Editor’s Perspective
What We Already Know about This Topic
  • Many if not all drugs used in anesthesia and analgesia can produce potentially severe respiratory depression

  • Maintenance of breathing under anesthesia is linked to the drive exerted by the retrotrapezoid nucleus on the respiratory central pattern generator

  • The retrotrapezoid nucleus neurons that stimulate breathing during anesthesia are carbon dioxide–sensitive noncatecholaminergic neurons that express Phox2b, a master gene for the development of autonomic neurons

  • The conditional mouse model with the +7Ala repeat mutation targeted to the retrotrapezoid nucleus (Phox2b27Alaki/+ mice) present a massive selective loss of retrotrapezoid nucleus neurons and lack carbon dioxide chemosensitivity at birth but survive normally and partially recover carbon dioxide chemosensitivity in adulthood

What This Article Tells Us That Is New
  • Ketamine, propofol, and fentanyl caused lethal respiratory failure in most mice with selective genetic loss of retrotrapezoid nucleus neurons, at doses that were safe in their wild type littermates

Background

Optimal management of anesthesia-induced respiratory depression requires identification of the neural pathways that are most effective in maintaining breathing during anesthesia. Lesion studies point to the brainstem retrotrapezoid nucleus. We therefore examined the respiratory effects of common anesthetic/analgesic agents in mice with selective genetic loss of retrotrapezoid nucleus neurons (Phox2b27Alacki/+ mice, hereafter designated “mutants”).

Methods

All mice received intraperitoneal ketamine doses ranging from 100 mg/kg at postnatal day (P) 8 to 250 mg/kg at P60 to P62. Anesthesia effects in P8 and P14 to P16 mice were then analyzed by administering propofol (100 and 150 mg/kg at P8 and P14 to P16, respectively) and fentanyl at an anesthetic dose (1 mg/kg at P8 and P14 to P16).

Results

Most mutant mice died of respiratory arrest within 13 min of ketamine injection at P8 (12 of 13, 92% vs. 0 of 8, 0% wild type; Fisher exact test, P < 0.001) and P14 to P16 (32 of 42, 76% vs. 0 of 59, 0% wild type; P < 0.001). Cardiac activity continued after terminal apnea, and mortality was prevented by mechanical ventilation, supporting respiratory arrest as the cause of death in the mutants. Ketamine-induced mortality in mutants compared to wild types was confirmed at P29 to P31 (24 of 36, 67% vs. 9 of 45, 20%; P < 0.001) and P60 to P62 (8 of 19, 42% vs. 0 of 12, 0%; P = 0.011). Anesthesia-induced mortality in mutants compared to wild types was also observed with propofol at P8 (7 of 7, 100% vs. 0 of 17,7/7, 100% vs. 0/17, 0%; P < 0.001) and P14 to P16 (8 of 10, 80% vs. 0 of 10, 0%; P < 0.001) and with fentanyl at P8 (15 of 16, 94% vs. 0 of 13, 0%; P < 0.001) and P14 to P16 (5 of 7, 71% vs. 0 of 11, 0%; P = 0.002).

Conclusions

Ketamine, propofol, and fentanyl caused death by respiratory arrest in most mice with selective loss of retrotrapezoid nucleus neurons, in doses that were safe in their wild type littermates. The retrotrapezoid nucleus is critical to sustain breathing during deep anesthesia and may prove to be a pharmacologic target for this purpose.

Respiratory depression associated with sedation and anesthesia is a persistent and puzzling problem in anesthesiology.1–4  In the last decade, particular attention has been paid to the respiratory depressant effects of opioids, which are particularly strong compared to those of other agents. Nonetheless, many if not all drugs used for anesthesia and analgesia can produce potentially severe respiratory depression.5,6  Although we probably cannot fully prevent respiratory depression during anesthesia,7  active research is underway to identify respiratory stimulants capable of minimizing this effect without overly affecting anesthesia.4  Knowledge about the toxicity and efficacy of these drugs is still insufficient to allow clinical use.4  Identifying the cell types and neural pathways that sustain breathing during deep anesthesia may facilitate the development of effective respiratory stimulants.

Lesion studies suggest that breathing maintenance during anesthesia may involve the drive supplied by the retrotrapezoid nucleus to the respiratory central pattern generator.5,8,9  The retrotrapezoid nucleus is a small group of glutamatergic neurons located in the rostral ventrolateral medulla, whose activation by arterial carbon dioxide stimulates breathing.10  In anesthetized cats and rats, neuronal lesions produced in the retrotrapezoid nucleus region by the neurotoxin kainic acid or by electrolysis caused apnea.11–13  However, both these lesion-induction techniques lack selectivity, as recently pointed out by Souza et al.14  Studies of saporin-substance P injections showed that the retrotrapezoid nucleus neurons responsible for stimulating breathing during anesthesia were carbon dioxide–sensitive noncatecholaminergic neurons that expressed Phox2b,14  a master gene for the development of autonomic neurons.15  In anesthetized rats, 70% destruction of these retrotrapezoid-nucleus carbon dioxide–sensitive chemoreceptors substantially raised the carbon dioxide apneic threshold, thereby predisposing to apnea.16 

To look for further evidence that the retrotrapezoid nucleus is pivotal in breathing maintenance during anesthesia, we examined breathing patterns in anesthetized mice with selective genetic depletion of the retrotrapezoid nucleus neurons. Loss of retrotrapezoid nucleus neurons has been reported in mouse models of congenital central hypoventilation syndrome, a life-threatening disorder characterized by hypoventilation during sleep and absence of the ventilatory response to carbon dioxide.17,18  The disease-causing mutations in congenital central hypoventilation syndrome are generally polyalanine repeat expansion mutations of PHOX2B.19  Knock-in mice bearing the 7-alanine expanded allele (i.e., 27Ala) of PHOX2B (the most frequent mutation in patients19 ) exhibit massive loss of retrotrapezoid nucleus neurons, lack carbon dioxide chemosensitivity, and die within hours after birth.20  In contrast, in the conditional mouse model with the +7Ala repeat mutation targeted to the retrotrapezoid nucleus (Phox2b27Alaki/+ mice, hereafter designated “mutants”), despite massive selective loss of retrotrapezoid nucleus neurons and absence of carbon dioxide chemosensitivity at birth, survival is normal and carbon dioxide chemosensitivity recovers partially in adulthood.21  We hypothesized that mice with selective genetic depletion of retrotrapezoid nucleus neurons would be prone to severe respiratory failure during anesthesia.

Mice

All experimental protocols were approved by local and national ethics committees (Ministry of Higher Education and Scientific Research, Directorate-General for Research and Innovation, Paris, France), in accordance with the European Communities Council Directive 2010/63/EU for animal care. The experiments in the main study were conducted in mutant mice and their wild type littermates at postnatal day (P) 8, which approximately corresponds to term in humans,22  and P14 to P16, corresponding to infancy in humans.22  Complementary studies were conducted at P29 to P31, corresponding to adolescence,23  and at P60 to P62, corresponding to early adulthood. The mice that survived anesthesia were killed at P8 by decapitation and beyond P8 by cervical dislocation, without anesthesia. The experiments were conducted in mice of both sexes. The proportions of males and females at birth were not significantly different in the wild types (125 males and 97 females in total; male-to-female ratio, 1.29) and mutants (86 males and 87 females in total; male-to-female ratio, 0.99, Fisher exact test; P = 0.192).

We generated conditional, tissue-specific, knock-in Phox2b27Alacki/+ mice as previously described.21  Briefly, upon cre recombinase-mediated recombination, mouse Phox2b exon 3 was replaced by the mutated human exon 3 bearing the seven-residue expansion. Because human and mouse Phox2b protein sequences are identical, the encoded protein is identical to mouse Phox2b except for the extension of the polyalanine stretch. Offspring with the recombined locus were produced by crossing Egr2cre/+ males with Phox2b27Alacki/27Alacki females. Genotyping was performed with tail DNA after completion of the protocol and data processing. To detect the presence of the Phox2b27Alacki allele, the primers GCCCACAGTGCCTCTTAACT and CTCTTAAACGGGCGTCTCAC were used, yielding bands of 330 bp for the wild type gene, 474 bp for the mutated gene, and 380 bp for the recombined allele. To detect cre, the primers AAATTTGCCTGCATTACCG and ATGTTTAGCTGGCCCAAATG were used, yielding a band of 200 bp.

Treatments

In the main study, ketamine anesthesia was administered to mice at P8 (8 wild types, 13 mutants) and P14 to P16 (59 wild types, 42 mutants) and in complementary studies at P29 to P31 (45 wild types, 36 mutants) and P60 to P62 (12 wild types, 19 mutants). We chose ketamine as an anesthetic/analgesic agent with minimal effects on central respiratory drive.24  We also tested propofol and fentanyl at P8 (propofol: 17 wild types, 7 mutants; fentanyl: 13 wild types, 16 mutants) and P14 to P16 (propofol: 10 wild types, 10 mutants; fentanyl: 11 wild types, 7 mutants). Both propofol and fentanyl markedly depress respiration.24  Ketamine, propofol, and fentanyl are widely used for pediatric and adult anesthesia/sedation and act mainly by different mechanisms.5,25 

All agents were administered by intraperitoneal injection in the lower left quadrant of the abdomen, near the midline and umbilicus, taking care to avoid any visible milk spots. A 19-mm/30-gauge needle was inserted at an angle of 45° to the abdominal wall. The maximal volume administered was 20 µl/g. All injections were performed by the same experimenter.

The doses were as follows: ketamine, 100 mg/kg at P8, 150 mg/kg at P14 to P16, and 250 mg/kg at P29 to P31 and P60 to P62; propofol, 100 and 150 mg/kg at P8 and P14 to P16, respectively; and fentanyl, 1 mg/kg at P8 and P14 to P16. These doses were previously shown to rapidly produce deep anesthesia at all studied ages, as assessed by the loss of righting and tail-pinch responses,26  with no or minimal mortality in wild type mice. They were at the upper end of the intraperitoneal ketamine,27  propofol,28  and fentanyl29  dose ranges previously used in mice. The investigators were blinded to genotype, which was visually indiscernible. The genotypes were determined after completion of each protocol and therefore, cardiorespiratory data processing and mortality recordings were performed without previous knowledge of genotypes.

Pharmacologic Manipulation of Serotonin Neuronal Function

In a separate experiment, we analyzed the possible involvement of the serotonergic system (which is not affected by Phox2b mutation20 ) in the response to anesthesia in mutants and wild type littermates. In rodents, serotonin produces an excitatory effect on the pre-Bötzinger complex, mediated by 5-hydroxytryptamine 2 receptors.30  We tested serotonin system involvement by using the 5-hydroxytryptamine 2A/C antagonist ketanserin at P29 to P31 (10 mg/kg intraperitoneal, combined with 150 mg/kg ketamine in 25 wild types, 16 mutants, vs. ketamine alone, in 22 wild types, 7 mutants). The litters were alternately assigned to the ketanserin and control groups.

Plethysmography

Breathing variables were measured noninvasively by using a battery of four custom-made, whole-body flow barometric plethysmographs (fig. 1, A–D), as previously described.21  Data were collected in all P8 mice (38 wild types, 36 mutants) and a large subset of P14 to P16 mice (69 wild types, 47 mutants). Each plethysmograph was composed of two 100-ml Plexiglas chambers (fig. 1B) immersed in a thermoregulated water bath to maintain the temperature at 32°C to prevent anesthesia-induced hypothermia. A 200 ml · min-1 flow of dry air (Brooks airflow stabilizer, Urlo, The Netherlands) was injected in each chamber. The differential pressure between the chambers (GE Sensing transducer, France; range, ± 0.1 millibar) was converted into a digital signal at a sampling rate of 100 Hz and processed by using Labview software (National Instruments, USA). The apparatus was calibrated before each session by using a built-in pump incorporating a microsyringe (Ito Inc., Japan), which injected a sinusoidal airflow into the animal chamber with maximal amplitude 2 µl and frequency 6 Hz. We measured breath duration (s) and tidal volume (ml · g-1) on a breath-by-breath basis to calculate minute ventilation (tidal volume · breath duration-1 · 60, ml · min-1 · g-1). The limitations of the plethysmographic method in newborn mice have been discussed elsewhere.31  Briefly, plethysmography is effective for measuring breath duration (or frequency) and apneas but only provides semiquantitative measurements of tidal volume and minute ventilation because of gas-compression effects related to airway resistance.31  In particular, upper airway obstruction may affect tidal volume and ventilation values. By allowing simultaneous measurements in the same pup, and in four pups simultaneously, our experimental setup contributed to reduce experimental variability and sample size, thereby meeting the ethical requirements of reducing and refining animal use in research.

Fig. 1.

Experimental setup and protocol. (A) General view of the cardiorespiratory recording system for simultaneous and noninvasive measurements of ventilation and electrocardiography. Four pups were tested simultaneously. (B) Each pup was installed in the plethysmograph chamber in which it could move freely. Electrocardiography was recorded via four rectangular gold electrodes embedded in the floor of each chamber. (C) Breathing variables were derived from the difference in pressure (P) between the two thermoregulated chambers. (D and E) Examples of ventilatory and electrocardiography traces during air, hypercapnic challenge, and air again in a postnatal day 15 mice before anesthesia. (F) Graphical representation of the experimental protocol in postnatal day 8 and postnatal day 14 to postnatal day 16 mice. Each mouse was initially placed in the plethysmograph for a 15-min recording comprising an 8% carbon dioxide (CO2) challenge (Phase 1). Then, the mice were extracted from the chamber to receive an intraperitoneal injection of ketamine, propofol, or fentanyl. Within a 4-min delay, mice were re-placed in the plethysmograph for a 30-min physiologic recording (Phase 2) followed by a second 8% CO2 challenge (Phase 3, identical to Phase 1). Mortality was assessed after completion of Phase 3.

Fig. 1.

Experimental setup and protocol. (A) General view of the cardiorespiratory recording system for simultaneous and noninvasive measurements of ventilation and electrocardiography. Four pups were tested simultaneously. (B) Each pup was installed in the plethysmograph chamber in which it could move freely. Electrocardiography was recorded via four rectangular gold electrodes embedded in the floor of each chamber. (C) Breathing variables were derived from the difference in pressure (P) between the two thermoregulated chambers. (D and E) Examples of ventilatory and electrocardiography traces during air, hypercapnic challenge, and air again in a postnatal day 15 mice before anesthesia. (F) Graphical representation of the experimental protocol in postnatal day 8 and postnatal day 14 to postnatal day 16 mice. Each mouse was initially placed in the plethysmograph for a 15-min recording comprising an 8% carbon dioxide (CO2) challenge (Phase 1). Then, the mice were extracted from the chamber to receive an intraperitoneal injection of ketamine, propofol, or fentanyl. Within a 4-min delay, mice were re-placed in the plethysmograph for a 30-min physiologic recording (Phase 2) followed by a second 8% CO2 challenge (Phase 3, identical to Phase 1). Mortality was assessed after completion of Phase 3.

Close modal

Electrocardiography

The plethysmograph chambers were equipped with electrocardiography recording platforms composed of four rectangular gold electrodes insulated from one another and embedded in the floor of the chamber (fig. 1, B–E). Conduction was enhanced by using electrode hydrogel (Sekisui Plastics, Japan). Signals were digitized at a sampling rate of 1,000 Hz (16 bits, PCI-6229, National Instruments, USA). An electrocardiograph signal was obtained when at least three paws were in contact with three different electrodes or, occasionally, when the pup was lying on the floor.

Mechanical Ventilation

In a separate study, P14 to P16 mice anesthetized with 150 mg/kg ketamine (22 wild types, 7 mutants) were mechanically ventilated with air until the anesthesia wore off. The negative pressure ventilator comprised a set of chambers connected to a common negative-pressure generator allowing for simultaneous ventilation of up to 60 mice. Each mouse was placed for 2 h in a custom setting that allowed mechanical ventilation by intermittent extrathoracic negative pressure. The head of the animal was carefully placed in a thin latex ring, which was then affixed to a polycarbonate chamber so that the body was inside and the head outside. An intermittent negative-pressure generator alternately connected the chamber to the atmosphere or to a low-pressure vacuum source. Ventilation frequency was set at 3 Hz, duty cycle at 0.2, and peak negative pressure in the chamber at 14 cm H2O. This pressure value was chosen to generate respiratory movements whose amplitude and frequency approximately corresponded to those during spontaneous breathing in mice of the same strain and age, according to preliminary pneumotachography measurements (not shown).

Design

The study design is summarized in figure 1F. In Phase 1 (preanesthesia carbon dioxide test, 15 min), mice were randomly taken from each litter in successive groups of four, weighed, and placed together in the plethysmograph chambers. After 5 min of familiarization with the chamber, baseline breathing and heart rate were recorded over 5 min. Then, the airflow through the plethysmograph was switched to 8% CO2 + 21% O2 + 71% N2 at the same flow rate (200 ml/min per chamber) for 5 min, after which the flow was switched back to normoxia for 5 min. In Phase 2 (air breathing under anesthesia, 30 min), the four mice were extracted from the plethysmograph chambers, given anesthetics, and replaced simultaneously in the plethysmograph chambers within 4 min after the injection, for a 30-min baseline recording. Phase 3 (posttreatment carbon dioxide test, 15 min) was identical to Phase 1. The primary outcome of the study was mortality caused by anesthesia in the mutants compared to the wild types. Mortality was assessed immediately after completion of Phase 3, based on cardiorespiratory signal monitoring while the mice were in the plethysmograph and on immobility, unresponsiveness to pinch, and discoloration. Survivors were reunited with their dams. Adolescent and adult mice (P29 to P31 and P60 to P62) did not undergo plethysmography; they were anesthetized with ketamine, and mortality was then assessed for 60 min. All the experiments were performed during the day, between 9:00 am and 6:30 pm.

Statistical Analysis

Anesthesia-induced mortality was compared between mutant and wild type mice, at each age and with each anesthetic agent, by applying the two-tailed Fisher exact test. The experimental unit was the animal (n refers to the number of animals in each group). Weights and baseline cardiorespiratory variables were compared by two-tailed independent t tests. The time-course of the ventilatory response to hypercapnia was analyzed by averaging minute ventilation over consecutive 30-s periods throughout the 15-min plethysmographic recording (5 min air, 5 min hypercapnia, 5 min air) at P8 and P14 to P16. These data were compared between mutant and wild type mice using two-way repeated measures ANOVA with genotype (wild types vs. mutants) as a between-subject factor and time period (1 to 30) as a repeated factor. Bonferroni post hoc tests were performed to compare time-matched minute ventilation data between genotypes. For all parametric tests, normality of data was assessed by the Shapiro–Wilk test. Data are mean ± SD in the text, tables, and figures. No statistical power calculation was conducted before the study. The sample size was based on our previous experience with respiratory variability in this genetic model. All analyses were done using R Studio v1.1.243 and JMP Pro 13 (SAS Institute, Inc., USA). Differences were considered statistically significant if P < 0.05. Sex had no statistically significant effect in any analyses and will not be mentioned further.

Disruption of Carbon Dioxide Sensitivity in Mutant Mice

At P8 and P14 to P16, the mutants (n = 36 and n = 47, respectively) weighed slightly less than their wild type littermates (n = 38 and n = 69, respectively; table 1) but were normal in general appearance and behavior. First, we verified that the Phox2b27Alacki/+ mutation effectively disrupted the carbon dioxide–sensitive drive, as previously shown,21  by performing whole-body plethysmography at P8 and P14 to P16. Mutant and wild type mice at P8 did not differ significantly regarding baseline breathing variables (i.e., before anesthesia, Phase 1, table 1). At P14 to P16, mutant mice showed significantly longer breath durations (i.e., lower breathing frequencies) and marginally smaller ventilation values than wild type mice. As expected, at P8, mutants lacked any ventilatory response to carbon dioxide, in contrast to wild type mice (genotype-by-time interaction, P < 0.001; fig. 2A). A slight response was present at P14 to P16 but was significantly smaller than in the wild type mice (genotype-by-time interaction, P < 0.001; fig. 2B). This disruption of carbon dioxide sensitivity in mutant mice confirms previous results.21  Wild type mice sustained a regular breathing pattern and a vigorous response to hypercapnia (fig. 2, A and B).

Table 1.

Baseline Respiratory Variables for Wild Type and Phox2b27Alacki/+ Mutant Mice at P8 and P14 to P16

Baseline Respiratory Variables for Wild Type and Phox2b27Alacki/+ Mutant Mice at P8 and P14 to P16
Baseline Respiratory Variables for Wild Type and Phox2b27Alacki/+ Mutant Mice at P8 and P14 to P16
Fig. 2.

Ventilatory responses to 8% carbon dioxide (CO2) stimulus for 5 min (shaded area) in wild type (empty circles) and Phox2b27Alacki/+ (mutant) mice (filled circles) with selective loss of retrotrapezoid nucleus before anesthesia at postnatal day (P) 8 (n = 38 wild type and n = 36 mutant mice; A) and P14 to P16 (n = 69 wild type and n = 47 mutant mice; B). Breathing was measured by whole-body plethysmography. *P < 0.05, #P < 0.005, and †P < 0.0005 indicate significant differences between wild types and mutants. Data are mean ± SD.

Fig. 2.

Ventilatory responses to 8% carbon dioxide (CO2) stimulus for 5 min (shaded area) in wild type (empty circles) and Phox2b27Alacki/+ (mutant) mice (filled circles) with selective loss of retrotrapezoid nucleus before anesthesia at postnatal day (P) 8 (n = 38 wild type and n = 36 mutant mice; A) and P14 to P16 (n = 69 wild type and n = 47 mutant mice; B). Breathing was measured by whole-body plethysmography. *P < 0.05, #P < 0.005, and †P < 0.0005 indicate significant differences between wild types and mutants. Data are mean ± SD.

Close modal

Ketamine-induced Mortality in Mutant Mice

Ketamine anesthesia produced high mortality rates in mutant mice at all studied ages, including adulthood, whereas all or most wild type mice survived (table 2). The sedative effect extended beyond completion of Phase 3 in all surviving mice, including mutants.

Table 2.

Ketamine-induced Mortality Rates in Phox2b27Alacki/+ Mutant and Wild Type Mice

Ketamine-induced Mortality Rates in Phox2b27Alacki/+ Mutant and Wild Type Mice
Ketamine-induced Mortality Rates in Phox2b27Alacki/+ Mutant and Wild Type Mice

Ketamine-induced Mortality Was Due to Respiratory Arrest

The cause of death was first determined by analyzing the cardiorespiratory recordings. Figure 3 shows a typical sequence of respiratory events in a P14 to P16 mutant mouse given ketamine. A period of eupneic breathing (fig. 3A) was followed by a sequence of short clusters of gasps, in pairs (“double gasps”) or triplets (“triple gasps”), separated by apneas (fig. 3B). This sequence was followed by a series of gasps of decreasing amplitude ending with terminal apnea (fig. 3C), usually during Phase 2 (within 13 min after ketamine administration). However, the time to terminal apnea varied widely across individuals, with some mice dying before being placed in the plethysmograph (i.e., within 4 min after ketamine administration). The electrocardiograph signal consistently outlasted the terminal gasps, although the heart rate was reduced (fig. 4). These findings support respiratory arrest as the cause of death, as opposed to cardiac arrest.

Fig. 3.

Respiratory arrest in a Phox2b27Alacki/+ mutant mouse at postnatal day 15 after ketamine administration. Breathing pattern, initially normal at the onset of recording (A), was followed after 3- to 4-min recording by apneas interspersed with clusters of gasps (B), and after 16-min recording by a series of gasps gradually diminishing in size until terminal apnea (C).

Fig. 3.

Respiratory arrest in a Phox2b27Alacki/+ mutant mouse at postnatal day 15 after ketamine administration. Breathing pattern, initially normal at the onset of recording (A), was followed after 3- to 4-min recording by apneas interspersed with clusters of gasps (B), and after 16-min recording by a series of gasps gradually diminishing in size until terminal apnea (C).

Close modal
Fig. 4.

Illustrative cardiorespiratory traces in a Phox2b27Alacki/+ mutant mouse at postnatal day (P) 8 exposed to ketamine. (A) The end of the series of gasps gradually diminishing in size followed by terminal apnea. (B) Cardiac activity outlasted the terminal apnea, showing that death was caused by respiratory arrest. P14 to P16 mice presented similar patterns of lethal respiratory arrests. ECG, electrocardiography.

Fig. 4.

Illustrative cardiorespiratory traces in a Phox2b27Alacki/+ mutant mouse at postnatal day (P) 8 exposed to ketamine. (A) The end of the series of gasps gradually diminishing in size followed by terminal apnea. (B) Cardiac activity outlasted the terminal apnea, showing that death was caused by respiratory arrest. P14 to P16 mice presented similar patterns of lethal respiratory arrests. ECG, electrocardiography.

Close modal

That death was due to respiratory arrest in ketamine-exposed mutants was further confirmed by exposing a separate group of P14 to P16 mutants (n = 7) and wild type littermates (n = 22) to mechanical ventilation immediately after a 150-mg/kg ketamine injection. The mice were ventilated until the effects of the anesthesia wore off, as reflected by gross body movements. The end of anesthesia varied widely among mice, from 60 min to 90 min after ketamine injection. In contrast to the high mortality rate caused by 150-mg/kg ketamine in nonventilated mutants (32 of 42, 76%; table 2), all ventilated mutants survived anesthesia (exact Fisher test, ventilated vs. nonventilated mutants, P < 0.001), as did all wild type mice, confirming respiratory arrest as the cause of ketamine-induced death in nonventilated mutant mice. Monitoring for 3 days after anesthesia confirmed that all ventilated mice survived and appeared normal, ruling out any major delayed effect of ketamine anesthesia.

To further analyze the respiratory effects of ketamine in mutants, we first compared breathing variables in mutants and their wild type littermates throughout the recording session (see Supplemental Digital Content 1, https://links.lww.com/ALN/B908, showing breathing variables in wild types and mutants from Phase 1 to Phase 3, and distinguishing surviving and nonsurviving mutants in Phase 1). Before ketamine treatment, the mutants had no ventilatory response to carbon dioxide. In both genotypes, ketamine depressed minute ventilation, mainly by decreasing tidal volume and increasing breath duration. However, the wild types tended to recover pre-ketamine levels for all variables, whereas ventilation in the mutant survivors remained depressed throughout Phases 2 and 3. The inspiratory-to-expiratory ratio was also markedly affected by ketamine in mutant survivors, suggesting impaired respiratory phase timing. Then, we analyzed individual plots of nonsurviving mutants (Supplemental Digital Content 2, https://links.lww.com/ALN/B909, which is a zoom of Phase 2, including wild types, mutant survivors, and mutant nonsurvivors with available respiratory data until the onset of gasping). No clear differences in respiratory variables were observed between surviving and nonsurviving mutants.

Effect of the Serotonin System on Ketamine-induced Mortality

We then examined whether survival of a subgroup of ketamine-exposed mutants was due to the serotonergic drive to the respiratory central pattern generator. Serotonergic terminals are found throughout brainstem respiratory regions, and both serotonin and the peptides released by serotonergic cells modify the activity of many types of respiratory neurons.8  In a separate experiment, we addressed the contribution of the serotonergic drive (which is not affected by Phox2b gene mutation20 ) by combining 10 mg/kg ketanserin, a serotonin 5-hydroxytryptamine 2A/C antagonist, with 150 mg/kg ketamine, administered to P29 to P31 mice (25 wild types, 16 mutants), and comparing the findings to those in a group given only ketamine (22 wild types, 7 mutants).

The 150 mg/kg ketamine dose was nonlethal in P29 to P31 mutants (table 3). However, when combined with ketanserin, ketamine caused death in 11 of 16 (69%) mutants (exact Fisher test, ketanserin-treated vs. untreated mutants, P = 0.005). All but one ketanserin-treated wild type mouse survived (table 3). These results support a role for the serotonin system in protecting mutants against ketamine-induced respiratory arrest. However, our attempt to decrease mortality in ketamine-exposed mutants by using the 5-hydroxytryptamine 2 agonist 1-(2,5-dimethoxy-4-iodophenyl)-2-aminopropane in doses of 0.75 to 1.5 mg/kg did not yield statistically significant results (0.75 mg/kg: P > 0.999; 1 mg/kg: P = 0.121; 1.5 mg/kg: P = 0.184).

Table 3.

Effect of Ketanserin on Ketamine-induced Mortality in Postweaning Phox2b27Alacki/+ Mutant Mice

Effect of Ketanserin on Ketamine-induced Mortality in Postweaning Phox2b27Alacki/+ Mutant Mice
Effect of Ketanserin on Ketamine-induced Mortality in Postweaning Phox2b27Alacki/+ Mutant Mice

Extrapolation to Propofol and Fentanyl

We then examined whether the lethal effects of ketamine anesthesia in mutants extended to other anesthetic/analgesic agents with different mechanisms of action. We exposed mutants and their wild type littermates to propofol (P8: 17 wild types, 7 mutants, 100 mg/kg; and P14 to P16: 10 wild types, 10 mutants, 150 mg/kg) or fentanyl (P8: 13 wild types, 16 mutants, 1 mg/kg; and P14 to P16, 11 wild types, 7 mutants, 1 mg/kg; table 3). Both agents produced very high mortality rates in mutants but no mortality in wild type mice (table 4), further supporting the mutants’ vulnerability to anesthesia previously suggested by ketamine-induced mortality. The respiratory traces in propofol-exposed mice displayed similar patterns of gasps and apneas as did those seen with ketamine (see Supplemental Digital Content 3, https://links.lww.com/ALN/B910, showing respiratory arrest in a P15 mutant mouse after propofol administration). Fentanyl-exposed mice exhibited severely disrupted ventilation from the onset of the recording, also followed by a sequence of gasps that gradually diminished in size until terminal apnea (see Supplemental Digital Content 4, https://links.lww.com/ALN/B911, showing respiratory arrest in a P15 mutant mouse after fentanyl administration).

Table 4.

Propofol and Fentanyl-induced Mortality in Phox2b27Alacki/+ Mutant Mice

Propofol and Fentanyl-induced Mortality in Phox2b27Alacki/+ Mutant Mice
Propofol and Fentanyl-induced Mortality in Phox2b27Alacki/+ Mutant Mice

In this study, ketamine, propofol, and fentanyl caused respiratory arrest in most mice with selective genetic loss of retrotrapezoid nucleus neurons, in doses that were safe in their wild type littermates. These results support a pivotal role for the retrotrapezoid nucleus in maintaining spontaneous breathing during deep anesthesia. The serotonin system probably contributed to sustain breathing during anesthesia in retrotrapezoid nucleus neuron-depleted mice.

Central Respiratory Failure

That the high mortality in the ketamine-exposed mutants was due to respiratory arrest was first established by examining the order of respiratory and cardiac arrests and was subsequently confirmed by showing that the mutants survived if given mechanical ventilation. In mutant mice, ketamine anesthesia produced apneas interspersed with clusters of gasps, a common marker of hypoxia. Gasps (and sighs) are generated by the pre-Bötzinger complex,32  which does not express Phox2b and therefore is spared by Phox2b gene mutations.33  Clusters of double or triple gasps have never been reported in ketamine-anesthetized animals; there is a single report in piglets sedated with 20 mg/kg intraarticular pentobarbital sodium administration.34  However, this pattern is often reported in dying preterm infants and in infants who subsequently experienced sudden infant death syndrome.35  Thus, in both these human cases and our mutant mice, the lack of successful autoresuscitation was not related to an inability to gasp.35,36  Rather, the anesthetized mutants probably died of hypoxic hypoventilation caused by disruption of the excitatory drive sent by the retrotrapezoid nucleus to the central pattern generator.

The recurrent apnea in anesthetized mutants suggested that absence of the retrotrapezoid nucleus may have caused an increase in the carbon dioxide apneic threshold. In an earlier study, the carbon dioxide apneic threshold was increased by 70% destruction of carbon dioxide–sensitive chemoreceptors in the retrotrapezoid nucleus of anesthetized rats.16  Our mutants exhibited similar or higher loss rates.21  Furthermore, ketamine, propofol, and mu-opioid agonists also increase the carbon dioxide apneic threshold, in a dose-dependent manner.25,37  Breathing was not impaired by anesthesia alone (as indicated by the breathing pattern in anesthetized wild types) or neuronal loss alone (as indicated by the breathing pattern in non-anesthetized mutants). However, when combined in anesthetized mutants, these two factors caused respiratory arrest, probably by increasing the carbon dioxide apneic threshold.

Interindividual Variability

Few mutant mice survived ketamine, propofol, or fentanyl anesthesia, and mortality was highest in the youngest animals. Individual differences in the vulnerability to anesthesia possibly reflected variations in retrotrapezoid nucleus cell loss caused by the Phox2b27Alacki/+ mutation.21  In previous studies, the number of residual cells in the retrotrapezoid nucleus of mutant mice was estimated at 15 to 20% of the mean value in wild types.21  This range is close to the threshold at which pharmacologic destruction of the retrotrapezoid nucleus disrupted phrenic nerve discharge in rats.16  In anesthetized and ventilated rats, bilateral 70% destruction of retrotrapezoid nucleus carbon dioxide–sensitive chemoreceptors (i.e., 30% spared neurons) substantially raised the carbon dioxide apneic threshold.16  Also, in rats with unilateral 70% retrotrapezoid nucleus neuron destruction, acute inhibition of the contralateral intact retrotrapezoid nucleus with muscimol instantly abolished phrenic nerve discharge.16  Cell counting was not done in the current study, but the mutant mice that survived anesthesia may have been those with more numerous spared cells in the retrotrapezoid nucleus and, therefore, with a stronger residual excitatory drive to the central pattern generator.14 

Serotonergic Contribution to Breathing

The key role of the retrotrapezoid nucleus in sustaining breathing during anesthesia does not preclude contributions of other neuronal systems. In particular, activation of the serotonergic raphe neurons stimulates ventilation, and the full effects of carbon dioxide on breathing require activity of these neurons.10  Therefore, we examined whether the serotonin system contributed to sustain breathing during anesthesia in mutants. We found that the 5-hydroxytryptamine 2 receptor antagonist ketanserin caused death in mutants despite being used in doses that were safe in wild types when administered alone. This finding supports a role in breathing maintenance during anesthesia of the excitatory drive from serotoninergic neurons to the respiratory network (especially the pre-Bötzinger complex), via 5-hydroxytryptamine 2 receptors.

This analysis, however, has several limitations. First, the 5-hydroxytryptamine 2 receptor agonist 2,5-dimethoxy-4-iodoamphetamine failed to significantly reduce ketamine-induced mortality in P14 to P16 mutants. Second, the role of the serotonin system revealed by ketanserin may reflect a compensatory adaptation to retrotrapezoid nucleus loss specific to mutant mice. Third, a previous report that knock-out mice with disrupted 5-hydroxytryptamine 2A receptor signaling survived anesthesia with ketamine 150 mg/kg38  would seem at variance with our findings. We focused on the 5-hydroxytryptamine 2 receptor because of its excitatory effect on respiratory rhythm generation, but the effects of 5-hydroxytryptamine 1,4 receptor agonists, which counteract opioid-induced ventilatory depression,1,4,39  also deserve attention. Finally, we cannot discount that ketanserin increased ketamine-induced respiratory depression and mortality via its alpha1-adrenergic receptor blocking properties in residual retrotrapezoid nucleus cells40  or in the pre-Bötzinger complex.41  This possible effect might explain why the 5-hydroxytryptamine 2 agonist 1-(2,5-dimethoxy-4-iodophenyl)-2-aminopropane did not decrease ketamine-induced mortality. The role for serotonin pathways in sustaining breathing during anesthesia clearly requires further investigation.

Mono- versus Polysynaptic Inputs

The critical role of the retrotrapezoid nucleus during deep anesthesia probably reflects its strategic location and function in the central pattern generator. One important characteristic of retrotrapezoid nucleus neurons is their monosynaptic input to the respiratory central pattern generator,42  which contrasts with the chiefly polysynaptic inputs from other sources (e.g., the peripheral chemosensory drive, arousal-related suprapontine activation, and exercise-related central command). Polysynaptic pathways are strongly affected by anesthetics due to cumulation of effects along the signaling chain.43  Furthermore, the pontine parabrachial-Koölliker-Fuse complex, which controls expiratory duration and the inspiratory on-switch,44,45  has strong reciprocal connections with the retrotrapezoid nucleus.46  The loss of retrotrapezoid input in mutants may have contributed to the disruption of respiratory phase timing in this group. Therefore, the cumulative effect of anesthetic agents on synaptic transmission, combined with retrotrapezoid neuron depletion, may account for the vulnerability of mutants to all anesthetics studied, despite their different cellular targets and properties.

The most parsimonious explanation of anesthesia-induced respiratory arrest in mutant mice is that massive loss of retrotrapezoid nucleus drive combined with silencing of polysynaptic respiratory pathways fully abolished the respiratory drive to the central pattern generator. However, our data do not rule out the possibility that respiratory arrest in the mutants involved mechanisms related to the specific cellular targets and respiratory central pattern generator effects of each anesthetic.5,25  Ketamine is an N-methyl-d-aspartate receptor antagonist that also depresses ventilation via γ-aminobutyric acid type A and opioid receptors.47  Propofol depresses ventilation via the γ-aminobutyric acid type A receptor. The glutamatergic pre-Boötzinger complex and the pontine parabrachial-Koölliker-Fuse complex, which, as noted above, control expiratory duration and the inspiratory on-switch,44,45  express opioid receptors. Possibly, activation of one of these pathways may be sufficient to disrupt the function of a system made inherently unstable by retrotrapezoid neuronal loss.

Finally, in vitro studies recently showed that isoflurane exposure aggravated the toxic effects of the mutated PHOX2B gene (i.e., PHOX2B protein misfolding, aggregation, and loss of nuclear localization) in cultured cells with the +7Ala expansion.48  Further research is needed to determine how these effects may affect the function of residual retrotrapezoid nucleus neurons in mutant mice.

The current results may explain why patients with congenital central hypoventilation syndrome, most of whom carry PHOX2B polyalanine expansions, are prone to major anesthesia-related complications that require profound perioperative precautions.49  They also suggest that the often used combination of opioids and propofol, which act additively on anesthesia induction,50  may be particularly detrimental to patients with congenital central hypoventilation syndrome.

Conclusions

Mice with selective loss of retrotrapezoid nucleus neurons were highly vulnerable to deep anesthesia induced by ketamine, propofol, or fentanyl. Most of them died of respiratory arrest when anesthetized, in contrast to their wild type littermates. These results confirm the pivotal role of the retrotrapezoid nucleus in breathing maintenance during deep anesthesia. They suggest that drugs capable of selectively activating retrotrapezoid nucleus neurons may hold promise for preventing anesthesia-induced respiratory depression.

Support was provided solely from institutional sources, Association Française du Syndrome d’Ondine (French Association of Ondine Syndrome; Paris, France) and Legs Poix (Poix Legacy Fund, Chancellery of Paris Universities; Paris, France).

The authors declare no competing interests.

1.
Ren
J
,
Ding
X
,
Greer
JJ
:
5-HT1A receptor agonist Befiradol reduces fentanyl-induced respiratory depression, analgesia, and sedation in rats.
Anesthesiology
2015
;
122
:
424
34
.
2.
Linz
K
,
Schröder
W
,
Frosch
S
,
Christoph
T
:
Opioid-type respiratory depressant side effects of cebranopadol in rats are limited by its nociceptin/orphanin FQ peptide receptor agonist activity.
Anesthesiology
2017
;
126
:
708
15
.
3.
van der Schier
R
,
Roozekrans
M
,
van Velzen
M
,
Dahan
A
,
Niesters
M
:
Opioid-induced respiratory depression: Reversal by non-opioid drugs.
F1000Prime Rep
2014
;
6
:
79
.
4.
Dahan
A
,
van der Schrier
R
,
Smith
T
,
Aarts
L
,
van Velzen
M
,
Niesters
M
:
Averting opioid-induced respiratory depression without affecting analgesia.
Anesthesiology
2018
;
128
:
1027
37
.
5.
Teppema
LJ
,
Baby
S
:
Anesthetics and control of breathing.
Respir Physiol Neurobiol
2011
;
177
:
80
92
.
6.
Dahan
A
,
Roozekrans
M
,
van der Schrier
R
,
Smith
T
,
Aarts
L
:
Primum non nocere or how to resolve drug-induced respiratory depression.
Anesthesiology
2013
;
118
:
1261
3
.
7.
Dahan
A
,
Teppema
LJ
:
Influence of anaesthesia and analgesia on the control of breathing.
Br J Anaesth
2003
;
91
:
40
9
.
8.
Guyenet
PG
:
The 2008 Carl Ludwig Lecture: Retrotrapezoid nucleus, CO2 homeostasis, and breathing automaticity.
J Appl Physiol (1985)
2008
;
105
:
404
16
.
9.
Smith
JC
,
Abdala
AP
,
Borgmann
A
,
Rybak
IA
,
Paton
JF
:
Brainstem respiratory networks: Building blocks and microcircuits.
Trends Neurosci
2013
;
36
:
152
62
.
10.
Guyenet
PG
,
Bayliss
DA
:
Neural control of breathing and CO2 homeostasis.
Neuron
2015
;
87
:
946
61
.
11.
Nattie
EE
,
Li
AH
:
Fluorescence location of RVLM kainate microinjections that alter the control of breathing.
J Appl Physiol (1985)
1990
;
68
:
1157
66
.
12.
Nattie
EE
,
Li
AH
,
St John
WM
:
Lesions in retrotrapezoid nucleus decrease ventilatory output in anesthetized or decerebrate cats.
J Appl Physiol (1985)
1991
;
71
:
1364
75
.
13.
Nattie
EE
,
Li
A
:
Retrotrapezoid nucleus lesions decrease phrenic activity and CO2 sensitivity in rats.
Respir Physiol
1994
;
97
:
63
77
.
14.
Souza
GMPR
,
Kanbar
R
,
Stornetta
DS
,
Abbott
SBG
,
Stornetta
RL
,
Guyenet
PG
:
Breathing regulation and blood gas homeostasis after near complete lesions of the retrotrapezoid nucleus in adult rats.
J Physiol
2018
;
596
:
2521
45
.
15.
Pattyn
A
,
Morin
X
,
Cremer
H
,
Goridis
C
,
Brunet
JF
:
The homeobox gene Phox2b is essential for the development of autonomic neural crest derivatives.
Nature
1999
;
399
:
366
70
.
16.
Takakura
AC
,
Moreira
TS
,
Stornetta
RL
,
West
GH
,
Gwilt
JM
,
Guyenet
PG
:
Selective lesion of retrotrapezoid Phox2b-expressing neurons raises the apnoeic threshold in rats.
J Physiol
2008
;
586
:
2975
91
.
17.
Weese-Mayer
DE
,
Berry-Kravis
EM
,
Ceccherini
I
,
Keens
TG
,
Loghmanee
DA
,
Trang
H
;
ATS Congenital Central Hypoventilation Syndrome Subcommittee
:
An official ATS clinical policy statement: Congenital central hypoventilation syndrome: Genetic basis, diagnosis, and management.
Am J Respir Crit Care Med
2010
;
181
:
626
44
.
18.
Gallego
J
:
Genetic diseases: Congenital central hypoventilation, Rett, and Prader-Willi syndromes.
Compr Physiol
2012
;
2
:
2255
79
.
19.
Amiel
J
,
Laudier
B
,
Attié-Bitach
T
,
Trang
H
,
de Pontual
L
,
Gener
B
,
Trochet
D
,
Etchevers
H
,
Ray
P
,
Simonneau
M
,
Vekemans
M
,
Munnich
A
,
Gaultier
C
,
Lyonnet
S
:
Polyalanine expansion and frameshift mutations of the paired-like homeobox gene PHOX2B in congenital central hypoventilation syndrome.
Nat Genet
2003
;
33
:
459
61
.
20.
Dubreuil
V
,
Ramanantsoa
N
,
Trochet
D
,
Vaubourg
V
,
Amiel
J
,
Gallego
J
,
Brunet
JF
,
Goridis
C
:
A human mutation in Phox2b causes lack of CO2 chemosensitivity, fatal central apnea, and specific loss of parafacial neurons.
Proc Natl Acad Sci USA
2008
;
105
:
1067
72
.
21.
Ramanantsoa
N
,
Hirsch
MR
,
Thoby-Brisson
M
,
Dubreuil
V
,
Bouvier
J
,
Ruffault
PL
,
Matrot
B
,
Fortin
G
,
Brunet
JF
,
Gallego
J
,
Goridis
C
:
Breathing without CO(2) chemosensitivity in conditional Phox2b mutants.
J Neurosci
2011
;
31
:
12880
8
.
22.
Semple
BD
,
Blomgren
K
,
Gimlin
K
,
Ferriero
DM
,
Noble-Haeusslein
LJ
:
Brain development in rodents and humans: Identifying benchmarks of maturation and vulnerability to injury across species.
Prog Neurobiol
2013
;
106-107
:
1
16
.
23.
Brust
V
,
Schindler
PM
,
Lewejohann
L
:
Lifetime development of behavioural phenotype in the house mouse (Mus musculus).
Front Zool
2015
;
12
(
suppl 1
):
S17
.
24.
Dahan
A
:
General anesthesia and control of respiration.
Semin Anesth
1996
;
15
:
328
34
.
25.
Stuth
EA
,
Stucke
AG
,
Zuperku
EJ
:
Effects of anesthetics, sedatives, and opioids on ventilatory control.
Compr Physiol
2012
;
2
:
2281
367
.
26.
Drobac
E
,
Durand
E
,
Laudenbach
V
,
Mantz
J
,
Gallego
J
:
A simple method for short-term controlled anesthesia in newborn mice.
Physiol Behav
2004
;
82
:
279
83
.
27.
Zuurbier
CJ
,
Koeman
A
,
Janssen
BJ
:
Letter to the editor: Ketamine-only versus isoflurane effects on murine cardiac function: Comparison at similar depths of anesthesia?
Am J Physiol Heart Circ Physiol
2015
;
309
:
H2160
.
28.
Kubo
K
,
Nishikawa
K
,
Hardy-Yamada
M
,
Ishizeki
J
,
Yanagawa
Y
,
Saito
S
:
Altered responses to propofol, but not ketamine, in mice deficient in the 65-kilodalton isoform of glutamate decarboxylase.
J Pharmacol Exp Ther
2009
;
329
:
592
9
.
29.
Kendall
LV
,
Hansen
RJ
,
Dorsey
K
,
Kang
S
,
Lunghofer
PJ
,
Gustafson
DL
:
Pharmacokinetics of sustained-release analgesics in mice.
J Am Assoc Lab Anim Sci
2014
;
53
:
478
84
.
30.
Peña-Ortega
F
:
Tonic neuromodulation of the inspiratory rhythm generator.
Front Physiol
2012
;
3
:
253
.
31.
Ren
J
,
Lenal
F
,
Yang
M
,
Ding
X
,
Greer
JJ
:
Coadministration of the AMPAKINE CX717 with propofol reduces respiratory depression and fatal apneas.
Anesthesiology
2013
;
118
:
1437
45
.
32.
Lieske
SP
,
Thoby-Brisson
M
,
Telgkamp
P
,
Ramirez
JM
:
Reconfiguration of the neural network controlling multiple breathing patterns: Eupnea, sighs and gasps [see comment].
Nat Neurosci
2000
;
3
:
600
7
.
33.
Pattyn
A
,
Morin
X
,
Cremer
H
,
Goridis
C
,
Brunet
JF
:
Expression and interactions of the two closely related homeobox genes Phox2a and Phox2b during neurogenesis.
Development
1997
;
124
:
4065
75
.
34.
Sanocka
UM
,
Donnelly
DF
,
Haddad
GG
:
Autoresuscitation: A survival mechanism in piglets.
J Appl Physiol (1985)
1992
;
73
:
749
53
.
35.
Sridhar
R
,
Thach
BT
,
Kelly
DH
,
Henslee
JA
:
Characterization of successful and failed autoresuscitation in human infants, including those dying of SIDS.
Pediatr Pulmonol
2003
;
36
:
113
22
.
36.
Tomori
Z
,
Donic
V
,
Benacka
R
,
Gresova
S
,
Peregrim
I
,
Kundrik
M
,
Pallayova
M
,
Jakus
J
:
Reversal of functional disorders by aspiration, expiration, and cough reflexes and their voluntary counterparts.
Front Physiol
2012
;
3
:
467
.
37.
Dahan
A
,
Nieuwenhuijs
D
,
Olofsen
E
:
Influence of propofol on the control of breathing
in
Advances in Modelling and Clinical Application of Intravenous Anaesthesia
. Edited by
Vuyk
J
,
Schraag
S
.
Boston, MA
,
Springer
,
2003
, pp
81
94
38.
Morici
JF
,
Ciccia
L
,
Malleret
G
,
Gingrich
JA
,
Bekinschtein
P
,
Weisstaub
NV
:
Serotonin 2a receptor and serotonin 1a receptor interact within the medial prefrontal cortex during recognition memory in mice.
Front Pharmacol
2015
;
6
:
298
.
39.
Czick
ME
,
Waldman
JC
,
Gross
JB
:
Sources of inspiration: A neurophysiologic framework for understanding anesthetic effects on ventilatory control.
Curr Anesthesiol Rep
2014
;
4
:
67
75
.
40.
Kuo
FS
,
Falquetto
B
,
Chen
D
,
Oliveira
LM
,
Takakura
AC
,
Mulkey
DK
:
In vitro characterization of noradrenergic modulation of chemosensitive neurons in the retrotrapezoid nucleus.
J Neurophysiol
2016
;
116
:
1024
35
.
41.
Viemari
JC
,
Ramirez
JM
:
Norepinephrine differentially modulates different types of respiratory pacemaker and nonpacemaker neurons.
J Neurophysiol
2006
;
95
:
2070
82
.
42.
Bochorishvili
G
,
Stornetta
RL
,
Coates
MB
,
Guyenet
PG
:
Pre-Bötzinger complex receives glutamatergic innervation from galaninergic and other retrotrapezoid nucleus neurons.
J Comp Neurol
2012
;
520
:
1047
61
.
43.
Banoub
M
,
Tetzlaff
JE
,
Schubert
A
:
Pharmacologic and physiologic influences affecting sensory evoked potentials: Implications for perioperative monitoring.
Anesthesiology
2003
;
99
:
716
37
.
44.
Zuperku
EJ
,
Stucke
AG
,
Hopp
FA
,
Stuth
EA
:
Characteristics of breathing rate control mediated by a subregion within the pontine parabrachial complex.
J Neurophysiol
2017
;
117
:
1030
42
.
45.
Zuperku
EJ
,
Stucke
AG
,
Krolikowski
JG
,
Tomlinson
J
,
Hopp
FA
,
Stuth
EA
:
Inputs to medullary respiratory neurons from a pontine subregion that controls breathing frequency.
Respir Physiol Neurobiol
2018
.
[Epub ahead of print]
.
46.
Rosin
DL
,
Chang
DA
,
Guyenet
PG
:
Afferent and efferent connections of the rat retrotrapezoid nucleus.
J Comp Neurol
2006
;
499
:
64
89
.
47.
Sarton
E
,
Teppema
LJ
,
Olievier
C
,
Nieuwenhuijs
D
,
Matthes
HW
,
Kieffer
BL
,
Dahan
A
:
The involvement of the mu-opioid receptor in ketamine-induced respiratory depression and antinociception.
Anesth Analg
2001
;
93
:
1495
500, table of contents
.
48.
Coghlan
M
,
Richards
E
,
Shaik
S
,
Rossi
P
,
Vanama
RB
,
Ahmadi
S
,
Petroz
C
,
Crawford
M
,
Maynes
JT
:
Inhalational anesthetics induce neuronal protein aggregation and affect ER trafficking.
Sci Rep
2018
;
8
:
5275
.
49.
Basu
SM
,
Chung
FF
,
AbdelHakim
SF
,
Wong
J
:
Anesthetic considerations for patients with congenital central hypoventilation syndrome: A systematic review of the literature.
Anesth Analg
2017
;
124
:
169
78
.
50.
Ben-Shlomo
I
,
Finger
J
,
Bar-Av
E
,
Perl
AZ
,
Etchin
A
,
Tverskoy
M
:
Propofol and fentanyl act additively for induction of anaesthesia.
Anaesthesia
1993
;
48
:
111
3
.