Respiratory and antinociceptive effects of dexmedetomidine and doxapram in ball pythons (Python regius)

Alyssa A. Karklus Department of Comparative Biosciences, School of Veterinary Medicine, University of Wisconsin-Madison, Madison, WI 53706.

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Kurt K. Sladky Department of Surgical Sciences, School of Veterinary Medicine, University of Wisconsin-Madison, Madison, WI 53706.

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Stephen M. Johnson Department of Comparative Biosciences, School of Veterinary Medicine, University of Wisconsin-Madison, Madison, WI 53706.

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Abstract

OBJECTIVE

To determine the effects of dexmedetomidine, doxapram, and dexmedetomidine plus doxapram on ventilation (e), breath frequency, and tidal volume (Vt) in ball pythons (Python regius) and of doxapram on the thermal antinociceptive efficacy of dexmedetomidine.

ANIMALS

14 ball pythons.

PROCEDURES

Respiratory effects of dexmedetomidine and doxapram were assessed with whole-body, closed-chamber plethysmography, which allowed for estimates of e and Vt. In the first experiment of this study with a complete crossover design, snakes were injected, SC, with saline (0.9% NaCl) solution, dexmedetomidine (0.1 mg/kg), doxapram (10 mg/kg), or dexmedetomidine and doxapram, and breath frequency, e, and Vt were measured before and every 30 minutes thereafter, through 240 minutes. In the second experiment, antinociceptive efficacy of saline solution, dexmedetomidine, and dexmedetomidine plus doxapram was assessed by measuring thermal withdrawal latencies before and 60 minutes after SC injection.

RESULTS

Dexmedetomidine significantly decreased breath frequency and increased Vt but did not affect e at all time points, compared with baseline. Doxapram significantly increased e, breath frequency, and Vt at 60 minutes after injection, compared with saline solution. The combination of dexmedetomidine and doxapram, compared with dexmedetomidine alone, significantly increased e at 30 and 60 minutes after injection and did not affect breath frequency and Vt at all time points. Thermal withdrawal latencies significantly increased when snakes received dexmedetomidine or dexmedetomidine plus doxapram, versus saline solution.

CONCLUSIONS AND CLINICAL RELEVANCE

Concurrent administration of doxapram may mitigate the dexmedetomidine-induced reduction of breathing frequency without disrupting thermal antinociceptive efficacy in ball pythons.

Abstract

OBJECTIVE

To determine the effects of dexmedetomidine, doxapram, and dexmedetomidine plus doxapram on ventilation (e), breath frequency, and tidal volume (Vt) in ball pythons (Python regius) and of doxapram on the thermal antinociceptive efficacy of dexmedetomidine.

ANIMALS

14 ball pythons.

PROCEDURES

Respiratory effects of dexmedetomidine and doxapram were assessed with whole-body, closed-chamber plethysmography, which allowed for estimates of e and Vt. In the first experiment of this study with a complete crossover design, snakes were injected, SC, with saline (0.9% NaCl) solution, dexmedetomidine (0.1 mg/kg), doxapram (10 mg/kg), or dexmedetomidine and doxapram, and breath frequency, e, and Vt were measured before and every 30 minutes thereafter, through 240 minutes. In the second experiment, antinociceptive efficacy of saline solution, dexmedetomidine, and dexmedetomidine plus doxapram was assessed by measuring thermal withdrawal latencies before and 60 minutes after SC injection.

RESULTS

Dexmedetomidine significantly decreased breath frequency and increased Vt but did not affect e at all time points, compared with baseline. Doxapram significantly increased e, breath frequency, and Vt at 60 minutes after injection, compared with saline solution. The combination of dexmedetomidine and doxapram, compared with dexmedetomidine alone, significantly increased e at 30 and 60 minutes after injection and did not affect breath frequency and Vt at all time points. Thermal withdrawal latencies significantly increased when snakes received dexmedetomidine or dexmedetomidine plus doxapram, versus saline solution.

CONCLUSIONS AND CLINICAL RELEVANCE

Concurrent administration of doxapram may mitigate the dexmedetomidine-induced reduction of breathing frequency without disrupting thermal antinociceptive efficacy in ball pythons.

Introduction

Untreated pain and causes of stress in reptiles are associated with immunocompromise, negative energy balance, and delayed wound healing.1 Well-defined guidelines exist regarding pain assessment and management in mammals,2,3,4 but not for reptiles, for which the neural mechanisms of pain are poorly understood and effective analgesic drugs are frequently lacking.5,6,7,8,9 Consequently, analgesia may infrequently be provided to reptilian patients in practice.5

Opioids are effective analgesics in turtles and lizards,10,11,12,13,14,15 but opioids are inconsistently effective analgesics in snakes, with many opioids failing to provide adequate antinociception in laboratory models of pain.11,16,17 Nonsteroidal anti-inflammatory drugs are commonly used in practice to mitigate signs of post-operative pain in reptiles; however, the few studies9 with reptiles, including snakes, reveal that NSAIDs have minimal efficacy.

Given the questionable efficacy of opioids and NSAIDs in snakes, we recently investigated and demonstrated that dexmedetomidine, an α2-adrenoceptor agonist, is a candidate analgesic for ball pythons (Python regius).18 α2-Adrenoceptor agonists are commonly used as an analgesic or anesthetic induction agent in veterinary medicine, particularly for mammals.19,20,21,22 Several α2-adrenoceptor agonists provide sedative and antinociceptive effects in turtles and lizards.23,24,25,26 Dexmedetomidine provides thermal antinociception for at least 8 hours in ball pythons, but it also significantly decreases breath frequency.18 α2-Adrenoceptor agonists acting at brainstem and peripheral α2-adrenoceptors27 cause inhibition of breathing in mammals,28,29,30,31,32 amphibians,33 and reptiles.23,26 Because respiratory depression in reptiles may alter arterial blood gases and reduce hypoxic ventilatory responses, leading to hypoxemia and tissue hypoxia,34 dexmedetomidine-induced respiratory depression in snakes could have similar physiologic effects. Reptiles are generally more tolerant of hypoxia than mammals,35,36 with many adaptations to cope with hypoxia.35,36,37,38,39,40,41,42 However, respiratory depression could negatively impact snakes, especially if they are already compromised by respiratory disease, hemorrhage, hypovolemia, or cardiac disturbances (eg, congenital heart defects, endocarditis, or heart failure). Respiratory depression induced by an α2-adrenoceptor agonist could also prolong the time to recover from trauma or surgical procedures, as well as prolong the time to expel volatile anesthetics in the postoperative period. Therefore, countering respiratory depression induced by dexmedetomidine is ideal, and doxapram, a respiratory stimulant, may do this.

Doxapram is a potassium channel blocker that acts on central respiratory centers and peripheral chemoreceptors to stimulate breathing in a dose-dependent manner.43,44 Doxapram is efficacious in crocodiles45 and, in anecdotal reports,46 stimulates breathing in reptilian patients in emergency settings, but no studies have yet specifically included an evaluation of doxapram in snakes. However, because reptiles have central respiratory centers and peripheral chemoreceptors similar to mammals,47,48,49 doxapram is expected to similarly stimulate breathing in snakes.

The primary goal of the study presented here was to determine whether the combination of dexmedetomidine and doxapram could effectively provide adequate nociception (dexmedetomidine) and simultaneously mitigate dexmedetomidine-induced respiratory depression (doxapram). First, dexmedetomidine-induced respiratory depression was confirmed. Then, the respiratory effects of doxapram alone and in combination with dexmedetomidine were determined. Lastly, the antinociceptive effects of dexmedetomidine alone and combined with doxapram were determined.

Materials and Methods

Animals

Fourteen ball pythons with mean (SEM; range) body weight of 243 g (35 g; 55 to 430 g) were obtained from multiple commercial vendorsa–c throughout the course of the study. The sex of each snake was not determined. All snakes were housed in the Animal Resource Center at the School of Veterinary Medicine, University of Wisconsin-Madison. Snakes were kept in standard laboratory enclosures (width, 26 cm; depth, 48 cm; height, 20 cm) and provided with a hideawayd and a water source at all times. Snake enclosures were cleaned at least weekly by animal care technicians but more frequently when the enclosures were wet or soiled. Environmental temperature was maintained between 25°C and 29°C, and snakes were subjected to a 12-hour light cycle. Snakes were fed thawed mice or neonatal rats once weekly (Friday afternoons). All tests were performed Monday through Friday, and no tests were performed < 60 hours after feeding. The study was approved by the Institutional Animal Care and Use Committee (protocol No. V5710).

Dose selection

A dose of 0.1 mg of dexmedetomidine/kg was used for the present study on the basis of results of a previous study18 of ball pythons, in which this dose was effective for thermal antinociception and associated with decreased breath frequency. A dose of 10 mg of doxapram/kg was selected on the basis of a previous study45 with alligators, in which an increase in breathing frequency was noted after its administration.

Respiratory experiment

A complete crossover-design experiment was conducted to evaluate the effects of dexmedetomidine alone, doxapram alone, and the 2 drugs in combination on changes in breathing. Respiratory function was measured in snakes (n = 11) through unrestrained, whole-body, closed-chamber plethysmography in a room maintained between 26°C and 28°C (measured by a thermometer next to the chamber). Snakes that weighed < 300 g were placed in a small air-tight chamber (inner dimensions, 10 × 20 × 5 cm), and snakes > 300 g were placed in a large air-tight chamber (inner dimensions, 22 × 11 × 7 cm) with a transparent lid (Figure 1). The chambers were large enough such that the snakes could move freely. Room air (21% O2) was supplied at a rate of approximately 0.10 L/min into one port of the chamber, verified with a flowmeter,e and room air was removed from another port at the same verified rate with the facility's vacuum system. The chamber was connected via a different port to a differential pressure transducer.f The pressure transducer was also connected to a similarly sized chamber (reference chamber). The reference chamber helped to minimize artifacts resulting from changes in ambient barometric pressure or noise, as noted in previous studies50,51 involving whole-body plethysmography. A visual barrier was placed around the chamber to minimize visual cues to the snake that could induce its movement (and therefore artifacts) or alter its breathing. The valves that controlled airflow for the flowmeter and vacuum were separate from the chamber, outside of the visual barrier, so that movements and vibrations resulting from adjusting the valves would not startle the snake (eg, a light tap on the chamber markedly increased breathing frequency for several minutes). Key improvements from our previously published18 method were a transparent chamber lid, so a snake did not sleep or enter a torpor-like state that can affect its breathing pattern during the 12-hour light cycle52,53; the airflow control valves were located away from the chamber;and e and Vt were calculated from a linear calibration curve that was generated by making expiratory traces with known volumes of a syringe, that mimicked snake breathing, attached to the recoding chamber. This technique allowed for reasonable estimation of e and Vt.

Figure 1
Figure 1

Drawing of the whole-body, closed-chamber plethysmography apparatus used for the respiratory experiment with 11 ball pythons (Python regius). A—The snake was placed into an air-tight chamber with a transparent lid. Airflow into and out of the chamber was manually controlled with valves located approximately 20 cm from the chamber to minimize chamber disturbances. An opaque barrier (visual barrier) surrounded the chamber to minimize visual stimulation or disturbance of the snake. The chamber was connected to a reference chamber of similar size via plastic tubing and a differential pressure transducer, which detected pressure changes attributable to breathing movements. An amplifier converted the pressure changes into voltage signals that were subsequently recorded by a data acquisition system to be later analyzed with computer software. B—A representative respiratory-related voltage trace (created by voltage signals) shows waveforms consistent with normal breathing plus an area of movement artifact. Downward deflections represent inspiratory movements, and upward deflections represent expiratory movements.

Citation: American Journal of Veterinary Research 82, 1; 10.2460/ajvr.82.1.11

Prior to the respiratory experiment, the snakes were conditioned to the chamber and portions of the experimental protocol for at least 6 hours on 2 or more days. Conditioning was conducted by placing a snake into the chamber and alternating between 20 minutes of airflow and 10 minutes of no airflow (ie, air inflow and outflow ports closed), with data recorded during the 10-minute period of no airflow. Alternating periods of airflow and no airflow prevented accumulation of CO2 in the chamber, which can stimulate breathing,54 increase Vt, increase breath frequency,

and increase e35,55,56,57,58 and allowed for nearly continuous recording for > 8 hours.18 When the inflow and outflow ports were closed, the pressure transducer detected pressure changes attributable to breathing movements. An amplifier converted these pressure changes to voltage signals, which were subsequently recorded by a data acquisition systemg and later analyzed with computer software.h On the digital tracing produced by the pressure transducer, upward deflections indicated increased pressure (expiration) and downward deflections indicated decreased pressure (inspiration).

For the experiment, snakes were placed in the chamber for approximately 2 hours to allow the snake to reach a steady rate and quality of breathing. For the time-control part of the experiment, the breathing of non-sedated snakes was recorded for 10 out of every 30 minutes (20 minutes of airflow and 10 minutes of no airflow, when breathing recorded) for 4 to 6 hours. For the treatment part of the experiment, the snakes were allowed to reach steady baseline breathing, then the chamber was opened, and the snakes randomly received the first of 4 treatments: SC injection of saline (0.9% NaCl) solution (isovolumetric to doxapram dose), dexmedetomidinei (0.1 mg/kg), doxapramj (10 mg/kg), or dexmedetomidine plus doxapram. Treatments were injected SC into the cranial one-third of the body at the approximate level of the heart to avoid the hepatic first-pass effect. Breathing data were collected for 240 minutes after injection. A minimum 7-day washout period was maintained between treatments.

Thermal antinociception study

A blinded, randomized, within-subjects complete crossover-design experiment was conducted to determine whether doxapram would alter the antinociceptive efficacy of dexmedetomidine. This experiment included use of a modified Hargreaves plantar thermal limb withdrawal latency apparatus,k with the original designed to assess nociception and thermal antinociception in rodents59 and which our laboratory had previously adapted for use with reptiles.10,11,12,13 The testing enclosure was comprised of 3 contiguous chambers on an elevated glass surface (allowing 3 snakes to be tested simultaneously), through which a noxious thermal stimulus could be applied from below. An opaque panel separated each chamber, such that the snakes could not see each other but could see the observer. The experiment was conducted at an ambient temperature of 26°C to 28°C. In the modified version, the Hargreaves apparatus was used to apply a noxious thermal stimulus, via an infrared beam, to the cranioventral one-third of the snake's body.16,18 Following activation of the infrared heat source, TWLs were measured automatically with a motion-sensitive timer. Stimulation strength of the infrared beam was adjusted to produce baseline latencies of 8 to 12 seconds. To minimize thermal injury, the heating apparatus automatically turned off at 32.6 seconds.

Three snakes were used for the thermal antinociception study, under 3 experimental conditions. Prior to the day of testing, snakes were conditioned to the test chamber and the presence of an observer for 2 to 4 hours, with intermittent exposure to a beam from the infrared heat source to minimize the snakes' anticipatory behaviors during testing. On the day of testing, snakes were acclimated to the test chamber for at least 30 minutes. Baseline TWLs were then recorded every 5 minutes for at least 2 trials. If the 2 TWLs differed by > 10 seconds, a third trial was performed. Mean baseline TWL was calculated from these trials. A blinded observer then administered a randomly assigned, SC injection of saline solution (isovolumetric to doxapram dose), dexmedetomidine (0.1 mg/kg) plus saline solution, or dexmedetomidine plus doxapram (10 mg/kg) to each snake. Thermal withdrawal latencies were measured 60 minutes after injection, the time of peak dexmedetomidine effects.18 A 7-day washout period was maintained between treatments.

Data analysis

Within each 10-minute data acquisition period for the respiratory experiment, all upward-deflecting (expiration), respiratory-related voltage traces (without obvious motion artifacts) were marked for analysis with commercially available software.h The area of the upward deflections was added and divided by the total number of breaths (breath frequency) to obtain the area per breath. Area per breath was then converted to volume per breath by comparison to a linear calibration curve generated by making similar expiratory traces (duration and shape) with known volumes of a syringe attached to the recording chamber. Separate calibration curves were generated for the small and large recording chambers. The calibration curve permitted calculation of e (mL/min/kg) and Vt (mL/breath/kg) by using the breath frequency data (e = Vt × breathing frequency). Mean values for respiratory variables were calculated for each snake within each 10-minute period and then mean values of all snakes for each 10-minute period of data acquisition were averaged. Whole-body plethysmography is an indirect and qualitative method to evaluate respiratory function because of the inherent pressure variability within the chamber.51,60 Thus, determination of actual values for e and Vt was not possible, so e and Vt data were regarded as estimates of their actual values. Nevertheless, this method provided information noninvasively with awake snakes behaving normally during a 12-hour light cycle.

Respiratory and thermal antinociception data were analyzed with a 2-way repeated-measures ANOVA with commercially available software.l Normality and equal assumption tests were not always satisfied, despite data transformation. Given that a 2-way repeated-measures ANOVA is not applicable for nonparametric data, the statistical results were interpreted cautiously. Post hoc comparisons were conducted with the Student-Newman-Keuls test. All data were reported as mean ± SEM, and values of P < 0.05 were considered significant.

Results

Respiratory experiment

Variability in the shape of voltage traces recorded by the plethysmograph was considerable, likely because of each snake's posture in the chamber and variability in each snake's movements during breathing (Figure 2). Nevertheless, rhythmic upward and downward traces indicated that breathing was observed and could be differentiated from movement artifacts. For the time-control and saline solution and dexmedetomidine treatment parts of the experiment, baseline e, breath frequency, and Vt were similar, which indicated that all snakes were breathing similarly prior to the start of the experiment (Figure 3).

Figure 2
Figure 2

Representative respiratory-related voltage traces obtained with whole-body, closed-chamber plethysmography illustrating breathing movements at 0 (baseline). 30, and 240 minutes under time-control conditions (no injection; A), with saline solution (B), or with 0.1 mg of dexmedetomidine/kg (C). Breathing frequency was assessed 2 hours after acclimation to the chamber (baseline) and then every 30 minutes for 240 minutes after SC injection. Saline solution transiently increased breathing frequency, whereas dexmedetomidine produced a highly irregular breathing pattern and decreased breathing frequency and increased Vt.

Citation: American Journal of Veterinary Research 82, 1; 10.2460/ajvr.82.1.11

Figure 3
Figure 3

Mean ± SEM of e (A), breath frequency (B), and Vt (C) during a 240-minute period for ball pythons (n = 11) under time-control conditions (no injection; squares) or after SC injection of saline solution (white circles) or 0.1 mg of dexmedetomidine/kg (black circles) in accordance with a complete crossover design. Data were acquired by use of whole-body, closed-chamber plethysmography before administration and every 30 minutes after administration for 240 minutes. *Within a treatment, value significantly (P < 0.05) differs from the value at 0 minutes (baseline). †Within a time point, value significantly (P < 0.05) differs from the value for time-control conditions. ‡Within a time point, value significantly (P < 0.05) differs from the value for dexmedetomidine §Within a time point, value significantly (P < 0.05) differs from the value for saline solution. llWithin an experiment, value for a treatment significantly (P < 0.05) differs from the value for time-control conditions (ie, drug effect). ¶Within an experiment, value for a treatment significantly (P < 0.05) differs from the value for saline solution (ie, drug effect). #Within an experiment, value for a treatment significantly (P < 0.05) differs from the value for dexmedetomidine (ie, drug effect).

Citation: American Journal of Veterinary Research 82, 1; 10.2460/ajvr.82.1.11

For the time-control part of the experiment, e was not altered through 240 minutes from baseline (mean ± SEM, 1.22 ± 0.22 mL/min/kg; Figure 3), but breath frequency was significantly (P < 0.036) different between baseline and all other time points (mean ± SEM; baseline, 5.59 ± 1.33 breaths/min vs 240 minutes, 2.56 ± 0.45 breaths/min). No significant differences were noted between Vt at baseline (0.31 ± 0.07 mL/breath/kg) and all other time points.

Saline solution significantly (P < 0.001) increased e at 30 (2.93 ± 0.64 mL/min/kg) and 60 minutes (2.11 ± 0.40 mL/min/kg), compared with baseline (1.21 ± 0.31 mL/min/kg), without significantly affecting Vt (Figure 3). A concurrent significant (P < 0.005) increase in breath frequency was noted (baseline, 4.85 ± 0.90 breaths/min vs 30 minutes, 9.26 ± 0.79 breaths/min or 60 minutes, 7.36 ± 0.85 breaths/min). Breath frequency decreased thereafter and achieved a value near baseline at 120 minutes through 240 minutes. Breath frequency increased with saline solution (P < 0.001) but not with time-control or dexmedetomidine.

Dexmedetomidine caused large expiratory and inspiratory deflections and irregular breathing patterns, compared with time-control and saline solution, starting at 30 minutes after administration. Compared with baseline (1.34 ± 0.31 mL/min/kg), e did not significantly differ throughout the experiment (Figure 3). Breath frequency at all time points was significantly (P < 0.001) decreased from baseline (eg, 5.76 ± 1.49 breaths/min vs 2.09 ± 0.57 breaths/min [240 minutes]). Overall, breath frequency was not significantly (P = 0.497) different between dexmedetomidine and time-control. Tidal volume with dexmedetomidine was significantly higher for all time points (range, 0.40 to 1.04 mL/breath/kg), compared with baseline (mean ± SEM, 0.25 ± 0.02 mL/breath/kg) and was overall significantly higher (P = 0.009), compared with saline solution. Dexmedetomidine did not alter e.

Thirty and 60 minutes after doxapram administration, e was significantly (P < 0.001) increased, compared with baseline (baseline, 1.43 ± 0.35 mL/min/kg; 30 minutes, 7.20 ± 1.09 mL/min/kg; 60 minutes, 3.96 ± 0.67 mL/min/kg; Figures 4 and 5). Breath frequency at each time point versus baseline was not affected by doxapram throughout the experiment (P > 0.294). Tidal volume was also significantly (P < 0.02) increased at 30 and 60 minutes (baseline, 0.42 ± 0.14 mL/breath/kg; 30 minutes, 1.26 ± 0.28 mL/breath/kg; 60 minutes, 0.80 ± 0.13 mL/breath/kg). Overall, e and Vt with doxapram were significantly (P = 0.002 and P < 0.001, respectively) increased versus saline solution.

Figure 4
Figure 4

Representative respiratory-related traces obtained with whole-body, closed-chamber plethysmography, illustrating breathing movements in a ball python prior to (baseline) and 30 and 240 minutes after SC injection of saline solution (A) or 10 mg of doxapram/kg (B). Note the increased breathing frequency, primarily attributable to increased Vt, 30 minutes after doxapram administration.

Citation: American Journal of Veterinary Research 82, 1; 10.2460/ajvr.82.1.11

Figure 5
Figure 5

Mean ± SEM e (A), breath frequency (B), and Vt (C), during a 240-minute period for ball pythons (n = 11) after SC injection of saline solution (circles) or 10 mg of doxapram/kg (triangles) in accordance with a complete crossover design. See Figure 3 for remainder of key.

Citation: American Journal of Veterinary Research 82, 1; 10.2460/ajvr.82.1.11

After administration of the combination of dexmedetomidine and doxapram, breath frequency increased and breathing pattern was less irregular, compared with dexmedetomidine alone (Figure 6). With the combination, e significantly increased from 1.23 ± 0.20 mL/min/kg at baseline to 3.52 ± 0.57 mL/min/kg (maximum value) at 30 minutes and to 2.62 ± 0.34 mL/min/kg at 60 minutes, then gradually decreased to values near baseline and similar to those for saline solution and dexmedetomidine (Figure 7). Overall, e was significantly (P = 0.029) higher, compared with dexmedetomidine alone. Breath frequency did not increase from baseline with the combination (P > 0.077), and added doxapram, compared with dexmedetomidine alone, prevented a significant (P = 0.016) decrease in breath frequency at 30 and 60 minutes. Dexmedetomidine plus doxapram significantly (P = 0.041) increased Vt from 0.25 ± 0.04 mL/breath/kg at baseline to 0.61 ± 0.11 mL/breath/kg (maximum value) at 30 minutes, but baseline Vt did not significantly differ from other time points. Tidal volume overall and at 120 minutes through 210 minutes for dexmedetomidine plus doxapram was significantly (P = 0.013 and P < 0.023, respectively) decreased, compared with dexmedetomidine alone.

Figure 6
Figure 6

Representative respiratory-related traces obtained with whole-body, closed-chamber plethysmography, illustrating breathing movements in a ball python prior to (baseline) and 30 and 240 minutes after SC injection of 0.1 mg of dexmedetomidine/kg (A) or dexmedetomidine plus 10 mg of doxapram/kg (B). Note that coadministration of dexmedetomidine and doxapram maintained breath frequency, compared with baseline, and a more regular breathing pattern, compared with dexmedetomidine alone.

Citation: American Journal of Veterinary Research 82, 1; 10.2460/ajvr.82.1.11

Figure 7
Figure 7

Mean ± SEM e (A), breath frequency (B), and Vt (C), during a 240-minute period for ball pythons (n = 11) after SC injection of saline solution (white circles), 0.1 mg of dexmedetomidine/kg (black circles), or dexmedetomidine plus 10 mg of doxapram/kg (diamonds) in accordance with a complete crossover design. See Figure 3 for remainder of key.

Citation: American Journal of Veterinary Research 82, 1; 10.2460/ajvr.82.1.11

Thermal antinociception experiment

Sixty minutes after injection of saline solution, TWL (mean ± SEM, 9.5 ± 1.5 seconds) did not significantly (P = 0.395) differ from baseline (Figure 8). Dexmedetomidine alone or combined with doxapram significantly (P < 0.001) increased TWL from baseline (dexmedetomidine: 10.7 ± 1.5 seconds [baseline], 31.9 ± 0.7 seconds [60 minutes]; dexmedetomidine and doxapram: 9.3 ± 1.8 seconds [baseline], 27.9 ± 1.5 seconds [60 minutes]).

Figure 8
Figure 8

Mean ± SEM noxious TWL for ball pythons (n = 3) before (white bars) and 60 minutes (black bars) after SC injection of saline (0.9 % NaCl) solution, 0.1 mg of dexmedetomidine/kg, or dexmedetomidine plus 10 mg of doxapram/kg, in accordance with a complete crossover design. *Within a treatment, value significantly (P < 0.05) differs from the value at baseline.

Citation: American Journal of Veterinary Research 82, 1; 10.2460/ajvr.82.1.11

Discussion

To our knowledge, this study revealed the first successful use of whole-body plethysmography without anesthesia, sedation, or invasive equipment to estimate e and Vt in unrestrained ball pythons. With this novel method, we demonstrated that within 60 minutes of dexmedetomidine administration, e and breath frequency decreased and normal breathing pattern was disrupted, compared with saline solution. Snakes compensated for the decreased breath frequency by increasing Vt, which caused e to remain at preinjection values. We also showed that doxapram increased e by increasing breath frequency and Vt and mitigated dexmedetomidine's effect on the breathing pattern. Plus, concurrent administration of dexmedetomidine and doxapram did not decrease the antinociceptive effect of dexmedetomidine. Thus, dexmedetomidine may be a candidate antinociceptive drug for use in snakes, and the irregular breathing pattern induced by dexmedetomidine may be ameliorated with coadministration of doxapram.

The method used to collect respiratory data in the present study was an improvement of techniques used in previous studies,55,61,62,63 which involved plethysmography with anesthesia, invasive instrumentation, or physical restraint. Though these techniques minimize movement artifacts, acquired respiratory data may not represent physiologic (normal) breathing.35 Unrestrained, whole-body plethysmography reduces artifacts and experimental variability that are often associated with invasive techniques because of the need for anesthesia and restraints for invasive techniques.60 To mitigate the occurrence of artifacts resulting from working with unrestrained awake snakes in the present study, snakes were acclimated to the transparent chamber before beginning the study, light was permitted through the transparent chamber to prevent snakes from sleeping or entering a torpor-like state, a visual barrier surrounded the chamber to prevent visual cues to the snakes, airflow valves were placed away from the chamber, and periods of airflow with no airflow (closed chamber), during which data were acquired, were alternated to prevent excessive CO2 accumulation. For each 10-minute period when the chamber was closed, no important alterations in breathing pattern suggestive of excessive CO2 accumulation in the chamber were observed. Yet, absolute Vt cannot be determined with whole-body plethysmography because of its well-known, inherent limitations.64 However, rather than using an invasive or complex methodology to obtain Vt and the other respiratory parameters, we elected to use a model that minimized restraint and natural activity (movement) of a snake to observe its undisturbed breathing. Nevertheless, Vt and e could be reasonably estimated with this model and drug-dependent changes in Vt and e were quantifiable.

The present study confirmed that dexmedetomidine decreases breath frequency in snakes, compared with saline solution, and revealed that e was unchanged from baseline. The lack of an increase in e with dexmedetomidine may have been secondary to rapid sedation of the snakes. In a previous study18 and repeatable in the present study, breath frequency reached a steady state of approximately 4 breaths/min with saline solution, whereas breath frequency reached a steady state of approximately 2 breaths/min with dexmedetomidine. However, the immediate response to saline solution differed between studies. Little to no change in breath frequency was observed immediately after injection of saline solution in the previous study,18 whereas breath frequency nearly doubled in the present study. These differences may be because of the different baseline breath frequencies and plethysmographic methods between studies. Surprisingly, breath frequency, without concurrent increased Vt, in the time-control part of the experiment was similar to that observed for dexmedetomidine-injected snakes. The importance of this observation is unclear, however.

Snakes and other reptiles breathe with a distinct pattern of active inspiration, active expiration, and a breath-holding period.35,65,66 Dexmedetomidine caused a dramatic change in the overall respiratory pattern of the ball pythons in the present study by making Vt irregular and inspiration and expiration vary in duration and amplitude throughout the entire 240-minute respiratory experiment. These changes may have been attributable to dexmedetomidine's effect on the respiratory rhythm–generating neurons in the brainstem. Disrupted breathing patterns caused by dexmedetomidine could have negative consequences (eg, increased arterial hypoxia and hypercapnia) for healthy and unhealthy snakes. Although e was not altered by dexmedetomidine, breathing disruptions may cause moment-by-moment fluctuations in Pao2 and Paco2. Switching from regularly spaced breaths to episodic breaths (ie, clusters of breaths) increases CO2 retention and subsequent acidemia in turtles,67 thereby illustrating the importance of breath patterns to blood gas homeostasis.

In reptiles, hypoxia and hypoxemia induce compensatory hypothermia and reduced metabolic rate.39,41,68,69 Reduced body temperature and metabolism could prolong recovery from sedation or anesthesia70,71 and could negatively impact wound healing.72 However, veterinarians frequently use supplemental heat sources to keep reptiles within the optimal temperature range.6,71 Alternatively, increased body temperature is associated with increased respiratory rate, oxygen uptake, e, and metabolic rate.62,73,74 Yet, a snake administered dexmedetomidine may not be able to respond accordingly to increased temperature. Ultimately, the effects of dexmedetomidine on hypoxic and hypercapnic responses in snakes are unknown; however, for snakes affected by trauma or disease, dexmedetomidine may exacerbate compromised respiratory function and prolong recovery from anesthesia.

The present study findings of doxapram-induced increase of e and, in some situations, of Vt were consistent with those previously reported for mammals43,75,76 and crocodilians.45 However, unlike previous studies, breath frequency after doxapram administration to the ball pythons of the present study did not significantly increase. Instead, the results aligned with those observed with people, in which doxapram's primary action on respiration is through changes in Vt and e.77 Unexpectedly, doxapram did not increase breath frequency similar to that observed immediately after injection of saline solution. Possibly, breath duration increased secondary to a 3-fold, doxapram-induced increase in Vt, such that breath frequency did not significantly increase concurrently.

In mammalian studies43,44,75,76 of doxapram, the effects of single doses are short-lived, typically lasting only 5 to 15 minutes. In a study45 of alligators, however, a measurable effect of doxapram on respiration was detected for hours after intra-arterial administration. In the present study of ball pythons, doxapram had respiratory stimulant effects for the first 60 minutes after injection. Doxapram's increased duration of action in alligators and snakes may be because of the lower metabolic rate of reptiles and subsequent slower drug metabolism.71

In the present study, results obtained after administration of the combination of dexmedetomidine and doxapram suggested that doxapram may counteract the respiratory disturbances caused by dexmedetomidine. With the addition of doxapram, the dexmedetomidine-induced decrease in breath frequency and irregular breathing pattern were blunted (ie, breath frequency was unchanged from baseline during the first 60 minutes after injection). Tidal volume was likely stable secondary to doxapram's blunting of the dexmedetomidine-induced decrease in breath frequency. However, the exact mechanism by which these drugs caused these changes is unknown, but we speculated that the stimulus to breathe induced by doxapram may have overridden the decreased breath frequency and stabilized the breathing pattern to reduce the disturbances caused by dexmedetomidine.

We did not anticipate any interference with the antinociceptive efficacy of dexmedetomidine by the addition of doxapram on the basis that they act on different types of receptors in the CNS.27,43 The thermal antinociception experiment showed that combined dexmedetomidine and doxapram did not block the antinociceptive efficacy of dexmedetomidine. Results of the respiratory and thermal antinociception experiments indicated that doxapram may effectively ameliorate respiratory disturbances induced by dexmedetomidine, while preserving its thermal antinociceptive effects. Our findings were similar to those of previous animal76 and human78 studies that indicate that doxapram can counteract the respiratory depressant effects of morphine without disrupting its thermal antinociceptive efficacy. However, in contrast to one of those studies76 in which only transient improvement of opioid-induced respiratory depression was detected after doxapram administration, we observed persistent improvement of dexmedetomidine-induced respiratory depression in the present study. When combined with dexmedetomidine, doxapram maintained a more normal respiratory rate, Vt, and breathing pattern over several hours. After the initial respiratory stimulation (ie, increased e) from doxapram waned, breath frequency was unchanged, compared with baseline, and snakes never demonstrated the decreased frequency or aberrant respiratory patterns observed when dexmedetomidine was administered alone. The snakes' slow metabolic rate and subsequent slow doxapram metabolism may have permitted sustained mitigation of dexmedetomidine-induced respiratory abnormalities.

Dexmedetomidine may be an important analgesic option for snakes, and doxapram may mitigate respiratory disturbances caused by dexmedetomidine, while also preserving its antinociceptive function. The results reported here may be applicable to other reptiles, such as chelonians and lizards, in which breath frequency also decreases after administration of α2-adrenoceptor agonists.23,26 However, doxapram is associated with decreased cerebral blood flow in dogs and goats79,80 and increased cardiac work in dogs,81 which may limit the usefulness of doxapram in reptiles. Yet, whether doxapram has these same effects in snakes is unknown. A clinically relevant model of pain for snakes needs to be developed before the analgesic effects of dexmedetomidine alone and combined dexmedetomidine and doxapram can be fully understood.

Acknowledgments

Supported in part by the Merial Summer Scholars Program (Karklus); Department of Comparative Biosciences, (Johnson); and a Companion Animal Fund grant, School of Veterinary Medicine, University of Wisconsin-Madison.

The authors declare that there were no conflicts of interest.

Abbreviations

TWL

Thermal withdrawal latency

e

Ventilation

Vt

Tidal volume

Footnotes

a.

Reptile Rapture, Monona, Wis.

b.

Pet Supplies Plus, Madison, Wis.

c.

LLL Reptile, Oceanside, Calif.

d.

Super Pet Mini Igloo, Petsmart, Phoenix, Ariz.

e.

Mass flow meter, model 4140, TSI Incorporated, Shoreview, Minn.

f.

Model DP45-14, Validyne Engineering Corp, Northridge, Calif.

g.

DigiData 1200, Axon Instruments, Sunnyvale, Calif.

h.

pClamp, Axon Instruments, Sunnyvale, Calif.

i.

Dexdomitor, 0.5 mg/mL, Iron Corp, Espoo, Finland.

j.

Dopram, West-Ward, Eatontown, NJ.

k.

Ugo Basile plantar analgesia instrument, model 37370, Ugo Basile Co, Comerio, Italy.

l.

Sigma Stat, Jandel Scientific Software, San Rafael, Calif.

References

  • 1.

    Mosley C. Pain and nociception in reptiles. Vet Clin North Am Exot Anim Pract 2011;14:45 60.

  • 2.

    Holton L, Reid J, Scott EM, et al. Development of a behavior-based scale to measure acute pain in dogs. Vet Rec 2001;148:525 531.

  • 3.

    Reid J, Scott EM, Calvo G, et al. Definitive Glasgow acute pain scale for cats: validation and intervention level. Vet Rec 2017;180:449 452.

  • 4.

    Epstein ME, Rodan I, Griffenhagen G, et al. 2015 AAHA/AAFP pain management guidelines for dogs and cats. J Feline Med Surg 2015;17:251 272.

  • 5.

    Read MR. Evaluation of the use of anesthesia and analgesia in reptiles. J Am Vet Med Assoc 2004;224:547 552.

  • 6.

    Sladky KK, Mans C. Clinical analgesia in reptiles. J Exot Pet Med 2012;21:158 167.

  • 7.

    Sladky KK. Analgesia. In: Mader DR, Divers S, eds. Current therapy in reptile medicine and surgery. 3rd ed. St Louis: Elsevier-Saunders, 2014;217 228.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 8.

    Mosley C. Reptile-specific considerations. In: Gaynor JS, Muir WW, eds. Handbook of veterinary pain management. 3rd ed. St Louis: Elsevier, 2015;42 60.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 9.

    Sladky KK, Mans C. Analgesia. In: Divers SJ, Stahl SJ, eds. Mader's reptile and amphibian medicine and surgery. St Louis: Elsevier, 2019;465 474.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 10.

    Sladky KK, Miletic V, Paul-Murphy J, et al. Analgesic efficacy and respiratory effects of butorphanol and morphine in turtles. J Am Vet Med Assoc 2007;230:1356 1362.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 11.

    Sladky KK, Kinney ME, Johnson SM. Analgesic efficacy of butorphanol and morphine in bearded dragons and corn snakes. J Am Vet Med Assoc 2008;233:267 273.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 12.

    Sladky KK, Kinney ME, Johnson SM. Effects of opioid receptor activation on thermal antinociception in red-eared slider turtles (Trachemys scripta). Am J Vet Res 2009;70:1072 1078.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 13.

    Baker BB, Sladky KK, Johnson SM. Evaluation of the analgesic effects of oral and subcutaneous tramadol administration in red-eared slider turtles. J Am Vet Med Assoc 2011;238:220 227.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 14.

    Kinney ME, Johnson SM, Sladky KK. Behavioral evaluation of red-eared slider turtles (Trachemys scripta elegans) administered either morphine or butorphanol following unilateral gonadectomy. J Herpetol Med Surg 2011;21:54 62.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 15.

    Leal WP, Carregaro AB, Bressan TF, et al. Antinociceptive efficacy of intramuscular administration of morphine sulfate and butorphanol tartrate in tegus (Salvator merianae). Am J Vet Res 2017;78:1019 1024.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 16.

    Kharbush RJ, Gutwillig A, Hartzler KE, et al. Antinociceptive and respiratory effects following application of transdermal fentanyl patches and assessment of brain μ-opioid receptor mRNA expression in ball pythons. Am J Vet Res 2017;78:785 795.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 17.

    Williams CJ, James LE, Bertelsen MF, et al. Tachycardia in response to remote capsaicin injection as a model for nociception in the ball python (Python regius). Vet Anaesth Analg 2016;43:429 434.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 18.

    Bunke LG, Sladky KK, Johnson SM. Antinociceptive efficacy and respiratory effects of dexmedetomidine in ball pythons (Python regius). Am J Vet Res 2018;79:718 726.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 19.

    Sinclair MD. A review of the physiological effects of α2-agonists related to the clinical use of medetomidine in small animal practice. Can Vet J 2003;44:885 897.

    • Search Google Scholar
    • Export Citation
  • 20.

    Murrell JC, Hellebrekers LJ. Medetomidine and dexmedetomidine: a review of cardiovascular effects and antinociceptive properties in the dog. Vet Anaesth Analg 2005;32:117 127.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 21.

    Kästner SBR. A2-agonists in sheep: a review. Vet Anaesth Analg 2006;33:79 96.

  • 22.

    Valverde A. Alpha-2 agonists as pain therapy in horses. Vet Clin North Am Equine Pract 2010;26:515 532.

  • 23.

    Sleeman JM, Gaynor J. Sedative and cardiopulmonary effects of medetomidine and reversal with atipamezole in desert tortoises (Gopherus agassizii). J Zoo Wildl Med 2000;31:28 35.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 24.

    Makau CM, Towett PK, Abelson KS, et al. Intrathecal administration of clonidine or yohimbine decreases the nociceptive behavior caused by formalin injection in the marsh terrapin (Pelomedusa subrufa). Brain Behav 2014;4:850 857.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 25.

    Makau CM, Towett PK, Abelson KS, et al. Modulation of formalin-induced pain-related behaviour by clonidine and yohimbine in the Speke's hinged tortoise (Kiniskys spekii). J Vet Pharmacol Ther 2017;40:439 446.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 26.

    Bisetto SP, Melo CF, Carregaro AB. Evaluation of sedative and antinociceptive effects of dexmedetomidine, midazolam and dexmedetomidine-midazolam in tegus (Salvator merianae). Vet Anaesth Analg 2018;45:320 328.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 27.

    Giovannoni MP, Ghelardini C, Vergelli C, et al. Alpha 2-agonists as analgesic agents. Med Res Rev 2009;29:339 368.

  • 28.

    Lerche P, Muir WW. Effect of medetomidine on breathing and inspiratory neuromuscular drive in conscious dogs. Am J Vet Res 2004;65:720 724.

  • 29.

    Tamiya J, Ide R, Takahashi M, et al. Effects of dexmedetomidine on cardiorespiratory regulation in spontaneously breathing newborn rats. Paediatr Anaesth 2014;24:1245 1251.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 30.

    Hilaire G, Viemari JC, Coulon P, et al. Modulation of the respiratory rhythm generator by the pontine noradrenergic A5 and A6 groups in rodents. Respir Physiol Neurobiol 2004;143:187 197.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 31.

    Oliveira LM, Moreira TS, Kuo FS, et al. α1- and α2-adrenergic receptors in the retrotrapezoid nucleus differentially regulate breathing in anesthetized adult rats. J Neurophysiol 2016;116:1036 1048.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 32.

    Tsuzawa K, Minoura Y, Takeda S, et al. Effects of α2-adorenoceptor agonist dexmedetomidine on respiratory rhythm generation of newborn rats. Neurosci Lett 2015;597:117 120.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 33.

    Fournier S, Kinkead R. Noradrenergic modulation of respiratory motor output during tadpole development: role of α-adrenoreceptors. J Exp Biol 2006;209:3685 3694.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 34.

    Malte CL, Bundgaard J, Jensen MS, et al. The effects of morphine on gas exchange, ventilation pattern and ventilatory responses to hypercapnia and hypoxia in dwarf caiman (Paleosuchus palpebrosus). Comp Biochem Physiol A Mol Integr Physiol 2018;222:60 65.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 35.

    Glass ML, Wood SC. Gas exchange and control of breathing in reptiles. Physiol Rev 1983;63:232 260.

  • 36.

    Bickler PE, Buck LT. Hypoxia tolerance in reptiles, amphibians, and fishes: life with variable oxygen availability. Annu Rev Physiol 2007;69:145 170.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 37.

    Boyer DR. Comparative effects of hypoxia on respiratory and cardiac function in reptiles. Physiol Zool 1966;39:307 316.

  • 38.

    Gratz RK. Ventilatory response of the diamondback water snake, Natrix rhombifera to hypoxia, hypercapnia and increased oxygen demand. J Comp Physiol B 1979;129:105 110.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 39.

    Wood SC, Hicks JW, Dupré RK. Hypoxic reptiles: blood gases, body temperature and control of breathing. Am Zool 1987;27:21 29.

  • 40.

    Ultsch GR. Ecology and physiology of hibernation and overwintering among freshwater fishes, turtles, and snakes. Biol Rev 1989;64:435 515.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 41.

    Hicks JW, Wang T. Hypometabolism in reptiles: behavioural and physiological mechanisms that reduce aerobic demands. Respir Physiol Neurobiol 2004;141:261 271.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 42.

    Jackson DC. Acid-base balance during hypoxic hypometabolism: selected vertebrate strategies. Respir Physiol Neurobiol 2004;141:273 283.

  • 43.

    Yost CS. A new look at the respiratory stimulant doxapram. CNS Drug Rev 2006;12:236 249.

  • 44.

    Heggem B. Doxapram. J Exot Pet Med 2011;20:237 240.

  • 45.

    Skovgaard N, Crossley DA, Wang T. Low cost of pulmonary ventilation in American alligators (Alligator mississippiensis) stimulated with doxapram. J Exp Biol 2016;219:933 936.

    • Search Google Scholar
    • Export Citation
  • 46.

    Martinez-Jimenez D, Hernandez-Divers SJ. Emergency care of reptiles. Vet Clin North Am Exot Anim Pract 2007;10:557 585.

  • 47.

    Smatresk NJ. Chemoreceptor modulation of endogenous respiratory rhythms in vertebrates. Am J Physiol 1990;259:R887 R897.

  • 48.

    Milsom WK, Burleson ML. Peripheral arterial chemoreceptors and the evolution of the carotid body. Respir Physiol Neurobiol 2007;157:4 11.

  • 49.

    Milsom WK. The phylogeny of central chemoreception. Respir Physiol Neurobiol 2010;173:195 200.

  • 50.

    Bartlett D Jr, Tenney SM. Control of breathing in experimental anemia. Respir Physiol 1970;10:384 395.

  • 51.

    Hernandez AB, Kirkness JP, Smith PL, et al. Novel whole body plethysmography system for the continuous characterization of sleep and breathing in a mouse. J Appl Physiol 2012;112:671 680.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 52.

    Coote JH. Respiratory and circulatory control during sleep. J Exp Biol 1982;100:223 244.

  • 53.

    Hicks JW, Riedesel ML. Diurnal ventilatory patterns in the garter snake, Thamnophis elegans. J Comp Physiol B 1983;149:503 510.

  • 54.

    Kimmel EC, Whitehead GS, Reboulet JE, et al. Carbon dioxide accumulation during small animal, whole body plethysmography: effects on ventilation, indices of airway function, and aerosol deposition. J Aerosol Med 2002;15:37 49.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 55.

    Furilla RA, Coates EL, Bartlett D Jr. The influence of venous CO2 on ventilation in garter snakes. Respir Physiol 1991;83:47 59.

  • 56.

    Nolan WF, Frankel HM. Ventilatory responses to CO2 at different body temperatures in the snake, Coluber constrictor. Experientia 1982;38:943 945.

  • 57.

    Coates EL, Ballam GO. Breathing and upper airway CO2 in reptiles: role of the nasal and vomeronasal systems. Am J Physiol 1989;256:R91 R97.

    • Search Google Scholar
    • Export Citation
  • 58.

    de Andrade DV, Tattersall GJ, Brito SP, et al. The ventilatory response to environmental hypercarbia in the South American rattlesnake, Crotalus durissus. J Comp Physiol B 2004;174:281 291.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 59.

    Hargreaves K, Dubner R, Brown F, et al. A new and sensitive method for measuring thermal nociception in cutaneous hyperalgesia. Pain 1988;32:77 88.

  • 60.

    Lim R, Zavou MJ, Milton PL, et al. Measuring respiratory function in mice using unrestrained whole-body plethysmography. J Vis Exp 2014;90:e51755.

    • Search Google Scholar
    • Export Citation
  • 61.

    Gratz RK. Ventilation and gas exchange in the diamondback water snake, Natrix rhombifera. J Comp Physiol B 1978;127:299 305.

  • 62.

    Stinner JN. Ventilation, gas exchange and blood gases in the snake, Pituophis melanoleucus. Respir Physiol 1982;47:279 298.

  • 63.

    Furilla RA, Bartlett D Jr. Intrapulmonary CO2 inhibits inspiration in garter snakes. Respir Physiol 1989;78:207 217.

  • 64.

    Enhorning G, van Schaik S, Lundgren C, et al. Whole-body plethysmography, does it measure tidal volume of small animals? Can J Physiol Pharmacol 1998;76:945 951.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 65.

    Milsom WK, Chatburn J, Zimmer MB. Pontine influences on respiratory control in ectothermic and heterothermic vertebrates. Respir Physiol Neurobiol 2004;143:263 280.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 66.

    Bartlett D Jr, Mortola JP, Doll EJ. Respiratory mechanics and control of the ventilatory cycle in the garter snake. Respir Physiol 1986;64:13 27.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 67.

    Malte CL, Malte H, Wang T. Episodic ventilation lowers the efficiency of pulmonary CO2 excretion. J Appl Physiol 2013;115:1506 1518.

  • 68.

    Hicks JW, Wood SC. Temperature regulation in lizards: effects of hypoxia. Am J Physiol 1985;248:R595 R600.

  • 69.

    Hicks JW, Wang T. Hypoxic hypometabolism in the anesthetized turtle, Trachemys scripta. Am J Physiol 1999;277:R18 R23.

  • 70.

    Heard DJ. Reptile anesthesia. Vet Clin North Am Exot Anim Pract 2001;4:83 117.

  • 71.

    Mosley CAE. Anesthesia and analgesia in reptiles. Semin Avian Exot Pet Med 2005;14:243 262.

  • 72.

    Smith DA, Barker IK, Allen OB. The effect of ambient temperature and type of wound on healing of cutaneous wounds in the common garter snake (Thamnophis sirtalis). Can J Vet Res 1988;52:120 128.

    • Search Google Scholar
    • Export Citation
  • 73.

    Nielsen B. On the regulation of the respiration in reptiles: the effect of temperature and CO2 on the respiration of lizards (Lacerta). J Exp Biol 1961;38:301 314.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 74.

    Glass M, Johansen K. Control of breathing in Acrochordus javanicus, an aquatic snake. Physiol Zool 1976;49:328 340.

  • 75.

    Gregoretti SM, Pleuvry BJ. Interactions between morphine and doxapram in the rabbit and mouse. Br J Anaesth 1977;49:323 329.

  • 76.

    Haji A, Kimura S, Ohi Y. Reversal of morphine-induced respiratory depression by doxapram in anesthetized rats. Eur J Pharmacol 2016;780:209 215.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 77.

    Golder FJ, Hewitt MM, McLeod JF. Respiratory stimulant drugs in the post-operative setting. Respir Physiol Neurobiol 2013;189:395 402.

  • 78.

    Gupta PK, Dundee JW. Morphine combined with doxapram or naloxone: a study of post-operative pain relief. Anaesthesia 1974;29:33 39.

  • 79.

    Gross PM, Marcus ML, Heistad DD. Regional distribution of cerebral blood flow during exercise in dogs. J Appl Physiol 1980;48:213 217.

  • 80.

    Miletich DJ, Ivankovich AD, Albrecht RF, et al. The effects of doxapram on cerebral blood flow and peripheral hemodynamics in the anesthetized and unanesthetized goat. Anesth Analg 1976;55:279 285.

    • Crossref
    • Search Google Scholar
    • Export Citation
  • 81.

    Brückner JB, Hess W, Schneider E, et al. Doxapram-induced changes in circulation and myocardial efficiency [in German]. Anaesthesist 1977;26:156 164.

    • Search Google Scholar
    • Export Citation
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