Research Note
Mortality associated with using medetomidine and ketamine for general anesthesia in pregnant and nonpregnant Wistar rats
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Lauren M. Callahan, BSc, BVMS1, Simone M. Ross, AdvCert Animal Tech, Cert IV Training and Assessment1, Megan L. Jones, BSc2 & Gabrielle C. Musk, BSc, BVMS, PhD, Dipl ECVAA1,3
Medetomidine and ketamine are injectable drugs that can be used in combination to induce general anesthesia in rats. After noticing a high incidence of morbidity and mortality in pregnant Wistar rats given medetomidine and ketamine for anesthesia, the authors further investigated the effects of this combination of anesthetic drugs in both pregnant and nonpregnant Wistar rats. The time to recumbency and the duration of general anesthesia were similar between pregnant and nonpregnant rats. Pregnancy status did not affect the rats’ pulse rate, respiratory rate, rectal temperature, oxygen saturation or perfusion index during 2 h of anesthesia. Pregnant rats had significantly lower blood glucose concentrations than nonpregnant rats at all time points, though blood glucose concentrations increased in both groups. The mortality rate was ~15% both for nonpregnant rats and for pregnant rats. Researchers using medetomidine and ketamine to anesthetize Wistar rats should carefully monitor the rats in order to minimize mortality. General anesthesia is commonly induced in rats undergoing surgical and non-surgical procedures for research purposes. Either injectable drugs or inhalational drugs may be used to induce general anesthesia, but there are several advantages of using injectable anesthetics rather than inhalational drugs: the administration of injectable drugs is relatively simple, does not require an anesthetic machine, results in a comparatively smooth induction of anesthesia, avoids the risk of waste anesthetic gas leakage, is usually more cost-effective than use of inhalants and has reversible effects in some cases1,2. Furthermore, certain inhalational anesthetics may interfere with primary physiological endpoints. For example, because preconditioning with inhalational anesthetics such as isoflurane can confer protection from ischemiareperfusion–induced tissue damage in heart3, kidney4 and brain5, inhalational anesthetics may confound ischemia-reperfusion injury studies.
Medetomidine and ketamine are injectable drugs that can be used in combination to induce general anesthesia in rats6,7. Medetomidine is an α2adrenoreceptor agonist that produces sedation and analgesia, and ketamine is a dissociative anaesthetic1. This combination of drugs can induce surgical anesthesia for 30–60 min and unconsciousness for 60–120 min in rats1. The specific α2-adrenoreceptor antagonist atipamezole can be used to reverse the effects of medetomidine, inducing recovery from anesthesia within a few minutes of its administration. Reported mortality rates associated with medetomidine and ketamine anesthesia of Wistar rats vary. Roughan et al.8 reported 56% mortality in rats that were premedicated with buprenorphine and then administered various doses of medetomidine and ketamine in combination. In contrast, Hedenqvist et al.9 and Moriondo et al.10 reported no mortality associated with the drug combination, despite using doses
1Animal Care Services, University of Western Australia, Crawley, Australia. 2School of Anatomy, Physiology and Human Biology, University of Western Australia, Crawley, Australia. 3School of Veterinary and Life Sciences, Murdoch University, Murdoch, Australia. Correspondence should be addressed to G.C.M. ([email protected]).
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as high as 0.5 mg per kg body weight medetomidine and 75 mg per kg body weight ketamine. Although the use of medetomidine and ketamine in nonpregnant rats is widespread, there is little evidence to demonstrate their safety and efficacy in pregnant rats. A study done at the University of Western Australia (M.L.J., unpublished observations) used a combination of medetomidine and ketamine in pregnant Wistar rats undergoing laparotomy and clamping of isolated uterine blood vessels to investigate the effect of placental ischemia-reperfusion on pregnancy outcome. When the rats were administered 0.1–0.5 mg per kg body weight medetomidine and 50–60 mg per kg body weight ketamine by the intraperitoneal (i.p.) route at gestational day 17, anesthetic and recovery times were substantially longer than expected, and four of six rats (67%) either died during the procedure or were euthanized owing to post-anesthetic complications. Despite initial investigations into potential contributing factors (such as inadequate monitoring during and after anesthesia, environmental conditions and poor drug administration technique), the cause of the high morbidity and mortality could not be determined. In light of these preliminary findings, we hypo thesized that pregnant Wistar rats administered medetomidine and ketamine might have higher morbidity and mortality than nonpregnant Wistar rats. We administered a single dose of medetomidine and ketamine by the i.p. route to both nonpregnant and pregnant Wistar rats, measured the duration of anesthesia and monitored several parameters to determine the effect of pregnancy status on the efficacy and safety of this anesthetic combination. METHODS Animals All procedures involving animals were approved by the Animal Ethics Committee of The University of Western Australia in accordance with the guidelines of the Australian Code for the Care and Use of Animals for Scientific Purposes11. We obtained 34 outbred, 8-week-old Wistar (Crl:WI) rats (27 nulliparous females and seven males) from the Animal Resources Centre (Murdoch, Australia). The rats were housed under specific pathogen–free conditions. Rooms were maintained at 22 ± 2 °C and 30–70% humidity on a 12-h: 12-h light:dark cycle. We housed the rats in filter top cages (Expanded Rat Cage, Thoren Caging Systems, Hazleton, PA) lined with aspen bedding (Tapvei, Harjumaa, Estonia). Females were housed in pairs, and males were housed individually. Rats were fed a pellet diet (AIN93G Rodent diet, Specialty Feeds, Glen Forest, Australia) and given ad libitum access to acidified water (pH 2.5–3.0). We tested all rats for specific pathogens in accordance with the recommendations LAB ANIMAL
of ComPath (South Australian Health and Medical Research Institute, Adelaide, Australia). Vendor reports indicated that rats were seronegative for pneumonia virus of mice, Theiler’s encephalomyelitis virus, Sendai virus, mouse adenovirus types 1 and 2, sialodacryoadenitis, rat parvovirus, lymphocytic choriomeningitis, Hantaan virus and reovirus-3 and that rats were free of bacterial and parasitic infections at the time of shipment. All rats were acclimatized to the animal facility for 1 week prior to our studies. The female rats were divided into two groups that were time-mated 6 weeks apart. We synchronized the estrus cycles of the female rats in each group by administering an i.p. injection of 5.7 IU serum gonadotropin (Folligon, Intervet, Bendigo East, Australia) 4 d prior to mating and an i.p. injection of 1.93 IU chorionic gonadotropin (Chorulon, Intervet, Bendigo East, Australia) 2 d prior to mating. On the mating day, we placed two females in a male’s cage overnight. We removed the females from the male’s cage the following morning; this was designated gestational day 0. Pregnancy was confirmed by abdominal palpation on gestation day 14. Female rats were allocated to one of two groups according to their pregnancy status: nonpregnant (n = 13) and pregnant (n = 14). Personnel were not blinded to the pregnancy status of the rats. Anesthesia and monitoring We anesthetized nonpregnant rats and pregnant rats on gestational day 17. All procedures were conducted between 8.00 a.m. and 4.00 p.m. We transported all rats in their home cages to the surgical theater. Temperatures in the surgical theater were maintained at 22 ± 2 °C. We weighed the rats using an electronic balance with an accuracy of 1 g (PGW Precision Balance, Adam Equipment, Milton Keynes, UK). Pregnant rats weighed 349 ± 18 g on average, and nonpregnant rats weighed 273 ± 25 g on average. We administered a single i.p. injection of 0.2 mg per kg body weight medetomidine (1 mg/ml; Domitor, Zoetis, West Ride, Australia) mixed with 50 mg per kg body weight ketamine (100 mg/ml diluted to 0.2 mg/ml in 0.9% sterile saline; Parnell Laboratories, Alexandria, Australia) by inserting a 0.5-in, 26-gauge needle (Precision Glide, BD, Singapore) to a depth of 5 mm at an angle of 30–40° into the caudal right abdominal region. The same investigator (L.M.C.) administered each injection. Drug solutions were prepared each morning, and unused portions were discarded at the end of the day. Following the injection, each rat was placed into an individual cage until it lost its righting reflex. We then transferred each rat to a pre-warmed heating pad (Redflex Pet Recovery Pad; DLC Australia, Hoppers Crossing, Australia) to maintain body temperature between 37.0 °C and 38.0 °C, and we Volume 43, No. 6 | JUNE 2014 2 09
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FIGURE 1 | Mean pulse rate (beats per min) during anesthesia. Open symbols, nonpregnant rats; filled symbols, pregnant rats. Error bars, s.d.
applied ophthalmic ointment (Lacrilube; Allergan, Jersey City, NJ) to the eyes to prevent corneal desiccation. We did not provide oxygen supplementation; the rats breathed room air. We recorded the time to recumbency, defined as the time from the injection of the anesthetic to the loss of the righting reflex, for each rat. We considered the rat to be anesthetized when it failed to exhibit a withdrawal reflex in response to hind limb and forelimb toe pinches, a routine indicator of surgical anesthesia. We obtained a 0.05-ml blood sample from the lateral tail vein12 before anesthetics were injected and 20 min, 30 min, 60 min, 90 min and 120 min after anesthetics were injected. We measured blood glucose concentration of each sample immediately after the sample was obtained, using a glucometer (ACCU-Chek Performa, Roche Diagnostics, Castle Hill, Australia). We recorded pulse rate, respiratory rate, rectal temperature, oxygen saturation (SpO2) and perfusion index (PI) every 10 min. We measured the pulse rate by positioning a Doppler probe (Doppler Flow Detector Model 811-B, Parks Medical Electronics, Aloha, OR) over the coccygeal artery and counting the audible pulses over a period of 15 s. We determined the respiratory rate by counting chest excursions over a period of 15 s. We measured rectal temperature using an electronic thermometer by placing the probe at least 1 cm into the rectum; the same thermometer was used for all temperature measurements. We monitored SpO2 and PI using a pulse oximeter (Masimo Radical-7, Masimo Corporation, Irvine, CA) by positioning the probe over the skin flap behind the elbow or at the tail base. Recovery Two hours after anesthetics were administered to the rats, we administered 0.1 mg per kg body weight atipamezole (5 mg/ml diluted to 0.25 mg/ml in 0.9% 210 Volume 43, No. 6 | JUNE 2014
sterile saline; Antisedan; Zoetis, West Ride, Australia) by i.p. injection. We determined the return of the swallowing reflex by challenging the rat with a drop of oral glucose syrup (Queen Fine Foods, Alderley, Australia). We determined that walking ability had returned when the rat was able to maintain itself in sternal recumbency and move around the cage. For each rat, we recorded the duration of surgical anesthesia, defined as the time from the loss of the withdrawal reflex to the return of the withdrawal reflex. Anesthetic mortality was defined as any death that occurred within 24 h of anesthesia. We carried out in-house postmortem gross anatomical examinations of the rats that died. We euthanized pregnant rats 24 h after anesthesia. We made nonpregnant rats available to other researchers, if appropriate. Statistical analysis We assessed the data for normality and made statistical comparisons using SigmaPlot 12.0 software (Systat Software Inc., San Jose, CA). We used unpaired t-tests to determine differences between pregnant and nonpregnant rats. We compared non-parametric data using a Wilcoxon Rank Sum test. We carried out a regression analysis of the relationship between SpO2 and PI values using Sigmaplot 12.0 software. Data are expressed as mean ± s.d. unless otherwise stated. P values < 0.05 were considered statistically significant. RESULTS Efficacy and safety The mean time to recumbency did not differ significantly between nonpregnant (4.68 ± 2.39 min) and pregnant rats (5.68 ± 4.60 min). There were no significant differences between the two groups in mean pulse rates (Fig. 1), mean respiratory rates (Fig. 2), mean rectal temperatures (Fig. 3) or mean SpO2 (Fig. 4) at any time point during the 2-h anesthesia period. 100
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The correlations between PI and SpO2 were similar for the two groups (Fig. 5). The mean blood glucose concentration was significantly higher in nonpregnant rats than in pregnant rats at all time points (Fig. 6). The mean blood glucose concentration increased from the baseline in both groups, peaking at 90 min. Although exact urine output was not measured, we did observe increased urination in both nonpregnant rats and pregnant rats during anesthesia. Recovery and mortality One pregnant rat and one nonpregnant rat died while under anesthesia. The pregnant rat experienced respiratory arrest after 112 min of anesthesia and was euthanized. This rat had an elevated rectal temperature (41.2 °C) for a period of 5 min while under anesthesia. An in-house post mortem examination of this rat had no relevant findings. The nonpregnant rat experienced dyspnea after 100 min of anesthesia and was euthanized. An in-house post mortem examination of this rat had no relevant findings. All of the other rats (12 nonpregnant and 13 pregnant)
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remained anaesthetized for 120 min until atipamezole was administered. All but one of the rats (11 nonpregnant and 13 pregnant) recovered from anesthesia within a few minutes of receiving atipamezole. One nonpregnant rat did not respond to atipamezole after 40 min and was euthanized. An independent necropsy of this rat, carried out by a pathologist at Murdoch University Veterinary Hospital (Australia), detected pulmonary hyperemia with alveolar hemoglobin crystals, but no other lesions were noted. One pregnant rat was found dead 22 h after anesthesia. This rat had an elevated rectal temperature (40.8 °C) for a period of 5 min while under anesthesia. An in-house post mortem examination of this rat had no relevant findings. The recovery of the remaining rats was uneventful. We immediately made food and water available to the rats, and we observed normal feeding, drinking and behavior. 25
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The overall mortality rate was 14.8%, with a mortality rate of 15.4% among nonpregnant rats and a mortality rate of 14.3% among pregnant rats. DISCUSSION A previous unpublished study done at our institution showed longer than expected anesthetic and recovery times and 67% mortality when medetomidine and ketamine were administered to pregnant Wistar rats for general anesthesia. These findings prompted us to investigate how the effects of medetomidine and ketamine anesthesia differed between pregnant and nonpregnant Wistar rats. Overall, pregnancy status did not affect the animal’s anesthetic depth in response to medetomidine and ketamine, as measured by pulse rate and respiratory rate. The pulse rates observed in this study were predominantly below 300 beats per min, consistent with the known pharmacological ability of medetomidine to decrease the heart rate12. As expected, the respiratory rates of the anesthetized rats were also slightly depressed compared with the normal respiratory rate of 70–115 breaths per min in the rat. Monitoring the depth of anesthesia in rats can be challenging, owing to their small size and high metabolic rate. The normal pulse rate of a rat is 250–450 beats per min13, making it difficult to manually count. Although the Doppler probe that we used was designed for use in animals, it may not be sensitive enough for reliable use in small rodents such as rats. As such, we observed some variability in the pulse rate that may be attributed to human error. The rectal temperature of the rats was also unaffected by pregnancy status. Maintaining the target body temperature (37.0–38.0 °C) during anesthesia in both pregnant and nonpregnant rats was difficult in this study, and rectal temperatures changed rapidly. It is vital during anesthesia that the target body temperature is maintained in order to avoid adverse physiological consequences of hypothermia and hyperthermia. Hypothermia can prolong recovery from anesthesia and can cause myocardial infarctions and coagulopathies, whereas hyperthermia can cause tachycardia, hyperventilation and cardiac arrhythmias14. Rats are especially prone to hypothermia during anesthesia because of their high metabolic rate and large surface-to-body-weight ratio, but hyperthermia is also a risk if an active heating device is used1. Given the temperature fluctuations we observed with this anesthetic regime, we suggest particular attention be paid to monitoring and maintaining body temperature when using medetomidine and ketamine for anesthesia, especially if active heating devices are being used. We did not find any differences in SpO2 or the relationship between SpO2 and PI between pregnant and nonpregnant rats. However, we were unable to 212 Volume 43, No. 6 | JUNE 2014
obtain consistent readings using the pulse oximeter. Despite observing SpO2 values below 90%, compared to normal SpO2 values of >95%, there was no obvious clinical evidence of hypoxemia in these rats. A limitation of our study was that we were unable to carry out arterial blood gas analyses; these would have allowed us to interpret SpO2 in the context of the partial pressure of oxygen in arterial blood. The majority of the SpO2 readings were associated with low PI, which may be due to the peripheral vasoconstrictive effects of medetomidine12. Pregnant rats had significantly lower blood glucose concentrations throughout the anesthetic period than nonpregnant rats, consistent with previous observations in rats15. An increased predisposition to hypoglycemia in mid- to late pregnancy has been demonstrated in other species as well16–18 and is attributed to maternal diversion of glucose to the developing fetus to meet demand during the period of maximal fetal growth19. Both nonpregnant and pregnant rats showed a steady increase in blood glucose concentrations in response to the administration of medetomidine and ketamine, likely owing to the hyperglycemic effects of medetomidine20,21. We suggest it may be prudent for researchers to avoid using medetomidine in studies in which persistent hyper glycemia interferes with their aims. We also observed in both groups of rats that urine output was qualitatively increased during anesthesia. This is a commonly reported side effect of α2adrenoreceptor agonists22–24. We recommend that perioperative fluid therapy be considered when anesthetizing animals with medetomidine, especially if intra-operative hemorrhage is likely. All rats that survived the anesthetic period remained anaesthetized for 2 h and required administration of atipamezole for arousal. This is not consistent with published anesthetic times6–10. Studies have shown that the strain and sex of an animal may affect its response to different injectable anesthetic agents25,26. Responses to equipotent doses of medetomidine vary widely between species owing to differences in α2-adrenoreceptor subtypes, densities and locations within the central nervous system and peripheral tissue27. Sex may also impact the response to medetomidine, because higher estrogen levels can increase the sensitivity of α2-adrenoreceptors27. Therefore, it is recommended when using medetomidine and ketamine that the doses be optimized to determine the safest and most efficacious combination for the particular strain. In addition, atipamezole should always be available to induce arousal. Mortality rate was not associated with pregnancy status, although both of the pregnant rats that died showed a temporary increase in body temperature that may have contributed to their death. The significance www.labanimal.com
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of this complication in this study is unknown. A limitation of this study is that comprehensive necropsy with histopathology was not done on all rats that died. The pulmonary hyperemia and edema detected upon necropsy examination in one nonpregnant rat may have compromised alveolar gas exchange. Given that xylazine may cause pulmonary edema in rats, albeit at high doses, it may be prudent to supplement oxygen in rats receiving α2-adrenoreceptor agonists28,29. Although oxygen supplementation during anesthesia is good practice, we aimed to use only equipment readily available in the laboratory animal research setting, whereas the use of oxygen requires delivery through an anesthetic machine and breathing system. In light of our findings, we urge researchers to provide supplementary oxygen to anaesthetized rats by a facemask if this combination of drugs is to be used for anesthesia. A possible cause of death in the rats that died during the anesthetic period is upper respiratory tract obstruction. Rats recovering from anesthesia must be continuously monitored until their swallowing reflex has returned, because the lack of this reflex poses a risk of choking in the case of any upper respiratory tract obstructions. Due to their small size, it can be difficult to determine when this reflex has returned. We have found that application of glucose syrup to the mouth and tongue after the rats show signs of regaining consciousness (such as head movements) is an effective technique for assessing the swallowing reflex. Other possible causes of anesthetic-related death in our study include cardiac dysrhythmias, cardiac arrest and hypoxia secondary to pulmonary edema or pulmonary vasoconstriction. It would have been useful to measure end tidal carbon dioxide concentrations, electrocardiogram and blood pressure. In future studies, specialized physiological monitoring equipment suitable for small rodents should be used to clarify the nature of anesthetic-related complications. As fetal viability was outside the scope of this study, and the rats were euthanized the following day, fetal viability was not assessed. Therefore, it is not possible to determine whether medetomidine and ketamine anesthesia affected placental perfusion. In principle, drugs such as medetomidine that stimulate the α2adrenoreceptor could increase the contractibility of the uterus in pregnant rats, leading to fetal resorption. The increase in progesterone levels that occur during pregnancy could counteract this effect by stimulating the sensitivity of β-adrenoreceptors, however, which would decrease the contractibility of the uterus27. Medetomidine and ketamine can be used to effectively induce general anesthesia in nonpregnant and pregnant Wistar rats, but this combination of
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a nesthetics resulted in a mortality rate of ~15% regardless of pregnancy status. Anesthetic monitoring equipment suitable for use in small rodents should be used in future studies to help detect physiological complications that may be managed or prevented to reduce mortality associated with these anesthetics. Furthermore, researchers should consider alternative approaches to anesthesia in Wistar rats. If that is not feasible, then we suggest researchers undertake an initial dose optimization study of these drugs for a specific strain of Wistar rat prior to commencing the study. Acknowledgments We thank Leah Attwood (Animal Services, The University of Western Australia) for providing technical support. COMPETING FINANCIAL INTERESTS The authors declare no competing financial interests. Received 22 January 2014; accepted 7 March 2014 Published online at http://www.labanimal.com/ 1. Flecknell, P. Laboratory Animal Anaesthesia 3rd edn. (Academic, Waltham, MA, 2009). 2. Hacker, S.O., White, C.E. & Black, I.H. A comparison of target-controlled infusion versus volatile inhalant anaesthesia for heart rate, respiratory rate, and recovery time in a rat model. Contemp. Top. Lab. Anim. Sci. 44, 7–12 (2005). 3. Stowe, D.F. & Kevin, L.G. Cardiac preconditioning by volatile anesthetic agents: a defining role for altered mitochondrial bioenergetics. Antioxid. Redox. Signal. 6, 439–448 (2004). 4. Hashiguchi, H. et al. Isoflurane protects renal function against ischemia and reperfusion through inhibition of protein kinases, JNK and ERK. Anesth. Analg. 101, 1584–1589 (2005). 5. Steiger, H.J. & Hänggi, D. Ischaemic preconditioning of the brain, mechanisms and applications. Acta Neurochir. (Wien) 149, 1–10 (2007). 6. Nevalainen, T., Pyhälä, L., Voipio, H.M. & Virtanen, R. Evaluation of anaesthetic potency of medetomidine-ketamine combination in rats, guinea-pigs and rabbits. Acta Vet. Scand. Suppl. 85, 139–143 (1989). 7. Jang, H.S., Choi, H.S., Lee, S.H., Jang, K.H. & Lee, M. Evaluation of the anaesthetic effects of medetomidine and ketamine in rats and their reversal with atipamezole. Vet. Anaesth. Analg. 36, 319–327 (2009). 8. Roughan, J.V., Ojeda, O.B. & Flecknell, P.A. The influence of pre-anaesthetic administration of buprenorphine on the anaesthetic effects of ketamine/medetomidine and pentobarbitone in rats and the consequences of repeated anaesthesia. Lab. Anim. 33, 234–242 (1999). 9. Hedenqvist, P., Roughan, J.V. & Flecknell, P.A. Effects of repeated anaesthesia with ketamine/medetomidine and of preanaesthetic administration of buprenorphine in rats. Lab. Anim. 34, 207–211 (2000). 10. Moriondo, A. et al. Impact of respiratory pattern on lung mechanics and interstitial proteoglycans in spontaneously breathing anaesthetized healthy rats. Acta Physiol. (Oxf.) 203, 331–341 (2011). 11. National Health and Medical Research Council, Australian Government. Australian Code of Practice for the Care and Use of Animals for Scientific Purposes 4th edn. (Australian Government, Canberra, Australia, 2004). 12. Plumb, D.C. Plumb’s Veterinary Drug Handbook 5th edn. (Blackwell, Ames, IA, 2005).
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13. Hrapkiewicz, K. & Medina, L. (eds.) Clinical Laboratory Animal Medicine: An Introduction 3rd edn. (Blackwell, Oxford, UK, 2007). 14. Dugdale, A. Veterinary Anaesthesia: Principles to Practice Ch.18 (Wiley Blackwell, West Sussex, UK, 2010). 15. Wharfe, M.D., Mark, P.J. & Waddell, B.J. Circadian variation in placental and hepatic clock genes in rat pregnancy. Endocrinology 152, 3552–3560 (2011). 16. Johnson, C.A. Glucose homeostasis during canine pregnancy: insulin resistance, ketosis, and hypoglycemia. Theriogenology 70, 1418–1423 (2008). 17. Reddi, A.S., Oppermann, W., Strugatz, L.H., Cole, H.S. & Camerini-Davalos, R.A. Effect of pregnancy on serum alanine concentration in normal and genetically diabetic mice. Horm. Metab. Res. 8, 478–482 (1976). 18. Schlumbohm, C. & Harmeyer, J. Twin-pregnancy increases susceptibility of ewes to hypoglycaemic stress and pregnancy toxaemia. Res. Vet. Sci. 84, 286–299 (2008). 19. Herrera, E. Metabolic adaptations in pregnancy and their implications for the availability of substrates to the fetus. Eur. J. Clin. Nutr. 54, S47–S51 (2000). 20. Kanda, T. & Hikasa, Y. Effects of medetomidine and midazolam alone or in combination on the metabolic and neurohormonal responses in healthy cats. Can. J. Vet. Res. 72, 332–339 (2008). 21. Zuurbier, C.J., Keijzers, P.J., Koeman, A., Van Wezel, H.B. & Hollmann, M.W. Anesthesia’s effects on plasma glucose and
insulin and cardiac hexokinase at similar hemodynamics and without major surgical stress in fed rats. Anesth. Analg. 106, 135–142 (2008). 22. Talukder, M.H. & Hikasa, Y. Diuretic effects of medetomidine compared with xylazine in healthy dogs. Can. J. Vet. Res. 73, 224–236 (2009). 23. Saleh, N. et al. Renal effects of medetomidine in isofluraneanesthetized dogs with special reference to its diuretic action. J. Vet. Med. Sci. 67, 461–465 (2005). 24. Gellai, M. Modulation of vasopressin antidiuretic action by renal alpha 2-adrenoceptors. Am. J. Physiol. 259, F1–F8 (1990). 25. Avsaroglu, H. et al. Strain differences in response to propofol, ketamine and medetomidine in rabbits. Vet. Rec. 152, 300 (2003). 26. Collins, T.B. Jr. & Lott, D.F. Stock and sex specificity in the response of rats to pentobarbital sodium. Lab. Anim. Care 18, 192–194 (1968). 27. Sinclair, M.D. A review of the physiological effects of α2-agonists related to the clinical use of medetomidine in small animal practice. Can. Vet. J. 44, 885–897 (2003). 28. Amouzadeh, H.R., Sangiah, S., Qualls, C.W. Jr., Cowell, R.L. & Mauromoustakos, A. Xylazine-induced pulmonary edema in rats. Toxicol. Appl. Pharmacol. 108, 417–427 (1991). 29. Amouzadeh, H. et al. Biochemical and morphological alterations in xylazine-induced pulmonary edema. Toxicol. Pathol. 21, 562–571 (1993).
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Mortality associated with using medetomidine and ketamine for general anesthesia in pregnant and nonpregnant Wistar rats
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Lauren M. Callahan, BSc, BVMS1, Simone M. Ross, AdvCert Animal Tech, Cert IV Training and Assessment1, Megan L. Jones, BSc2 & Gabrielle C. Musk, BSc, BVMS, PhD, Dipl ECVAA1,3
Medetomidine and ketamine are injectable drugs that can be used in combination to induce general anesthesia in rats. After noticing a high incidence of morbidity and mortality in pregnant Wistar rats given medetomidine and ketamine for anesthesia, the authors further investigated the effects of this combination of anesthetic drugs in both pregnant and nonpregnant Wistar rats. The time to recumbency and the duration of general anesthesia were similar between pregnant and nonpregnant rats. Pregnancy status did not affect the rats’ pulse rate, respiratory rate, rectal temperature, oxygen saturation or perfusion index during 2 h of anesthesia. Pregnant rats had significantly lower blood glucose concentrations than nonpregnant rats at all time points, though blood glucose concentrations increased in both groups. The mortality rate was ~15% both for nonpregnant rats and for pregnant rats. Researchers using medetomidine and ketamine to anesthetize Wistar rats should carefully monitor the rats in order to minimize mortality. General anesthesia is commonly induced in rats undergoing surgical and non-surgical procedures for research purposes. Either injectable drugs or inhalational drugs may be used to induce general anesthesia, but there are several advantages of using injectable anesthetics rather than inhalational drugs: the administration of injectable drugs is relatively simple, does not require an anesthetic machine, results in a comparatively smooth induction of anesthesia, avoids the risk of waste anesthetic gas leakage, is usually more cost-effective than use of inhalants and has reversible effects in some cases1,2. Furthermore, certain inhalational anesthetics may interfere with primary physiological endpoints. For example, because preconditioning with inhalational anesthetics such as isoflurane can confer protection from ischemiareperfusion–induced tissue damage in heart3, kidney4 and brain5, inhalational anesthetics may confound ischemia-reperfusion injury studies.
Medetomidine and ketamine are injectable drugs that can be used in combination to induce general anesthesia in rats6,7. Medetomidine is an α2adrenoreceptor agonist that produces sedation and analgesia, and ketamine is a dissociative anaesthetic1. This combination of drugs can induce surgical anesthesia for 30–60 min and unconsciousness for 60–120 min in rats1. The specific α2-adrenoreceptor antagonist atipamezole can be used to reverse the effects of medetomidine, inducing recovery from anesthesia within a few minutes of its administration. Reported mortality rates associated with medetomidine and ketamine anesthesia of Wistar rats vary. Roughan et al.8 reported 56% mortality in rats that were premedicated with buprenorphine and then administered various doses of medetomidine and ketamine in combination. In contrast, Hedenqvist et al.9 and Moriondo et al.10 reported no mortality associated with the drug combination, despite using doses
1Animal Care Services, University of Western Australia, Crawley, Australia. 2School of Anatomy, Physiology and Human Biology, University of Western Australia, Crawley, Australia. 3School of Veterinary and Life Sciences, Murdoch University, Murdoch, Australia. Correspondence should be addressed to G.C.M. ([email protected]).
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as high as 0.5 mg per kg body weight medetomidine and 75 mg per kg body weight ketamine. Although the use of medetomidine and ketamine in nonpregnant rats is widespread, there is little evidence to demonstrate their safety and efficacy in pregnant rats. A study done at the University of Western Australia (M.L.J., unpublished observations) used a combination of medetomidine and ketamine in pregnant Wistar rats undergoing laparotomy and clamping of isolated uterine blood vessels to investigate the effect of placental ischemia-reperfusion on pregnancy outcome. When the rats were administered 0.1–0.5 mg per kg body weight medetomidine and 50–60 mg per kg body weight ketamine by the intraperitoneal (i.p.) route at gestational day 17, anesthetic and recovery times were substantially longer than expected, and four of six rats (67%) either died during the procedure or were euthanized owing to post-anesthetic complications. Despite initial investigations into potential contributing factors (such as inadequate monitoring during and after anesthesia, environmental conditions and poor drug administration technique), the cause of the high morbidity and mortality could not be determined. In light of these preliminary findings, we hypo thesized that pregnant Wistar rats administered medetomidine and ketamine might have higher morbidity and mortality than nonpregnant Wistar rats. We administered a single dose of medetomidine and ketamine by the i.p. route to both nonpregnant and pregnant Wistar rats, measured the duration of anesthesia and monitored several parameters to determine the effect of pregnancy status on the efficacy and safety of this anesthetic combination. METHODS Animals All procedures involving animals were approved by the Animal Ethics Committee of The University of Western Australia in accordance with the guidelines of the Australian Code for the Care and Use of Animals for Scientific Purposes11. We obtained 34 outbred, 8-week-old Wistar (Crl:WI) rats (27 nulliparous females and seven males) from the Animal Resources Centre (Murdoch, Australia). The rats were housed under specific pathogen–free conditions. Rooms were maintained at 22 ± 2 °C and 30–70% humidity on a 12-h: 12-h light:dark cycle. We housed the rats in filter top cages (Expanded Rat Cage, Thoren Caging Systems, Hazleton, PA) lined with aspen bedding (Tapvei, Harjumaa, Estonia). Females were housed in pairs, and males were housed individually. Rats were fed a pellet diet (AIN93G Rodent diet, Specialty Feeds, Glen Forest, Australia) and given ad libitum access to acidified water (pH 2.5–3.0). We tested all rats for specific pathogens in accordance with the recommendations LAB ANIMAL
of ComPath (South Australian Health and Medical Research Institute, Adelaide, Australia). Vendor reports indicated that rats were seronegative for pneumonia virus of mice, Theiler’s encephalomyelitis virus, Sendai virus, mouse adenovirus types 1 and 2, sialodacryoadenitis, rat parvovirus, lymphocytic choriomeningitis, Hantaan virus and reovirus-3 and that rats were free of bacterial and parasitic infections at the time of shipment. All rats were acclimatized to the animal facility for 1 week prior to our studies. The female rats were divided into two groups that were time-mated 6 weeks apart. We synchronized the estrus cycles of the female rats in each group by administering an i.p. injection of 5.7 IU serum gonadotropin (Folligon, Intervet, Bendigo East, Australia) 4 d prior to mating and an i.p. injection of 1.93 IU chorionic gonadotropin (Chorulon, Intervet, Bendigo East, Australia) 2 d prior to mating. On the mating day, we placed two females in a male’s cage overnight. We removed the females from the male’s cage the following morning; this was designated gestational day 0. Pregnancy was confirmed by abdominal palpation on gestation day 14. Female rats were allocated to one of two groups according to their pregnancy status: nonpregnant (n = 13) and pregnant (n = 14). Personnel were not blinded to the pregnancy status of the rats. Anesthesia and monitoring We anesthetized nonpregnant rats and pregnant rats on gestational day 17. All procedures were conducted between 8.00 a.m. and 4.00 p.m. We transported all rats in their home cages to the surgical theater. Temperatures in the surgical theater were maintained at 22 ± 2 °C. We weighed the rats using an electronic balance with an accuracy of 1 g (PGW Precision Balance, Adam Equipment, Milton Keynes, UK). Pregnant rats weighed 349 ± 18 g on average, and nonpregnant rats weighed 273 ± 25 g on average. We administered a single i.p. injection of 0.2 mg per kg body weight medetomidine (1 mg/ml; Domitor, Zoetis, West Ride, Australia) mixed with 50 mg per kg body weight ketamine (100 mg/ml diluted to 0.2 mg/ml in 0.9% sterile saline; Parnell Laboratories, Alexandria, Australia) by inserting a 0.5-in, 26-gauge needle (Precision Glide, BD, Singapore) to a depth of 5 mm at an angle of 30–40° into the caudal right abdominal region. The same investigator (L.M.C.) administered each injection. Drug solutions were prepared each morning, and unused portions were discarded at the end of the day. Following the injection, each rat was placed into an individual cage until it lost its righting reflex. We then transferred each rat to a pre-warmed heating pad (Redflex Pet Recovery Pad; DLC Australia, Hoppers Crossing, Australia) to maintain body temperature between 37.0 °C and 38.0 °C, and we Volume 43, No. 6 | JUNE 2014 2 09
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FIGURE 1 | Mean pulse rate (beats per min) during anesthesia. Open symbols, nonpregnant rats; filled symbols, pregnant rats. Error bars, s.d.
applied ophthalmic ointment (Lacrilube; Allergan, Jersey City, NJ) to the eyes to prevent corneal desiccation. We did not provide oxygen supplementation; the rats breathed room air. We recorded the time to recumbency, defined as the time from the injection of the anesthetic to the loss of the righting reflex, for each rat. We considered the rat to be anesthetized when it failed to exhibit a withdrawal reflex in response to hind limb and forelimb toe pinches, a routine indicator of surgical anesthesia. We obtained a 0.05-ml blood sample from the lateral tail vein12 before anesthetics were injected and 20 min, 30 min, 60 min, 90 min and 120 min after anesthetics were injected. We measured blood glucose concentration of each sample immediately after the sample was obtained, using a glucometer (ACCU-Chek Performa, Roche Diagnostics, Castle Hill, Australia). We recorded pulse rate, respiratory rate, rectal temperature, oxygen saturation (SpO2) and perfusion index (PI) every 10 min. We measured the pulse rate by positioning a Doppler probe (Doppler Flow Detector Model 811-B, Parks Medical Electronics, Aloha, OR) over the coccygeal artery and counting the audible pulses over a period of 15 s. We determined the respiratory rate by counting chest excursions over a period of 15 s. We measured rectal temperature using an electronic thermometer by placing the probe at least 1 cm into the rectum; the same thermometer was used for all temperature measurements. We monitored SpO2 and PI using a pulse oximeter (Masimo Radical-7, Masimo Corporation, Irvine, CA) by positioning the probe over the skin flap behind the elbow or at the tail base. Recovery Two hours after anesthetics were administered to the rats, we administered 0.1 mg per kg body weight atipamezole (5 mg/ml diluted to 0.25 mg/ml in 0.9% 210 Volume 43, No. 6 | JUNE 2014
sterile saline; Antisedan; Zoetis, West Ride, Australia) by i.p. injection. We determined the return of the swallowing reflex by challenging the rat with a drop of oral glucose syrup (Queen Fine Foods, Alderley, Australia). We determined that walking ability had returned when the rat was able to maintain itself in sternal recumbency and move around the cage. For each rat, we recorded the duration of surgical anesthesia, defined as the time from the loss of the withdrawal reflex to the return of the withdrawal reflex. Anesthetic mortality was defined as any death that occurred within 24 h of anesthesia. We carried out in-house postmortem gross anatomical examinations of the rats that died. We euthanized pregnant rats 24 h after anesthesia. We made nonpregnant rats available to other researchers, if appropriate. Statistical analysis We assessed the data for normality and made statistical comparisons using SigmaPlot 12.0 software (Systat Software Inc., San Jose, CA). We used unpaired t-tests to determine differences between pregnant and nonpregnant rats. We compared non-parametric data using a Wilcoxon Rank Sum test. We carried out a regression analysis of the relationship between SpO2 and PI values using Sigmaplot 12.0 software. Data are expressed as mean ± s.d. unless otherwise stated. P values < 0.05 were considered statistically significant. RESULTS Efficacy and safety The mean time to recumbency did not differ significantly between nonpregnant (4.68 ± 2.39 min) and pregnant rats (5.68 ± 4.60 min). There were no significant differences between the two groups in mean pulse rates (Fig. 1), mean respiratory rates (Fig. 2), mean rectal temperatures (Fig. 3) or mean SpO2 (Fig. 4) at any time point during the 2-h anesthesia period. 100
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The correlations between PI and SpO2 were similar for the two groups (Fig. 5). The mean blood glucose concentration was significantly higher in nonpregnant rats than in pregnant rats at all time points (Fig. 6). The mean blood glucose concentration increased from the baseline in both groups, peaking at 90 min. Although exact urine output was not measured, we did observe increased urination in both nonpregnant rats and pregnant rats during anesthesia. Recovery and mortality One pregnant rat and one nonpregnant rat died while under anesthesia. The pregnant rat experienced respiratory arrest after 112 min of anesthesia and was euthanized. This rat had an elevated rectal temperature (41.2 °C) for a period of 5 min while under anesthesia. An in-house post mortem examination of this rat had no relevant findings. The nonpregnant rat experienced dyspnea after 100 min of anesthesia and was euthanized. An in-house post mortem examination of this rat had no relevant findings. All of the other rats (12 nonpregnant and 13 pregnant)
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remained anaesthetized for 120 min until atipamezole was administered. All but one of the rats (11 nonpregnant and 13 pregnant) recovered from anesthesia within a few minutes of receiving atipamezole. One nonpregnant rat did not respond to atipamezole after 40 min and was euthanized. An independent necropsy of this rat, carried out by a pathologist at Murdoch University Veterinary Hospital (Australia), detected pulmonary hyperemia with alveolar hemoglobin crystals, but no other lesions were noted. One pregnant rat was found dead 22 h after anesthesia. This rat had an elevated rectal temperature (40.8 °C) for a period of 5 min while under anesthesia. An in-house post mortem examination of this rat had no relevant findings. The recovery of the remaining rats was uneventful. We immediately made food and water available to the rats, and we observed normal feeding, drinking and behavior. 25
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The overall mortality rate was 14.8%, with a mortality rate of 15.4% among nonpregnant rats and a mortality rate of 14.3% among pregnant rats. DISCUSSION A previous unpublished study done at our institution showed longer than expected anesthetic and recovery times and 67% mortality when medetomidine and ketamine were administered to pregnant Wistar rats for general anesthesia. These findings prompted us to investigate how the effects of medetomidine and ketamine anesthesia differed between pregnant and nonpregnant Wistar rats. Overall, pregnancy status did not affect the animal’s anesthetic depth in response to medetomidine and ketamine, as measured by pulse rate and respiratory rate. The pulse rates observed in this study were predominantly below 300 beats per min, consistent with the known pharmacological ability of medetomidine to decrease the heart rate12. As expected, the respiratory rates of the anesthetized rats were also slightly depressed compared with the normal respiratory rate of 70–115 breaths per min in the rat. Monitoring the depth of anesthesia in rats can be challenging, owing to their small size and high metabolic rate. The normal pulse rate of a rat is 250–450 beats per min13, making it difficult to manually count. Although the Doppler probe that we used was designed for use in animals, it may not be sensitive enough for reliable use in small rodents such as rats. As such, we observed some variability in the pulse rate that may be attributed to human error. The rectal temperature of the rats was also unaffected by pregnancy status. Maintaining the target body temperature (37.0–38.0 °C) during anesthesia in both pregnant and nonpregnant rats was difficult in this study, and rectal temperatures changed rapidly. It is vital during anesthesia that the target body temperature is maintained in order to avoid adverse physiological consequences of hypothermia and hyperthermia. Hypothermia can prolong recovery from anesthesia and can cause myocardial infarctions and coagulopathies, whereas hyperthermia can cause tachycardia, hyperventilation and cardiac arrhythmias14. Rats are especially prone to hypothermia during anesthesia because of their high metabolic rate and large surface-to-body-weight ratio, but hyperthermia is also a risk if an active heating device is used1. Given the temperature fluctuations we observed with this anesthetic regime, we suggest particular attention be paid to monitoring and maintaining body temperature when using medetomidine and ketamine for anesthesia, especially if active heating devices are being used. We did not find any differences in SpO2 or the relationship between SpO2 and PI between pregnant and nonpregnant rats. However, we were unable to 212 Volume 43, No. 6 | JUNE 2014
obtain consistent readings using the pulse oximeter. Despite observing SpO2 values below 90%, compared to normal SpO2 values of >95%, there was no obvious clinical evidence of hypoxemia in these rats. A limitation of our study was that we were unable to carry out arterial blood gas analyses; these would have allowed us to interpret SpO2 in the context of the partial pressure of oxygen in arterial blood. The majority of the SpO2 readings were associated with low PI, which may be due to the peripheral vasoconstrictive effects of medetomidine12. Pregnant rats had significantly lower blood glucose concentrations throughout the anesthetic period than nonpregnant rats, consistent with previous observations in rats15. An increased predisposition to hypoglycemia in mid- to late pregnancy has been demonstrated in other species as well16–18 and is attributed to maternal diversion of glucose to the developing fetus to meet demand during the period of maximal fetal growth19. Both nonpregnant and pregnant rats showed a steady increase in blood glucose concentrations in response to the administration of medetomidine and ketamine, likely owing to the hyperglycemic effects of medetomidine20,21. We suggest it may be prudent for researchers to avoid using medetomidine in studies in which persistent hyper glycemia interferes with their aims. We also observed in both groups of rats that urine output was qualitatively increased during anesthesia. This is a commonly reported side effect of α2adrenoreceptor agonists22–24. We recommend that perioperative fluid therapy be considered when anesthetizing animals with medetomidine, especially if intra-operative hemorrhage is likely. All rats that survived the anesthetic period remained anaesthetized for 2 h and required administration of atipamezole for arousal. This is not consistent with published anesthetic times6–10. Studies have shown that the strain and sex of an animal may affect its response to different injectable anesthetic agents25,26. Responses to equipotent doses of medetomidine vary widely between species owing to differences in α2-adrenoreceptor subtypes, densities and locations within the central nervous system and peripheral tissue27. Sex may also impact the response to medetomidine, because higher estrogen levels can increase the sensitivity of α2-adrenoreceptors27. Therefore, it is recommended when using medetomidine and ketamine that the doses be optimized to determine the safest and most efficacious combination for the particular strain. In addition, atipamezole should always be available to induce arousal. Mortality rate was not associated with pregnancy status, although both of the pregnant rats that died showed a temporary increase in body temperature that may have contributed to their death. The significance www.labanimal.com
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of this complication in this study is unknown. A limitation of this study is that comprehensive necropsy with histopathology was not done on all rats that died. The pulmonary hyperemia and edema detected upon necropsy examination in one nonpregnant rat may have compromised alveolar gas exchange. Given that xylazine may cause pulmonary edema in rats, albeit at high doses, it may be prudent to supplement oxygen in rats receiving α2-adrenoreceptor agonists28,29. Although oxygen supplementation during anesthesia is good practice, we aimed to use only equipment readily available in the laboratory animal research setting, whereas the use of oxygen requires delivery through an anesthetic machine and breathing system. In light of our findings, we urge researchers to provide supplementary oxygen to anaesthetized rats by a facemask if this combination of drugs is to be used for anesthesia. A possible cause of death in the rats that died during the anesthetic period is upper respiratory tract obstruction. Rats recovering from anesthesia must be continuously monitored until their swallowing reflex has returned, because the lack of this reflex poses a risk of choking in the case of any upper respiratory tract obstructions. Due to their small size, it can be difficult to determine when this reflex has returned. We have found that application of glucose syrup to the mouth and tongue after the rats show signs of regaining consciousness (such as head movements) is an effective technique for assessing the swallowing reflex. Other possible causes of anesthetic-related death in our study include cardiac dysrhythmias, cardiac arrest and hypoxia secondary to pulmonary edema or pulmonary vasoconstriction. It would have been useful to measure end tidal carbon dioxide concentrations, electrocardiogram and blood pressure. In future studies, specialized physiological monitoring equipment suitable for small rodents should be used to clarify the nature of anesthetic-related complications. As fetal viability was outside the scope of this study, and the rats were euthanized the following day, fetal viability was not assessed. Therefore, it is not possible to determine whether medetomidine and ketamine anesthesia affected placental perfusion. In principle, drugs such as medetomidine that stimulate the α2adrenoreceptor could increase the contractibility of the uterus in pregnant rats, leading to fetal resorption. The increase in progesterone levels that occur during pregnancy could counteract this effect by stimulating the sensitivity of β-adrenoreceptors, however, which would decrease the contractibility of the uterus27. Medetomidine and ketamine can be used to effectively induce general anesthesia in nonpregnant and pregnant Wistar rats, but this combination of
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a nesthetics resulted in a mortality rate of ~15% regardless of pregnancy status. Anesthetic monitoring equipment suitable for use in small rodents should be used in future studies to help detect physiological complications that may be managed or prevented to reduce mortality associated with these anesthetics. Furthermore, researchers should consider alternative approaches to anesthesia in Wistar rats. If that is not feasible, then we suggest researchers undertake an initial dose optimization study of these drugs for a specific strain of Wistar rat prior to commencing the study. Acknowledgments We thank Leah Attwood (Animal Services, The University of Western Australia) for providing technical support. COMPETING FINANCIAL INTERESTS The authors declare no competing financial interests. Received 22 January 2014; accepted 7 March 2014 Published online at http://www.labanimal.com/ 1. Flecknell, P. Laboratory Animal Anaesthesia 3rd edn. (Academic, Waltham, MA, 2009). 2. Hacker, S.O., White, C.E. & Black, I.H. A comparison of target-controlled infusion versus volatile inhalant anaesthesia for heart rate, respiratory rate, and recovery time in a rat model. Contemp. Top. Lab. Anim. Sci. 44, 7–12 (2005). 3. Stowe, D.F. & Kevin, L.G. Cardiac preconditioning by volatile anesthetic agents: a defining role for altered mitochondrial bioenergetics. Antioxid. Redox. Signal. 6, 439–448 (2004). 4. Hashiguchi, H. et al. Isoflurane protects renal function against ischemia and reperfusion through inhibition of protein kinases, JNK and ERK. Anesth. Analg. 101, 1584–1589 (2005). 5. Steiger, H.J. & Hänggi, D. Ischaemic preconditioning of the brain, mechanisms and applications. Acta Neurochir. (Wien) 149, 1–10 (2007). 6. Nevalainen, T., Pyhälä, L., Voipio, H.M. & Virtanen, R. Evaluation of anaesthetic potency of medetomidine-ketamine combination in rats, guinea-pigs and rabbits. Acta Vet. Scand. Suppl. 85, 139–143 (1989). 7. Jang, H.S., Choi, H.S., Lee, S.H., Jang, K.H. & Lee, M. Evaluation of the anaesthetic effects of medetomidine and ketamine in rats and their reversal with atipamezole. Vet. Anaesth. Analg. 36, 319–327 (2009). 8. Roughan, J.V., Ojeda, O.B. & Flecknell, P.A. The influence of pre-anaesthetic administration of buprenorphine on the anaesthetic effects of ketamine/medetomidine and pentobarbitone in rats and the consequences of repeated anaesthesia. Lab. Anim. 33, 234–242 (1999). 9. Hedenqvist, P., Roughan, J.V. & Flecknell, P.A. Effects of repeated anaesthesia with ketamine/medetomidine and of preanaesthetic administration of buprenorphine in rats. Lab. Anim. 34, 207–211 (2000). 10. Moriondo, A. et al. Impact of respiratory pattern on lung mechanics and interstitial proteoglycans in spontaneously breathing anaesthetized healthy rats. Acta Physiol. (Oxf.) 203, 331–341 (2011). 11. National Health and Medical Research Council, Australian Government. Australian Code of Practice for the Care and Use of Animals for Scientific Purposes 4th edn. (Australian Government, Canberra, Australia, 2004). 12. Plumb, D.C. Plumb’s Veterinary Drug Handbook 5th edn. (Blackwell, Ames, IA, 2005).
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13. Hrapkiewicz, K. & Medina, L. (eds.) Clinical Laboratory Animal Medicine: An Introduction 3rd edn. (Blackwell, Oxford, UK, 2007). 14. Dugdale, A. Veterinary Anaesthesia: Principles to Practice Ch.18 (Wiley Blackwell, West Sussex, UK, 2010). 15. Wharfe, M.D., Mark, P.J. & Waddell, B.J. Circadian variation in placental and hepatic clock genes in rat pregnancy. Endocrinology 152, 3552–3560 (2011). 16. Johnson, C.A. Glucose homeostasis during canine pregnancy: insulin resistance, ketosis, and hypoglycemia. Theriogenology 70, 1418–1423 (2008). 17. Reddi, A.S., Oppermann, W., Strugatz, L.H., Cole, H.S. & Camerini-Davalos, R.A. Effect of pregnancy on serum alanine concentration in normal and genetically diabetic mice. Horm. Metab. Res. 8, 478–482 (1976). 18. Schlumbohm, C. & Harmeyer, J. Twin-pregnancy increases susceptibility of ewes to hypoglycaemic stress and pregnancy toxaemia. Res. Vet. Sci. 84, 286–299 (2008). 19. Herrera, E. Metabolic adaptations in pregnancy and their implications for the availability of substrates to the fetus. Eur. J. Clin. Nutr. 54, S47–S51 (2000). 20. Kanda, T. & Hikasa, Y. Effects of medetomidine and midazolam alone or in combination on the metabolic and neurohormonal responses in healthy cats. Can. J. Vet. Res. 72, 332–339 (2008). 21. Zuurbier, C.J., Keijzers, P.J., Koeman, A., Van Wezel, H.B. & Hollmann, M.W. Anesthesia’s effects on plasma glucose and
insulin and cardiac hexokinase at similar hemodynamics and without major surgical stress in fed rats. Anesth. Analg. 106, 135–142 (2008). 22. Talukder, M.H. & Hikasa, Y. Diuretic effects of medetomidine compared with xylazine in healthy dogs. Can. J. Vet. Res. 73, 224–236 (2009). 23. Saleh, N. et al. Renal effects of medetomidine in isofluraneanesthetized dogs with special reference to its diuretic action. J. Vet. Med. Sci. 67, 461–465 (2005). 24. Gellai, M. Modulation of vasopressin antidiuretic action by renal alpha 2-adrenoceptors. Am. J. Physiol. 259, F1–F8 (1990). 25. Avsaroglu, H. et al. Strain differences in response to propofol, ketamine and medetomidine in rabbits. Vet. Rec. 152, 300 (2003). 26. Collins, T.B. Jr. & Lott, D.F. Stock and sex specificity in the response of rats to pentobarbital sodium. Lab. Anim. Care 18, 192–194 (1968). 27. Sinclair, M.D. A review of the physiological effects of α2-agonists related to the clinical use of medetomidine in small animal practice. Can. Vet. J. 44, 885–897 (2003). 28. Amouzadeh, H.R., Sangiah, S., Qualls, C.W. Jr., Cowell, R.L. & Mauromoustakos, A. Xylazine-induced pulmonary edema in rats. Toxicol. Appl. Pharmacol. 108, 417–427 (1991). 29. Amouzadeh, H. et al. Biochemical and morphological alterations in xylazine-induced pulmonary edema. Toxicol. Pathol. 21, 562–571 (1993).
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