Efficacy of Warmed Inspired Air for Prevention of Perianesthetic Hypothermia in New Zealand White Rabbits (Oryctolagus cuniculus)
Warmed inspired air circuits have proved to be effective in semiclosed heating modalities in veterinary species such as dogs, cats, and nonhuman primates, and thus, there is a gap in species requiring a nonrebreathing circuit. This study evaluated the efficacy of using warmed inspired air in addition to conductive mattress warming in the prevention of hypothermia in anesthetized rabbits (Oryctolagus cuniculus). Rabbits were divided into 2 groups: conductive warming only (control) or conductive warming and warmed inspired air. Our results showed that the addition of a warmed air anesthesia circuit had a significant positive effect on perianesthetic body temperature, maintaining a higher rectal temperature starting 10 minutes after induction and a higher final rectal temperature after a 45-minute anesthetic procedure. At 20 minutes after induction, the body temperature of the warmed air group was not significantly different from baseline compared with a significant drop from baseline in the control group. Infrared pinnal temperatures did not show a pairwise significance in the effect of heating modality and time; however, a clinically significant difference of 2-3 °F between groups was seen. There were no statistically significant differences between groups for time to full recovery and time to extubation. For procedures using rabbits, the addition of warmed inspired air should be considered a significant refinement that promotes normothermia during anesthesia based on consistent and improved overall body temperatures.
Introduction
The effort to combat hypothermia-associated sequelae during veterinary anesthesia such as altered hemostasis, increased risk of wound infection, and prolonged recovery time focuses on addressing the different mechanisms of heat loss.1,2 Body heat is lost under anesthesia through 4 primary mechanisms: radiation (infrared heat loss to the environment), convection (heat transfer to moving air), conduction (direct heat transfer to surfaces in contact with the patient), and evaporation (loss of moisture from the respiratory tract, skin, or open surgical sites).3 Additional factors, such as fur clipping, the use of isopropyl alcohol or other antiseptics during surgical site preparation, and patient exposure to ambient temperature during preoperative procedures and transport, can further contribute to heat loss. To offset this, warming methods such as the use of conductive heating pads, forced-air warming blankets, and heated surgical tables are traditionally employed in veterinary medicine. However, these modalities only address radiative and conductive heat losses. The standard of care in human medicine is to use warmed and humidified air to prevent evaporative heat loss, but there is minimal objective data available regarding the effectiveness in species such as the rabbit.
Rabbits are commonly used in biomedical research worldwide. Refinement of rabbit surgical and anesthetic techniques improves clinical outcomes and data quality, while standardizing these refinements in studies is key to ensuring reproducibility. We sought to investigate the utility of warming the inspired air during anesthesia of New Zealand White rabbits to address the evaporative mechanism of heat loss from the respiratory tract during anesthesia. There is currently no other literature using an in-line warmer on an intubated rabbit.4
Available equipment to test our hypothesis included an inexpensive, low-maintenance heated breathing circuit designed for use in veterinary medicine and easily integrated into any anesthesia machine as a free-standing unit (Darvall Heated Breathing Circuit; Advanced Anesthesia Specialists, Gladesville, NSW, Australia). The breathing circuit warms inspired air to 104 °F using a feedback loop to prevent overheating and is easily cleaned and disinfected between patients. The feedback loop increases or decreases the temperature, as needed (within the range of the device 68-122 °F) to maintain a temperature range of normothermia in a rabbit (100-104 °F) based on the physiologic conditions of the patient for the duration of anesthesia.
The efficacy of this heated breathing circuit was previously explored in intubated rhesus macaques (Macaca mulatta) resulting in an appreciably faster return to baseline temperature and a higher final temperature after a 2-hour anesthetic procedure.5 Another study incorporated the Darvall circuit with anesthesia of callimicos, smaller nonhuman primates weighing 544 ± 97 g, but found that the rate of heat loss did not differ between methods using a reflective blanket alone or with a heated anesthetic circuit via face mask. The authors attributed this to the small size of the animal, the high body surface area-to-mass ratio, and the relatively minimal amount of heat lost via the respiratory tract compared with radiative and conductive heat losses.6 As seen in dogs, cats, humans, and nonhuman primates, a heated anesthesia breathing circuit may have a more substantial effect on maintaining normothermia in animals >1 kg, suggesting that it may have value for anesthesia in rabbits.3,5,7–10 Thus, use of a heated breathing circuit as an additional warming method in rabbits may reduce the risk of inhalation anesthesia hypothermia-associated adverse effects and serve as a valuable refinement.
Implementation of an additional warming method has been shown to be effective at preventing perioperative hypothermia and reducing the risk of hypothermia-associated adverse effects. Conductive warming is as effective as other warming methods in dogs3 and is inexpensive and available at many research institutions; and it is likely that a heated breathing circuit could have synergistic benefits in providing normothermia. This would address evaporative heat loss from the respiratory tract and radiative and conductive heat loss using the conductive warming mattress.
We hypothesized that the use of a heated anesthesia breathing circuit to provide warmed anesthetic gas to rabbits when used in conjunction with conductive warming would result in reduced heat loss and a more stable body temperature compared with anesthetized rabbits that received solely conductive warming support.
Ethical Review
The Walter Reed Army Institute of Research (WRAIR) has reviewed the material described here. There is no objection to its presentation and/or publication. The opinions or assertions contained herein are the private views of the authors and are not to be construed as official, or as reflecting true views of the Department of the Army or the Department of Defense. Research was conducted in accordance with an IACUC-approved animal use protocol in a facility accredited by AAALAC International, with a Public Health Services Animal Welfare Assurance, and in compliance with the Animal Welfare Act and other federal statutes and regulations relating to laboratory animals. The study adhered to the principles outlined in the Guide for the Care and Use of Laboratory Animals (National Research Council, 2011 edition).
Materials and Methods
Study design.
Anesthetic events 1 and 2.
Each rabbit was anesthetized, fitted with a supraglottic airway device, clipped, and aseptically prepped as they would for a surgical procedure (Figure 1). Animals served as their own controls with each animal undergoing 2 anesthetic events of approximately 45 minutes. The duration used was to mimic the length of a routine surgical procedure performed on a rabbit.11 Rabbits were randomized and evenly distributed for which group they were placed in for the first anesthetic event. Rabbits had 6 days of recovery between anesthetic events, with the second anesthetic event procedure for each rabbit conducted at the same time of day as the first. The room temperature and humidity of the procedure room were approximately 66 °F and 30% humidity for all procedures.
Citation: Journal of the American Association for Laboratory Animal Science 2025; 10.30802/AALAS-JAALAS-25-118
Groups were as follows:
Control group: the heated breathing circuit was connected to the anesthesia unit, but not turned on, while the conductive heating mattress provided heat support with the temperature set at 104 °F.
Warmed air group: the heated breathing circuit and conductive heating mattress (temperature set at 104 °F) were both turned on. The heated breathing circuit was set to the standard setting of 104 °F.
Sample size.
To approximate the comparison of interest in the primary outcomes, the pairwise comparisons resulting from the interaction term in the mixed model or ANOVA, a 2-sided paired samples t test was used to compute the appropriate sample size a priori. It was anticipated that the effect size, using Cohen’s dz for paired samples, of the difference between treatments (control/warmed air) would be dz = 1. The study would be powered at 80% to find this difference using 10 animals that would contribute data twice, since each animal served as its own control. This difference was assumed to be constant for all between-group comparisons in both the outcomes of rectal and pinnal temperatures over time.5 Sample size computation was conducted in G*Power version 3.1.9.7 (https://www.psychologie.hhu.de/arbeitsgruppen/allgemeine-psychologie-und-arbeitspsychologie/gpower).
Inclusion and exclusion criteria.
Exclusion criteria during the experiment included if the core body temperature dropped below 94 °F or higher than 104 °F for more than 30 minutes without improving using multimodal current warming or cooling methods. Animals displaying signs of pain or distress including, but not limited to, vocalization and avoidance or aggressive behavior, hyperactivity and/or restless and nervous behavior, listless behavior and failure to rise when approached by personnel, self-mutilation or persistent rubbing or licking, altered gait or abnormal posture, and changes in skin texture or appearance that cannot be alleviated with veterinary consultation were criteria for excluding animals. However, using these criteria, no animals were excluded from this study. Criteria for inclusion or exclusion for data points were not set. However, vulva infrared temperature measurements were excluded from this study as they did not reflect an accurate measurement of core body temperature and were too variable between each animal and within each animal. The sample size for this crossover study used 10 animals.
Randomization.
Randomization was used to allocate animals to initial groups by block randomization using Sealed Envelope to avoid the potential confounder of the order of anesthetic events and measurements. All animals received both warmed air and no warmed air treatments 6 days apart. Animals were chosen based on the same sex, age, and similar weight.
Blinding.
No blinding was possible in the preparation, conduct, and data analysis of this experiment due to the nature of its design.
Outcome measures.
The primary outcome of change in temperature (°F), both rectal and pinnal, was evaluated. Secondary outcomes of time (minutes) to extubation and full recovery were explored.
Statistical methods.
GraphPad Prism version 10.4.1 for Windows (GraphPad Software; www.graphpad.com) was used to analyze the data. To assess whether the data met the assumptions of the statistical approach, measures taken included looking at the main effect of group (control/warmed air), main effect of time, and the interaction of the 2. Comparison of the respective time point back to the initial time point for the groups occurred before the pairwise comparison to analyze the significance of the interactions occurring in the experiment.
Experimental animals.
The study group was 10 purpose-bred female New Zealand White rabbits (Oryctolagus cuniculus) purchased from Charles River Canada (8 months old; weight [mean ± 1 SD]: 4.25 ± 0.25 kg). Study animals were selected to be approximately the same size and body condition. These animals were serologically negative for Treponema cuniculi, Encephalitozoon cuniculi, Filobacterium rodentium, and Pasteurella multocida, and negative for all fecal helminthes and external arthropod parasites. The rabbits were singly housed in cages with perforated-bottom caging (Lenderking Caging Products, Millersville, MD) and fed a commercial diet (Prolab Hi-Fiber Rabbit Diet; LabDiet, Richmond, IN). Food was provided once daily and water ad libitum via automatic lixit, being checked at least twice daily while assigned to the study. Enrichment was provided in the form of toys, dietary enrichment, and exercise, at a minimum of twice weekly for no less than 20 minutes. Timothy hay was provided once daily as well as papaya tablets daily with the pelleted food, excluding weekends. Animals were housed in ABSL1 rooms at 61-72 °F with a relative humidity of 30%-70%. Anesthesia events were conducted in a surgical suite with a temperature set point of 66 °F and 30% humidity.
Experimental procedures.
Anesthesia and warming.
Each rabbit was anesthetized via intramuscular injection with 30 mg/kg ketamine (Covetrus, Portland, ME) and 4 mg/kg xylazine (Covetrus, Portland, ME) for the 2 separate anesthetic events. The animals were removed from their home enclosures, just before administration of the anesthetic injection, which was done while the animals were manually restrained. They were returned to their home enclosures until reaching a level of sedation safe for transport to the procedure room. Each animal was fitted with a rabbit supraglottic airway device (SGAD; v-gel advanced; DocsInnovent, Hemel Hempstead, United Kingdom). The SGAD was chosen based on body mass as per the manufacturer’s recommendations. In cases in which an animal fell between sizes, the larger SGAD was selected. An insertion attempt began after the rabbit was preoxygenated and fully anesthetized with the jaw relaxed and reflexes absent after at least 5 minutes of isoflurane administered via face mask. A laryngoscope was used to insert a lubricated SGAD (Surgilube Non-Spermicidal Lubricating Topical Jelly; Fougera Pharmaceuticals, Melville, NY) into the oropharynx until either resistance was encountered or the incisors were within 1-2 cm of the device’s fixation tabs. Acceptable placement was confirmed by testing for a patent airway according to the manufacturer with auscultation of bilateral breath sounds during positive pressure ventilation and use of a pulse oximeter to monitor oxygen saturation above 95%. Next, the rabbit was connected to the nonrebreathing circuit. A small area (approximately 10 × 10 cm) of the ventral abdomen was shaved of fur followed by a sterile surgical scrub of 3 rounds of alcohol, mimicking the sterile prep that precedes an abdominal laparotomy. Isoflurane anesthesia (Baxter, Deerfield, IL) was administered using an isoflurane vaporizer (A.M. Bickford, Wales Center, NY) on a veterinary anesthesia machine (Surgivet, Waukesha, WI) at 1%-3% to maintain surgical anesthetic depth. The oxygen flow rate for each procedure was 0.5 L/min. The Darvall Heated Breathing Controller with LACK SWT Nonrebreathing Circuit Kit (Darvall nonrebreathing LACK circuit for use in patients 1-25 lbs; no. 8305; Advanced Anesthesia Specialists, Gladesville, NSW, Australia; Figure 2A) was used as the breathing circuit for all anesthetized patients. The LACK circuit kit can be used with the Darvall heat controller unit and the small 12 mm smooth wall heated tubing 2-30 kg, as a nonrebreathing heated circuit suitable for animals 0.5-7 kg. Each rabbit underwent two 45-minute anesthetic events, each beginning from the time of connection to the nonrebreathing circuit. For both procedures, the rabbit was placed in dorsal recumbency on a vinyl-covered foam v-trough with the head extending beyond the v-trough and monitoring equipment connected. A conductive heating mattress was placed in direct contact with the animal, in accordance with the manufacturer’s instructions, underneath the patient set to 104 °F with the temperature management controller heating unit (V104–Large 22” x 31” Warming Blanket with Multifunction Warming Controller WC52; HotDog; Augustine Surgical, Eden Prairie, MN). The heating mattress was positioned under each rabbit in the v-trough with the black side facing the rabbit to always ensure contact with the sensor according to the manufacturer’s instructions for use. The patient was covered with a disposable surgical drape, leaving only the head and a 10 × 10 cm fenestration over the abdomen exposed (HALYARD* Minor Procedure Drape; Halyard Health, Alpharetta, GA; Figure 2B).
Citation: Journal of the American Association for Laboratory Animal Science 2025; 10.30802/AALAS-JAALAS-25-118
Following the 45-minute anesthetic event, the anesthesia gas vaporizer was turned off, and the animal was given at least 5 minutes of oxygen. The rabbit was monitored closely during recovery and allowed to regain consciousness naturally following the cessation of isoflurane anesthesia. Extubation was performed based on clinical signs indicating readiness, including coughing, voluntary movement, and stable respiratory patterns. Each rabbit was returned to its home cage once fully awake, responsive, and able to move voluntarily upright with a normal body temperature (100-104 °F), a point defined as full recovery. The heated nonrebreathing circuit unit was sanitized according to the manufacturer’s recommendations between procedures.
Physiologic parameter monitoring.
During anesthesia, a rectal thermometer probe was inserted 3 cm into the rectum of each rabbit. The rectal probe was plugged into the Darvall Heated Breathing Circuit port to display both the thermometer probe temperature and temperature at the end of the breathing circuit of the anesthetic gas delivered to the patient. A manual rectal thermometer probe (MWI Animal Health, Boise, ID) was also inserted approximately 3 cm into the rectum. In addition, an infrared thermometer (Etekcity Lasergrip 630; Vesync, Anaheim, CA) was used to take the temperature of the pinna. The location used for taking the pinnal temperature was the bisection of the long axis (L), the length from base to the tip of the pinna, and the short axis (W), the width of the pinna perpendicular to the length measurement. The average reading was measured when the laser points overlapped or within 1-2 cm of each other on the pinna as demonstrated in Figure 2B. Other vital signs including heart rate, respiratory rate, and SpO2 were measured via a Vetcorder (Vetcorder; MAI Animal Health, Elmwood, WI) for the duration of the anesthetic event.
Body temperature was determined in degrees Fahrenheit every 5 minutes via the manual rectal thermometer probe and infrared thermometer probe and recorded. Baseline temperature was designated as the body temperature measured when the patient was initially connected to the breathing circuit and monitoring equipment (“time = 0”). Other vital signs were also measured every 5 minutes.
Data analysis.
Data was entered in an Excel spreadsheet and imported for analysis in GraphPad Prism version 10.4.1 for Windows. The primary outcomes of change in rectal and pinnal temperature were evaluated using a repeated 2-way ANOVA and a linear mixed model, respectively. Both models had the main effects of time and group, as well as the interaction of the 2. Because animals served as their own controls, both factors were considered repeated. Subsequent pairwise comparisons explored both within-groups and between-group effects. Each time point was compared back to the baseline value using a Dunnett test. Comparisons between groups used a Sidak multiple comparisons test at each time point.
Secondary outcomes of time (minutes) to extubation and to full recovery were explored using a series of paired t tests between the 2 groups. Normality was visually assessed and tested through the D’Agostino-Pearson Omnibus K2 test. Two out of the 3 secondary outcomes passed this assumption. The results were compared with the Wilcoxon signed rank test, though these results are not shown. For ease of understanding and consistency, the parametric test was run on all outcomes.
Results from the primary outcome models are reported using means, SEs, and 95% CIs. Secondary outcome results are summarized using means and SDs. P values are considered significant at the 5% level, and all tests are 2 sided.
Results
The mean difference (SE) between average baseline and final body temperatures for the warmed air group compared with control was 0.12 (0.13) and 0.97 (0.13) °F, respectively, for rectal temperatures (Table 1). The mean difference and SE between average baseline and final body temperatures for the warmed air group compared with control were 1.01 (0.79) and 1.24 (0.81) °F, respectively for pinnal temperatures (Table 1). There was statistical significance for the main effects of time and group, as well as the interaction of the 2, when comparing change in rectal temperature (P < 0.0001, P = 0.0037, P < 0.0001, respectively; Figure 3). For the control group, rectal temperature varied significantly starting 20 minutes after baseline (P = 0.0003 at minute 25, and P < 0.0001 for remaining time points). There was no statistical difference in temperature at any time point compared with baseline for the warmed air group (P > 0.05 for every time point). When comparing the warmed air with control groups, differences in rectal temperature were seen starting at 10 minutes with the warmed air group maintaining a higher temperature throughout (P = 0.0211 at minute 10, and P ≤ 0.001 for remaining time periods). The pinnal temperature in the rabbits did not display any statistical significance for the main effects of time and group, nor the interaction of the 2 (P = 0.1529, P = 0.1215, P = 0.5621, respectively; Figure 3). In addition, there was no statistical significance between treatment groups for time to extubation or time to recovery (P = 0.1647, P = 0.3919, respectively; Figure 4).
| Treatment group | Reading | Baseline (°F) | Final (°F) | Mean difference (SE) |
|---|---|---|---|---|
| Warmed air | Rectal | 102.5 | 102.4 | 0.12 (0.13) |
| Control | Rectal | 102.4 | 101.4 | 0.97 (0.13) |
| Warmed air | Pinna | 90.55 | 89.54 | 1.01 (0.79) |
| Control | Pinna | 88.32 | 87.08 | 1.24 (0.81) |
Note that the final reading was at 45 min.
Citation: Journal of the American Association for Laboratory Animal Science 2025; 10.30802/AALAS-JAALAS-25-118
Citation: Journal of the American Association for Laboratory Animal Science 2025; 10.30802/AALAS-JAALAS-25-118
Discussion
These results support our hypothesis that the use of a heated anesthesia breathing circuit to provide warmed anesthetic gas to anesthetized rabbits when used in conjunction with conductive warming results in reduced heat loss and a more stable body temperature compared with anesthetized rabbits that received solely conductive warming support. When compared with conductive mattress warming alone, the addition of a warmed air anesthesia circuit had a significant positive effect on perianesthetic body temperature with maintenance of a higher rectal temperature starting 10 minutes after induction and a higher final rectal temperature after a 45-minute anesthetic procedure. Normothermia was superior for the warmed air group, with no significant variation in temperature from baseline for the duration of the anesthetic event, whereas the control group experienced a significant decrease in temperature after 20 minutes of anesthesia. Maintaining normothermia is essential for ensuring the stability of the cardiovascular system, respiratory system, and drug metabolism during anesthesia and surgical procedures. Hypothermia can lead to significant physiologic disturbances, including bradycardia, impaired gas exchange, and altered drug metabolism.
Hypothermia results in a cascade of systemic physiologic changes that place significant stress on the body. Maintenance of normothermia preserves normal physiologic processes and minimizes stress. The benefits of reducing hypothermia are particularly significant for animals undergoing longer procedures or those in compromised health, where the risks of hypothermia are amplified. During prolonged anesthesia, animals are exposed to cumulative heat loss due to factors such as cold surgical environments, evaporative cooling from surgical scrubs, and the use of cold anesthetic gases. Compromised animals, such as those with preexisting conditions or those immunocompromised, are at higher risk of infection. Preventing hypothermia helps preserve immune function and reduces the likelihood of postoperative infections. Actively preventing hypothermia not only improves outcomes but also aligns with ethical responsibility in veterinary and research settings and thus improves animal welfare.
Our results align with those found in rhesus macaques using the same equipment, but differ from those in callimicos.5,6 A possible explanation for the differences observed in our study compared to those in callimicos could be accounted for by the use of a SGAD as compared with a facemask. A SGAD, such as a v-gel, creates a sealed airway that directs heated and humidified air directly into the trachea, allows for controlled ventilation, and minimizes heat loss into the surrounding environment. Heat loss can occur due to the open design of the facemask, especially in small animals such as callimicos with high surface area-to-body mass ratios.
There was no difference in time to extubation or to full recovery, a finding that may be attributed to our anesthetic cocktail of ketamine and xylazine, and unrelated to heating. In addition, this finding may have been due to the shorter anesthetic period, as it has been concluded that heated inspired air was beneficial in procedures greater than one hour for these variables.5 Despite this finding, maintenance of normothermia should be emphasized due to its direct correlation to improved patient outcomes and reduced complications when considering the addition of a heated anesthesia breathing circuit as a refinement to a multimodal plan for prevention of hypothermia.
To minimize confounding variables, this study did not perform any concurrent surgical procedures that would have introduced an open incision or blood loss as variables outside of evaporative heat loss from the respiratory tract. Room temperature and humidity, temperature recorders, anesthesia, procedure preparation, time from premedication to intubation, and other factors like draping and positioning were consistent between patients to allow the heated breathing circuit to be the only significant difference between anesthetic events.
It should be noted that the heated nonrebreathing circuit was a modified heated Bain for use in smaller animals. This study supports that, despite the high flow rate in the Bain circuit, patients appear to have received warmed air, due to Darvall’s smooth wall tubing and minimum dead space Y piece connector.
We were unable to use an esophageal probe in this study due to placement of v-gels, which seal off the esophagus. During anesthesia, thermoregulatory mechanisms are often impaired, and peripheral temperature may not accurately reflect the body’s thermal state, and thus, measurement of core body temperature at the heart level is the gold standard. However, the rectal temperature data we obtained support our hypothesis. The pinnal temperatures taken with the infrared thermometer did not show a statistically significant difference across groups (warmed air compared with control) and time. However, when looking at the data, there appeared to be a clinically significant 2-3 °F higher difference in the warmed air group compared with the control group (Figure 4B). These findings may be due to the rabbits having thermoregulated by peripheral vasoconstriction to shunt blood flow to the core more efficiently than other species because of the large size of their pinnae having prominent vasculature.12 It seems highly likely that more significant changes in the body temperature are needed to impact the ear temperature.
A limitation of this study is that the Darvall unit was used at the time of induction and did not evaluate its efficacy in restoring already hypothermic patients to normothermia. In addition, the Darvall unit does not humidify the warmed air, which may damage the mucociliary apparatus. Proper humidification and hydration remain critical strategies to maintain respiratory health and prevent complications associated with dry air exposure. Another unit uses humidified warm air delivery to the patient, but there is currently no literature to compare it to the Darvall and no nonrebreathing circuit option supplied by the manufacturer for use in smaller species (BreatheWarm Intro Kit; Veterinary Surgical Supply; www.vssinfo.com). Another consideration would be to evaluate temperature probe measurements at the end of the endotracheal tube in comparison to rectal temperatures.
The use of a v-gel SGAD compared with an endotracheal tube deserves some consideration. Two limitations of using a SGAD include the following: seal integrity, particularly if the device is not properly sized or positioned; and risk of displacement during procedures, which can compromise gas delivery.
In conclusion, we recommend the use of a heated anesthesia breathing circuit to provide thermal support to rabbits during anesthesia. Further research is recommended to measure and compare the endotracheal temperature and the efficacy of a heated breathing circuit for thermal support during anesthesia for other laboratory animal species.
Study Design Timeline.
Heated anesthesia circuit (A) connected to the veterinary anesthesia machine and (B) connected to the patient. The patient’s head and the skin below a 10 × 10 cm fenestration was exposed and an absorbent pad and conductive warming blanket set to 104 °F was placed underneath the patient. The 2 laser points reflect the spot where the average temperature in the pinna was taken.
Mean change of temperatures comparing anesthesia events with a heated breathing circuit, and with only conductive heating (“standard care”) for (A) rectal and (B) pinna. Error bars represent the 95% CI. The asterisks are differences from baseline and plus signs are differences between the 2 groups at that time point.
Box plot of (A) time to full recovery from anesthesia and (B) time to extubation. Mean with SEM represented.
Contributor Notes