiate neurotransmitters and neural connectivity in fish is comparable to the neurocircuitry that mediates pain in mammals (Gonzalez-Nunez and Rodriguez 2009). However, because it is unclear whether analogous or homologous evolutionary mechanisms could have given rise to similar structures, the debate about pain perception in fish continues to be unresolved, even though analogous evolution is less likely based on molecular data (Gonzalez-Nunez and Rodriguez 2009). Several external behavioral signs can be used to differentiate between a normal and distressed state, including nociception and pain: excessive movement of the opercula (tachyventilation, hyperventilation), cough rate (reversal of water flow direction over the gills), color (pigmentation) changes, and increased movement (hypertaxia) or cessation of movement (ataxia) (Ross and Ross 1999) are useful indicators of stress in some species. However, species-specific differences do exist, making a generalized interpretation of several stress indicators difficult. Interestingly, when comparing opercular movements, use of cover, and swim rates, zebrafish share fewer stress indicators with the closely related carp (Cyprinus carpio) and more with the distantly related salmonid, trout (Oncorhynchus mykiss). Trout and zebrafish respond to a noxious stressor with increased ventilation, increased use of cover, and a decreased swim rate, whereas, based on these indicators, carp appear to be rather indifferent to the same stressor. Carp, however, respond with several anomalous (or, perhaps, species-specific) stress indicators such as off-balance swimming, loss of equilibrium, or rubbing of lips against the aquarium wall (Reilly et al. 2008). Methods of Zebrafish Analgesia, Sedation, and Anesthesia In fish, species-specific modes of respiration play a key role for the route of anesthetic administration. For example, injection of anesthetics is more efficient for species that are capable of piping, or performing surface aerial respiration under hypoxic conditions (Bruecker 1993). Zebrafish are cyprinids, and as such, they can resort to surface respiration under hypoxic conditions (University of Oregon 2007; Varga et al., unpublished study). Under normal conditions, however, they are obligatory gill breathers. Therefore, the most widely used method to anesthetize (Harper and Lawrence 2011; Westerfield 2007) or sedate (Trevarrow 2007, 2001) zebrafish is immersion, which, in fish, is analogous to inhalant anesthesia in terrestrials (Neiffer and Stamper 2009). As with the assessment of discomfort and pain by behavioral observations, levels of sedation or anesthesia can also be characterized by several behavioral criteria. For fish specifically, six levels of anesthesia have been described, and of these, opercular movement is probably the most significant indicator to assess levels of anesthesia (McFarland and Klontz 1969). During light sedation (level 1), reaction to some external stimuli (visual and tactile) is slightly reduced, whereas under deep sedation (level 2), fish have slightly reduced opercular movement rates and do not react to any external stimuli except pressure. This is followed by partial loss of equilibrium (level 3), during which muscle tone is decreased, swimming is erratic, and the opercular rate is increased. At this level, fish still continue to react to strong tactile and vibrational stimuli. At the next level (level 4), however, equilibrium and muscle tone are completely lost, the opercular movement rate decreases again, and fish respond only to strong pressure as an external stimulus. After more prolonged anesthetic exposure (level 5), opercular movements become shallow and the heart rate is decreased. There is no reaction to external stimuli and all reflex reactivity is lost. Finally, the last stage (stage 6) is described as medullary collapse: opercular movement completely stops and the heartbeat is shallow and almost completely absent. If the anesthetic is not removed and attempts are not made to revive the fish, death follows due to cardiac arrest and hypoxia (McFarland and Klontz 1969). These levels of fish anesthesia are, to some degree, similar to the stages observed in mammals. However, deviations may exist from these criteria. For example, even after opercular movements have ceased in tricaine methanesulfonate anesthetized zebrafish, a hard knock on the work surface can still elicit an occasional, single-jolt flight reflex (Varga et al., unpublished study), indicating that some reflexes, and hence some neural activity, persist. Ethyl 3-Aminobenzoate Methanesulfonic Acid Tricaine methanesulfonate, also known as TMS, MS-222, Tricaine mesylate, Finquel (Argent Laboratories, Redmond, Washington), and Tricaine-S (Western Chemical, Inc., Ferndale, Washington), is currently the only anesthetic approved for some aquatic species by the US Food and Drug Administration (FDA) and the most widely used sedative and anesthetic for zebrafish. MS-222 is a muscle relaxant that blocks sodium and to a lesser degree potassium currents in nerve membranes. Action potentials as well as spontaneous contractions of muscles are eliminated by MS-222, including sensory input and reflexes (Frazier and Narahashi 1975). MS-222 is available as a water-soluble powder; however, it is acidic and lowers the ph of unbuffered or weakly buffered solutions to as low as 5. Some studies suggested that the anesthetic effect of MS-222 is reduced under acidic conditions in several fish species (Gilderhus et al. 1973) but not in channel catfish (Welker et al. 2007). Welker and colleagues (2007) also showed that following buffered or unbuffered MS-222 exposure, the plasma cortisol levels remained essentially unaffected in catfish. In these studies, plasma cortisol and glucose levels also increased with higher MS-222 concentrations, suggesting that the stress responses resulted 194 ILAR Journal from the anesthetic rather than the ph of exposure water. In general, however, the lowering of water ph by MS-222 increases the risk of plasma acidosis in fish (Harper and Lawrence 2011) and should be avoided. Zebrafish immersed in buffered or unbuffered MS-222 typically respond with hypertaxia and tachyventilation before righting reflex, body, and opercular movements cease (University of Oregon 2007; Wilson et al. 2009; Varga et al., unpublished study). Occasionally, gill bleeding occurs, and some fish do not recover from anesthesia. Presumably this is due to a general, anesthetic-induced stress response that includes acid stress and/or acidosis (in the unbuffered solution), changes in plasma osmolality, and a blood pressure increase that could damage the gills. MS-222 can be neutralized with sodium bicarbonate or sodium hydroxide (Harper and Lawrence 2011) or with Tris buffer (Westerfield 2007). Another side effect of the muscle relaxant is a reduced heart rate, which increases the risk of accidental death during anesthesia. Simultaneous administration of isoflurane and MS-222 in the zebrafish reduces negative physiological effects, such as reduced heart rate, and effectively helps extend anesthesia duration and shorten recovery time (Huang et al. 2010). In zebrafish, and presumably other fish, MS-222 dose response depends on age, size, and metabolic state. An increase in sensitivity to MS-222 (Rombough 2007) was observed in 3- to 9-day larvae, presumably due to changes in liver detoxification activity. Median lethal concentrations for 4- to 7-day zebrafish larvae suggested that key developmental changes, consistent with the maturation of gills (Shadrin and Ozernyuk 2002) and liver (Field et al. 2003; Sakaguchi et al. 2008), occur at this time. The renowned toxicologist and zebrafish scholar von Hohenheim stated: Poison is in everything, and nothing is without poison. The dosage makes it either a poison or a remedy (Borzelleca 2000). Consistent with this observation, MS-222 has been used for zebrafish sedation [0.01 mg/ml (Trevarrow 2007, 2011)], anesthesia [0.168 mg/ml (Westerfield 2007)], and euthanasia [ mg/ml (University of Oregon 2007; Wilson et al. 2009; Varga et al., unpublished study)]. In addition, studies in Tilapia (Oreochromis niloticus, O. aureus, and O. mossambicus) suggested that repeated ane
Recommended
View more...
We Need Your Support
Thank you for visiting our website and your interest in our free products and services. We are nonprofit website to share and download documents. To the running of this website, we need your help to support us.

Thanks to everyone for your continued support.

No, Thanks