The superior cervical ganglia modulate ventilatory responses to hypoxia independently of preganglionic drive from the cervical sympathetic chain

Abstract

Superior cervical ganglia (SCG) postganglionic neurons receive preganglionic drive via the cervical sympathetic chains (CSC). The SCG projects to structures like the carotid bodies (e.g., vasculature, chemosensitive glomus cells), upper airway (e.g., tongue, nasopharynx), and to the parenchyma and cerebral arteries throughout the brain. We previously reported that a hypoxic gas challenge elicited an array of ventilatory responses in sham-operated (SHAM) freely moving adult male C57BL6 mice and that responses were altered in mice with bilateral transection of the cervical sympathetic chain (CSCX). Since the CSC provides preganglionic innervation to the SCG, we presumed that mice with superior cervical ganglionectomy (SCGX) would respond similarly to hypoxic gas challenge as CSCX mice. However, while SCGX mice had altered responses during hypoxic gas challenge that occurred in CSCX mice (e.g., more rapid occurrence of changes in frequency of breathing and minute ventilation), SCGX mice displayed numerous responses to hypoxic gas challenge that CSCX mice did not, including reduced total increases in frequency of breathing, minute ventilation, inspiratory and expiratory drives, peak inspiratory and expiratory flows, and appearance of noneupneic breaths. In conclusion, hypoxic gas challenge may directly activate subpopulations of SCG cells, including subpopulations of postganglionic neurons and small intensely fluorescent (SIF) cells, independently of CSC drive, and that SCG drive to these structures dampens the initial occurrence of the hypoxic ventilatory response, while promoting the overall magnitude of the response. The multiple effects of SCGX may be due to loss of innervation to peripheral and central structures with differential roles in breathing control.

NEW & NOTEWORTHY We present data showing that the ventilatory responses elicited by a hypoxic gas challenge in male C57BL6 mice with bilateral superior cervical ganglionectomy are not equivalent to those reported for mice with bilateral transection of the cervical sympathetic chain. These data suggest that hypoxic gas challenge may directly activate subpopulations of superior cervical ganglia (SCG) cells, including small intensely fluorescent (SIF) cells and/or principal SCG neurons, independently of preganglionic cervical sympathetic chain drive.

INTRODUCTION

The cell bodies of postganglionic sympathetic neurons (1–5) and small intensely fluorescent (SIF) cells (6–9) within the left and right superior cervical ganglia (SCG) receive their preganglionic innervation from thoracic spinal cord (T1–T4) nerves that course within the ipsilateral cervical sympathetic chains (CSC). The majority of postganglionic nerves project from the SCG via two main trunks, the external and internal carotid nerves (10–13). In addition, a population of postganglionic SCG fibers in the external carotid nerve branch into the ganglioglomerular nerve (GGN). These GGN fibers innervate structures in the carotid bodies such as, the vasculature, chemoreceptor afferent nerve terminals, and glomus (type I) cells (8–11, 14–19). These fibers also innervate the terminals of baroreceptor afferent nerves in the carotid sinus (20–24). The above SCG projections, in addition to those directed to nuclei within the brainstem, such as the nucleus tractus solitarius (NTS) and hypothalamus (21–34) as well as the tongue and upper airway (35–41), support the possibility that the SCG is vital to the integration of cardiorespiratory function.

Until recently, no in vivo studies have characterized the effects of transection of the CSC or surgical removal of the SCG on breathing or the ventilatory responses that occur during hypoxic gas exposures. We recently reported that the responses of several breathing parameters that occurred during a 5-min challenge with a hypoxic gas mixture (10% O₂-90% N₂) were modified in C57BL6 mice that had undergone bilateral CSC transection (CSCX) 4 days previously (42). First, breathing parameters at rest in CSCX mice were similar to those of mice that underwent sham operation (SHAM). However, we found that the hypoxic gas challenge (hypoxic challenge)-induced increases in frequency of breathing (Freq) and accompanying decreases in inspiratory time (Ti), expiratory time (Te), end expiratory pause (EEP; pause between end of expiration and start of inspiration), and relaxation time (i.e., decay of respiration to 36% of maximum peak inspiratory flow), as well as increases in minute ventilation (MV), expiratory drive, and expiratory flow at 50% exhaled tidal volume (EF₅₀) occurred faster (i.e., reached peak values more quickly) in CSCX mice than in SHAM mice, although the total responses that occurred during the hypoxic challenge were similar in CSCX and SHAM mice. In addition, the total increase in tidal volume (TV) during the latter half of the hypoxic challenge was higher in CSCX mice than SHAM mice, whereas the early and total increases in peak inspiratory flow (PIF) were higher in the CSCX mice. These findings suggest that in C57BL6 mice, the CSC innervation of the SCG drives the SCG cells to provide inhibitory control to central structures (e.g., nuclei within the brainstem) and/or the carotid bodies that mediate the ventilatory responses to hypoxic challenge. The reasons for choosing male C57BL6 mice for this study in which we bilaterally removed the SCG (SCGX) and our companion CSCX study (42), including descriptions of genetic, morphological, neurophysiological and neurochemical factors that participate in the expression of the ventilatory responses of C57BL6 mice to hypoxic challenge are described in detail by Getsy et al (42). The particular problems raised by the use of these C57BL6 mice were also discussed in detail (42), and it is important to note that these mice (both males and females) present with substantial ventilatory instability compared with other mouse strains. This can be viewed as problematic but also as an opportunity to further our understanding of the genetic and gender aspects of ventilatory performance.

As mentioned, the SCG contains postganglionic sympathetic cell bodies that receive preganglionic (cholinergic) innervation from nerve fibers in the CSC (10–13) and SIF cells that also receive preganglionic innervation from the CSC (6–9). Evidence that neural activity in the CSC and ganglioglomerular nerve increases during hypoxia exposure (43–45) suggests that hypoxic challenge elicits preganglionic nerve-induced activation of postganglionic SCG neurons and SIF cells. As such, a carotid sinus (chemoreceptor afferent) nerve-brainstem-descending spinal cord pathway may be activated during hypoxic challenge, which in turn increases the activity of preganglionic CSC nerves projecting to SCG cells that in turn project to key ventilatory structures, such as the carotid body via the ganglioglomerular nerve branch. Thus, it can be hypothesized that mice with bilateral superior cervical ganglionectomy (SCGX) would present with similar responses to hypoxic challenge as those with bilateral CSCX. It is important to note that while it appears that principal SCG cells are not hypoxia-sensitive (46–51), there is convincing evidence that SIF cells/interneurons in the SCG directly respond to hypoxic challenges (52–55). Indeed, Dinger et al (53) clearly demonstrated that SIF cells in the SCG are selectively activated by hypoxic stimuli and that the neurochemical responses of these cells are comparable to those observed with chemosensory type I (glomus) cells of the carotid body.

The objectives of the present study were to use the whole body plethysmography method (42, 56–60) to 1) compare resting ventilation in adult male C57BL6 mice in which both SCG were surgically removed 4 days previously to those that were sham-operated (SHAM), and 2) compare the ventilatory responses of these mice to the hypoxic challenge. The data show that SCGX mice displayed numerous altered responses to hypoxic challenge that also occurred in CSCX mice but also displayed numerous changes in hypoxic challenge responses that CSCX mice did not, including diminished total increases in minute ventilation. As such, the major finding of this study is that bilateral removal of the SCG does not elicit the same effects as CSC transection (42) in response to hypoxic challenge, which suggests that hypoxia exerts multiple changes in neural signaling in the CSC-SCG pathway. These data raise the possibility that hypoxic challenge directly activates subpopulations of SCG cells, including SIF cells and/or subpopulations of postganglionic neurons independently of projections from the CSC, and that SCG innervation of cardiorespiratory structures is of vital importance to hypoxic ventilatory responses.

MATERIALS AND METHODS

Permissions

All of the studies described in the manuscript were performed in strict accordance with the National Institutes of Health (NIH) Guide for the Care and Use of Laboratory Animals (NIH Publication No. 80-23) revised in 1996. In addition, we obtained prior approval from the Animal Care and Use Committee of Case Western Reserve University for all protocols.

Mice

Male C57BL6 mice were obtained from Jackson Laboratories (Bar Harbor, ME). The mice were delivered pathogen-free 48 h before surgery and were housed in a pathogen-free room on a 12:12-h light-dark cycle in our Animal Resource Center. On the days of surgery, the mice were randomized into SHAM (n = 14) and SCGX (n = 14) groups. No mouse had difficulty surviving the SHAM or SCGX surgeries. For postsurgical recovery, the mice were returned to the quarantine room until the whole body plethysmography studies were performed. The mice had ad libitum access to water and standard rat chow before and after surgery. After plethysmography studies were performed, the mice were taken to the post mortem room in the Animal Resource Center and were euthanized via overexposure to CO₂ and then cervical dislocation. All mice used in the present study were completely different from those used in the study of Getsy et al (42).

SHAM and SCGX Surgeries

Adult C57BL6 mice (95 ± 1 days of age at time of surgery) were anesthetized via an intraperitoneal injection of ketamine (80 mg/kg, Ketaset, Zoetis, Parsippany, NJ, containing 100 mg/mL of solvent) and xylazine (10 mg/kg, Akorn Animal Health, Lake Forest, IL, containing 20 mg/mL solvent) and placed on a surgical table. The body temperatures of the mice were maintained at 37°C via a heating pad (SurgiSuite, Kent Scientific Corporation, Torrington, CT). Toe-pinch was regularly performed to assess the adequacy of anesthesia. A midline incision in the neck was made to expose the left and right CSC and SCG that were located behind each carotid artery bifurcation (Fig. 1). Both SCGs were surgically removed after transection of the CSC at its junction with the SCG and the points where the internal and external carotid nerves exit the SCG. The left and right CSC and SCG were identified but not transected in the SHAM mice. The incision in the neck was closed using nonabsorbable monofilament sutures. None of the mice showed signs of inflammation or pain from the surgery and began moving about the cages and eating and drinking ∼1 h after surgery. SCGX mice (but not SHAM mice) displayed Horner’s Syndrome (constriction of the pupil due to loss of postganglionic drive from the SCG) after they woke up from the anesthesia. All mice were given an injection of the nonsteroidal anti-inflammatory drug carprofen (5 mg/kg ip: Rimadyl, Zoetis, Parsippany, NJ, containing 50 mg/mL of solvent), 24 h and 48 h postsurgery to reduce possible pain or inflammation at the incision site. Mice were weighed daily to ensure proper weight gain. Both SCGX and SHAM mice were given a total of 4 days to recover from the surgical procedures before whole body plethysmography experiments were performed. This 4-day recovery time point was chosen because tyrosine hydroxylase-positive nerve terminals within rat carotid bodies were substantially diminished 3–4 days after the surgical removal of the SCG ipsilateral to the carotid body (14, 61–63). We have determined that these two injections of carprofen do not affect resting ventilation or the response to hypoxic gas challenge on day 4 postsurgery [e.g., peak MV response in naïve mice (n = 6) and carprofen-injected mice (n = 5) were 282 ± 23 and 275 ± 22 mL/min, respectively; P > 0.05]. An additional note is that since sutures were to be removed 10 days postsurgery, the mice still had sutures in at the time of the whole body plethysmography experiments.

Figure 1.

Picture of the left superior cervical ganglion (SCG) of a C57BL6 mouse and the associated cervical sympathetic chain (CSC), left internal carotid nerve (ICN), and external carotid nerve (ECN) nerves emanating from the SCG. The common carotid artery (CCA) and vagus (Xth cranial) nerve are shown. The scale bar in the photograph represents a length of 100 μm.

Whole Body Plethysmography

The methods for performing the whole body plethysmography in unrestrained freely moving mice have been detailed previously (42, 56–60). In brief, each mouse was put into a whole body plethysmograph (Buxco Small Animal Whole Body Plethysmography, DataSciences, Inc., St. Paul, MN) to continuously record breathing. Flow rates of the room air and hypoxic gas mixture (10% O₂-90% N₂) were set at 0.5 L/min. Chamber temperature (°C) and chamber relative humidity (%) were constantly monitored via sensors built into the plethysmography chambers, and the breathing waveforms were continuously recorded by system algorithms to allow the FinePointe software to accurately analyze recorded waveforms (https://www.datasci.com/products/buxco-respiratory-products/finepointe-whole-body-plethysmography). Note that the temperature of the air in the chambers housing the SHAM or SCGX mice changed minimally during each stage of the protocol (∼0.2°C), whereas the relative percent humidity of the chambers during the hypoxic challenge (relatively dry air) fell by −8.0 ± 0.6 (P < 0.05) and −6.7 ± 0.6 (P < 0.05) in the SHAM and SCGX mice, respectively (P > 0.05 for between-group response). The choice of parameters was based on the value of each of these parameters to understanding ventilatory control processes (42, 56–60, 64–66) in order to allow detailed analyses of the effects of SCGX on breathing patterns and the responses to hypoxic challenge. The parameters include Freq; TV; MV; Ti; Te; Ti/Te; Ti/(Ti + Te); EIP; end expiratory pause (EEP; pause between end of expiration and start of inspiration); PIF; peak expiratory flow (PEF); PIF/PEF; EF₅₀; relative rate of achieving PEF (Rpef); inspiratory drive (TV/Ti); expiratory drive (TV/Te); noneupneic breathing index (i.e., %noneupneic breaths including apneas during each epoch); noneupneic breathing index corrected for respiratory frequency (noneupneic breathing index/Freq); and degree of apneic pause during each recorded epoch (Te/relaxation time)-1. A full description of the methods by which the plethysmography software accepted or rejected breaths is provided by Getsy et al (42).

Protocols for the Recording of Ventilatory Responses during and following Hypoxic Challenge

On the particular day of study, each SCGX or SHAM mouse was placed in an individual plexiglass plethysmography chamber and given at least 60 min to acclimatize to the chamber. Once the mouse had settled and stable ventilatory parameters were obtained, the animal was exposed to a hypoxic challenge (10% O₂-90% N₂) for 5 min and then reexposed to room air for 15 min. The hypoxic challenge was given for 5 min because a 5 min time frame is associated with pronounced changes in ventilatory parameters that reached plateau levels before introduction of room air and also because the return to room air is associated with its own distinct ventilatory pattern.

Statistics

With respect to data collection, a data point was recorded every 15 s, before the hypoxic challenge (i.e., the 5-min period immediately before hypoxic challenge, referred to as prevalues), during the hypoxic challenge (the entire 5-min challenge) and after the hypoxic challenge (the entire 15 min of room air following the hypoxic challenge). To calculate the total responses that occurred during the hypoxic challenge and upon return to room air for the parameters of each of the mice, the values recorded during the hypoxic challenge phase and during the room-air phase were summed and expressed as the total cumulative %change from prevalues. The cumulative response was then determined for each mouse using the formulas, 1) total hypoxic challenge response = (sum of 20 values during hypoxic challenge) − (mean of the prevalues × 20); and 2) total room-air response = (sum of 60 values during room-air phase) − (mean of the pre values × 60). The means ± SE of the group data were then calculated. There were 20 values (i.e., data points) for each mouse recorded during the hypoxic challenge (four 15-s data points per min × 5-min recording period = 20 data points). Similarly, 60 values were collected for the post-hypoxic (room air) phase for each mouse (four 15-s data points per min × 15-min recording period = 60 data points). We also determined the total responses during the 0- to 105-s epoch and 106- to 300-s epoch of the hypoxic challenge, in addition to the entire 5-min hypoxic challenge (0- to 300-s epoch). Note that the 106- to 300-s epoch during the hypoxic challenge was when the differences between SHAM and SCGX mice were most evident. This epoch in the SHAM and SCGX mice had noneupneic breathing index values that were equivalent to prelevels and as such differences in ventilatory parameters between the groups were not complicated by data sampling issues.

All summary data are presented as means ± SE. The data were analyzed by one-way or two-way ANOVA followed by Student’s modified t test with Bonferroni corrections for multiple comparisons between means using the error mean square terms from each ANOVA (67–70). A value of P < 0.05 denoted the initial level of statistical significance that was modified according to the number of comparisons between means as detailed by Wallenstein et al (68). The modified t statistic is t = (mean of group 1 − mean of group 2)/[s² × (1/n₁ + 1/n₂)^1/2], where s² = the mean square within groups term from the ANOVA (the square root of this value is taken for the modified t statistic formula) and n₁ and n₂ are the number of mice in each group that is being compared. Based on an elementary inequality called Bonferroni’s inequality, a conservative critical value for the modified t statistics taken from tables of t distribution using a significance level of P/m, where m is the number of comparisons between groups to be performed. The degrees of freedom are those for the mean square for within group variation from the ANOVA table. In most cases, the critical Bonferroni value cannot be obtained from conventional tables of the t distribution but may be approximated from widely available tables of the normal curve by t* = z + (z + z3)/4n, with n being the degrees of freedom and z being the critical normal curve value for P/m (67–70). Wallenstein et al (68) first reported that the Bonferroni procedure is preferable for general use since it is easiest to apply, has the widest range of applications, and gives critical values that are lower than those of other procedures if the investigator can limit the number of comparisons and that will be only slightly larger than those of other procedures if many comparisons are made. The practical application of the Bonferroni procedure demonstrated by Wallenstein et al (68) was confirmed and expanded on in separate studies by Ludbrook (69) and by McHugh (70). A value of P < 0.05 was taken as the initial level of statistical significance (67, 68).

RESULTS

Resting Parameters

The ages and body weights of the mice and their baseline (resting) ventilatory parameters before exposure to hypoxic challenge are summarized in Table 1. There were 14 mice in each group. There were no differences in the ages or body weights of the two groups of mice (P > 0.05, for each comparison), and therefore, it was not necessary to correct for body weights for the volume parameters (i.e., TV, PIF, and PEF). No between-group differences were found for any variable (P > 0.05, for all comparisons) except for the greater degree of apneic pauses occurring in SCGX mice (P < 0.05). Described below (see Fig. 3, A, D, and G; Figs. 4, 5, 6, and 7, A and D; and Figs. 8, 9, and 10, A, D, and G) are the values recorded before, during the hypoxic challenge (10% O₂-90% N₂ for 5 min), and following reintroduction of room air. The arithmetic changes recorded over the first 60 s of hypoxic challenge are displayed below (see Fig. 3, B, E, and H; Figs. 4, 5, 6, and 7, B and E; and Figs. 8, 9, and 10, B, E, and H). The total response (%change from prevalues) during three epochs of the 300-s (5 min) hypoxic challenge, 0–105 s, 106–300 s, and 0–300 s, are displayed below (see Fig. 3, C, F, and I; Figs. 4, 5, 6, and 7, C and F; and Figs. 8, 9, and 10, C, F, and I). These epochs best represented the clearest differences between the SCGX and SHAM mice.

Table 1.

Baseline parameters in sham-operated mice and in mice with bilateral removal of the superior cervical ganglia

Parameter	SHAM	SCGX
Number of mice	14	14
Age, days	99 ± 1	98 ± 1
Body weight, g	27.6 ± 0.5	28.1 ± 0.5
Frequency (Freq), breaths/min	194 ± 5	190 ± 4
Tidal volume (TV), ml	0.151 ± 0.004	0.150 ± 0.006
Minute ventilation, ml/min	28.7 ± 0.7	28.2 ± 0.8
Inspiratory time (Ti), s	0.114 ± 0.002	0.115 ± 0.002
Expiratory time (Te), s	0.212 ± 0.008	0.216 ± 0.005
End inspiratory pause (EIP), ms	2.89 ± 0.09	2.92 ± 0.07
End expiratory pause (EEP), ms	47.5 ± 11.6	42.9 ± 7.7
Ti/Te	0.555 ± 0.018	0.542 ± 0.012
Ti/(Ti + Te)	0.354 ± 0.008	0.350 ± 0.005
Peak inspiratory flow (PIF), ml/s	2.34 ± 0.05	2.31 ± 0.04
Peak inspiratory flow (PIF), ml/s	1.58 ± 0.06	1.60 ± 0.09
PIF/PEF	1.51 ± 0.06	1.50 ± 0.07
Expiratory flow at 50% exhaled TV (EF50), ml/s	0.072 ± 0.002	0.073 ± 0.002
Relaxation time, s	0.099 ± 0.003	0.093 ± 0.002
Rate of achieving PEF (Rpef)	0.141 ± 0.009	0.127 ± 0.009
Inspiratory drive (TV/Ti), ml/s	1.34 ± 0.03	1.32 ± 0.03
Expiratory drive (TV/Te) ml/s	0.73 ± 0.03	0.71 ± 0.03
Noneupneic breathing index (NEBI), %	15.0 ± 2.0	14.1 ± 1.6
(NEBI/frequency) × 100	8.1 ± 1.2	7.5 ± 0.9
Apneic pause	1.15 ± 0.06	1.33 ± 0.04*

Values are means ± SE. *P < 0.05, superior cervical ganglionectomy (SCGX) vs. sham operated (SHAM).

Figure 3.

A, D, and G: frequency of breathing (Freq), tidal volume (TV), and minute ventilation (MV) values recorded before, during, and following a 5-min hypoxic (HX; 10% O₂-90% N₂) gas challenge in sham-operated (SHAM) mice and mice with bilateral superior cervical ganglionectomy (SCGX). B, E, and H: responses recorded during the first 60 s of hypoxic challenge in SHAM and SCGX mice (expressed as %prevalues). C, F, and I: total responses recorded during the first 105 s, between 106 and 300 s, and between 0 and 300 s (expressed as the sum of all %changes from prevalues). All data are presented as means ± SE. *P < 0.05, significant change from prevalues. †P < 0.05, SCGX mice vs. SHAM mice. There were 14 mice in each of the SHAM and SCGX groups.

Figure 4.

A and D: inspiratory time (Ti) and expiratory time (Te) values recorded before, during, and following a 5-min hypoxic (HX; 10% O₂-90% N₂) gas challenge in sham-operated (SHAM) mice and mice with bilateral superior cervical ganglionectomy (SCGX). B and E: responses of inspiratory times and expiratory times recorded during the first minute of hypoxic challenge in SHAM and SCGX mice (expressed as %prevalues). C and F: total responses recorded during the first 105 s, between 106 and 300 s, and between 0 and 300 s (expressed as the sum of all %changes from prevalues). All results are presented as means ± SE. *P < 0.05, significant from prevalues. †P < 0.05, SCGX vs. SHAM. There were 14 mice in each of the SHAM and SCGX groups.

Figure 5.

A and D: inspiratory time/expiratory time (Ti/Te) ratios and Ti/(Ti + Te) ratios recorded before, during, and following a 5-min hypoxic (HX; 10% O₂-90% N₂) gas challenge in sham-operated (SHAM) mice and mice with bilateral superior cervical ganglionectomy (SCGX). B and E: responses recorded during the first minute of hypoxic challenge in the SHAM and SCGX mice (expressed as %prevalues). C and F: total responses recorded during the first 105 s, between 106 and 300 s, and between 0 and 300 s (expressed as the sum of all %changes from prevalues). All results are presented as means ± SE. *P < 0.05, significant from prevalues. There were 14 mice in each of the SHAM and SCGX groups.

Figure 6.

A and D: end inspiratory pause (EIP) and end expiratory pause (EEP) values recorded before, during, and following a 5-min hypoxic (HX; 10% O₂-90% N₂) gas challenge in sham-operated (SHAM) mice and mice with bilateral superior cervical ganglionectomy (SCGX). B and E: responses recorded during the first minute of hypoxic challenge in the SHAM and SCGX mice (expressed as %prevalues). C and F: total responses recorded during the first 105 s, between 106 and 300 s, and between 0 and 300 s (expressed as the sum of all %changes from prevalues). All results are presented as means ± SE. *P < 0.05, significant change from prevalues. †P < 0.05, SCGX vs. SHAM. There were 14 mice in each of the SHAM and SCGX groups.

Figure 7.

A and D: inspiratory drive (TV/Ti) and expiratory drive (TV/Te) values recorded before, during, and following a 5-min hypoxic (HX; 10% O₂-90% N₂) gas challenge in sham-operated (SHAM) mice and mice with bilateral superior cervical ganglionectomy (SCGX). B and E: Responses recorded during the first minute of hypoxic challenge in the SHAM and SCGX mice (expressed as %prevalues). C and F: total responses recorded during the first 105 s, between 106 and 300 s, and between 0 and 300 s (expressed as the sum of all %changes from prevalues). All results are presented as means ± SE. *P < 0.05, significant change from prevalues. †P < 0.05, SCGX vs. SHAM. There were 14 mice in each of the SHAM and SCGX groups.

Figure 8.

A, D, and G: peak inspiratory flow (PIF) and peak expiratory flow (PEF) values and PIF/PEF ratios recorded before, during, and following a 5-min hypoxic (HX; 10% O₂-90% N₂) gas challenge in sham-operated (SHAM) mice and mice with bilateral superior cervical ganglionectomy (SCGX). B, E, and H: responses recorded during the first minute of hypoxic challenge in SHAM and SCGX mice (expressed as %prevalues). C, F, and I: total responses recorded during the first 105 s, between 106 and 300 s, and between 0 and 300 s (expressed as the sum of all %changes from prevalues). All results are presented as means ± SE. *P < 0.05, significant change from prevalues. †P < 0.05, SCGX vs. SHAM. There were 14 mice in each of the SHAM and SCGX groups.

Figure 9.

A, D, and G: EF₅₀, Rpef and relaxation time (RT) values recorded before, during, and following a 5-min hypoxic (HX; 10% O₂-90% N₂) gas challenge in sham-operated (SHAM) mice and mice with bilateral superior cervical ganglionectomy (SCGX). B, E, and H: responses recorded during the first minute of hypoxic challenge in SHAM and SCGX mice (expressed as %prevalues). C, F, and I: total responses recorded during the first 105 s, between 106 and 300 s, and between 0 and 300 s (expressed as the sum of all %changes from prevalues). All results are presented as means ± SE. *P < 0.05, significant change from prevalues. †P < 0.05, SCGX vs. SHAM. There were 14 mice in each of the SHAM and SCGX groups.

Figure 10.

A, D, and G: noneupneic breathing index (NEBI) values, NEBI/frequency (Freq) ratios, and apneic pause (AP) values recorded before, during and following a 5-min hypoxic (HX; 10% O₂-90% N₂) gas challenge in sham-operated (SHAM) mice and in mice with bilateral superior cervical ganglionectomy (SCGX). B, E, and H: responses recorded during the first minute of hypoxic challenge in SHAM and SCGX mice (expressed as %prevalues). C, F, and I: total responses recorded during the first 105 s, between 106 and 300 s and between 0 and 300 s (expressed as the sum of all %changes from prevalues). All results are presented as means ± SE. *P < 0.05, significant change from prevalues. †P < 0.05, SCGX vs. SHAM. There were 14 mice in each of the SHAM and SCGX groups.

Before introduction of the hypoxic challenge, the mice were calm and still although they would occasionally groom or move about the chamber. Upon introduction of the hypoxic challenge, the mice would not change behavior (i.e., there was no obvious direct response of the mice when switching to the hypoxic gas), and they became progressively more still as the hypoxic challenge progressed, as described previously (42, 56–60). The mice returned to normal levels of behaviors such as grooming upon reintroduction of room air and accordingly, the long-lasting increase in noneupneic breathing index upon return to room air was not explained by an increase in behaviors such as locomotion, rearing, grooming, or sniffing (42, 56–60).

Hypoxic Challenge and Return to Room-Air Phases: Frequency, Tidal Volume, Minute Ventilation

Selected examples of the ventilatory waveforms recorded in a SHAM and SCGX mouse during each phase of the protocol are displayed in Fig. 2. A bar denoting 1.25 s can be seen between Fig. 2, A and B. The baseline Freq (prehypoxic challenge) value was 192 breaths/min in the SHAM mouse and 216 breaths/min in the SCGX mouse. The initial response to hypoxic challenge (at 15 s) was very similar in size in the SHAM mouse (192 to 240 breaths/min = 48 breaths/min, +25%) and SCGX mouse (216 to 264 breaths/min = 48 breaths/min, +22%). The decline in Freq (roll-off) that occurred at the end of the 5-min hypoxic challenge was much more pronounced in the SCGX mouse than in the SHAM mouse. The value (hypoxic challenge at 5 min) in the SHAM mouse was equivalent to the prevalue (192 to 192 breaths/min = 0 breaths/min, +0%) whereas the value in the SCGX mouse at 5 min hypoxic challenge was considerably lower than the prevalue (216 to 168 breaths/min = −48 breaths/min, –22%). The initial increase in Freq upon return to room air (RA at 15 s) was more pronounced in the SCGX mouse (216 to 504 breaths/min = 288 breaths/min, +133%) than in the SHAM mouse (192 to 216 breaths/min = 24 breaths/min, +13%). Freq values 5 min following return to room air (RA at 5 min) were similar in the SHAM mouse (192 to 168 breaths/min = −24 breaths/min, −13%) and SCGX mouse (216 to 192 breaths/min = −24 breaths/min, −11%).

Figure 2.

Selected sections of the respiratory waveforms that were recorded during key times of the protocol in a sham-operated (A; SHAM) mouse and in a mouse with bilateral superior cervical ganglionectomy (B; SCGX). A 1.25-s time bar is displayed between A and B. HXC, phase of challenge with hypoxic gas; RA, the room-air phase.

The Freq, TV, and MV responses in SHAM and SCGX mice recorded during the 5 min hypoxic challenge and then return to room air are shown in Fig. 3. Figure 3A shows that hypoxic challenge elicited a prompt rise in Freq in SHAM and SCGX mice that decayed substantially (i.e., displayed roll-off) toward the end of the hypoxic challenge. Figure 3B shows that the initial rise in Freq in SCGX mice was similar to that of SHAM mice except that the response was higher in SCGX mice at 45 s. Figure 3C shows the total increases in Freq in the 0- to 105-s, 106- to 300-s, and 0- to 300-s epochs in the SHAM and SCGX mice. As can be seen, the faster roll-off evident in the SCGX mice in Fig. 3A resulted in SCGX mice displaying significantly lower total increases in Freq responses during the 106- to 300-s epoch. The return to room air elicited immediate and pronounced increases in Freq that were similar in SHAM and SCGX mice (Fig. 3A). As summarized in Table 2, both groups displayed similar total Freq responses with return to room air if all of the mice were used in the analyses. The relatively large standard error term in the SHAM mice prompted us to examine the data more closely, and we found that the SHAM mice separate into a larger subgroup of 10 that had a net positive change in Freq over the 15 min room-air recording period (+Freq) whereas 4 SHAM mice had a net negative response (−Freq). There were 13 mice that fell into the net positive responses in the SCGX mice and only 1 that fell into the net negative category. The total Freq responses with return to room air in the SCGX mice that fell into the net positive category were substantially smaller than those in the net positive category of SHAM mice (Table 2).

Table 2.

Total changes that occurred during the first 15 min upon return to room air

Parameter	SHAM	SCGX
Number of mice	14	14
Frequency, %	+41.1 ± 10.8 (14)	+30.9 ± 5.5 (14)
+Freq, %	+58.8. ± 8.9 (10)	+33.5 ± 5.0* (13)
−Freq, %	−3.3 ± 0.5 (4)	−2.9 (1)
Tidal volume (TV), %	+39.2 ± 8.0 (14)	+28.0 ± 6.3 (14)
+TV, %	+43.4 ± 7.1 (13)	+33.3 ± 5.6 (12)
−TV, %	−15.8 (1)	−2.9 ± 0.1 (2)
Minute ventilation, %	+113 ± 26.9	+79.7 ± 14.1 (14)
+MV, %	+123.2 ± 25.9 (13)	+79.7 ± 14.1 (14)
−MV, %	−20 (1)	None
Inspiratory time (Ti), %	−30.4 ± 5.3 (14)	−30.3 ± 3.1 (14)
Expiratory time (Te), %	−4.1 ± 6.0 (14)	+8.6 ± 4.9 (14)
+Te, %	+16.5 ± 2.2 (6)	+17.0 ± 3.7 (10)
−Te, %	−19.5 ± 4.2 (8)	−12.3 ± 2.3 (4)
Ti/Te, %	−25.9 ± 2.3 (14)	−33.3 ± 1.9* (14)
Ti/(Ti + Te), %	−19.0 ± 1.7 (14)	−25.1 ± 1.5* (14)
End inspiratory pause (EIP), %	−8.3 ± 2.7 (14)	−12.1 ± 2.5 (14)
End expiratory pause (EEP), %	+185 ± 55 (14)	+259 ± 66 (14)
Inspiratory drive (TV/Ti), %	+143 ± 29 (14)	+116 ± 16 (14)
Expiratory drive (TV/Te), %	+77.8 ± 21.4 (14)	+44.6 ± 11.8 (14)
+TV/Te, %	+100.9 ± 19.9 (11)	+53.2 ± 11.2* (12)
−TV/Te, %	−7.2 ± 3.6 (3)	−6.9 ± 1.8 (2)
Peak expiratory flow (PEF), %	+122 ± 18 (14)	+83 ± 16 (14)
PIF/PEF, %	+11.1 ± 5.6 (14)	+25.8 ± 6.3 (14)
+PIF/PEF, %	+18.2 ± 4.7 (11)	+34.6 ± 4.8* (11)
−PIF/PEF, %	−15.0 ± 1.6 (3)	−6.6 ± 1.5 (3)
Expiratory flow at 50% exhaled TV (EF50), %	+117 ± 27 (14)	+80 ± 17 (14)
Relaxation Time (RT), %	−5.3 ± 5.5 (14)	+15.0 ± 4.6* (14)
+RT, %	+12.1 ± 3.3 (8)	+24.5 ± 2.1* (10)
−RT, %	−16.0 ± 3.9 (6)	−8.9 ± 2.1 (4)
Rate of achieving PEF (Rpef), %	+14.5 ± 10.7 (14)	+19.6 ± 13.9 (14)
+Rpef, %	+40.4 ± 6.8 (8)	+69.9 ± 9.6* (6)
−Rpef, %	−20.1 ± 3.3 (6)	−18.0 ± 4.8 (8)
Noneupneic breathing index (NEBI), %	+315 ± 84 (14)	+280 ± 52 (14)
(NEBI/frequency) x 100, %	+175 ± 41 (14)	+194 ± 37 (14)
Apneic pause (Te/RT)-1, %	+5.3 ± 4.8 (14)	−4.9 ± 6.1 (14)
+(Te/RT) - 1, %	+19.1 ± 3.7 (7)	+25.9 ± 4.1 (4)
−(Te/RT) - 1, %	−8.6 ± 1.9 (7)	−17.2 ± 2.3* (10)

Values are means ± SE; (n), number of mice in the subgroups for Te, relaxation time (RT), relative rate of achieving peak expiratory flow (Rpef), and (Te/RT)-1. *P < 0.05, superior cervical ganglionectomy (SCGX) vs. sham operated (SHAM).

As summarized in Fig. 3, D–F, hypoxic challenge elicited prompt and sustained increases in TV that were very similar in the SHAM and SCGX mice, except that the initial rise in TV happened more quickly in SCGX mice, and this rise in TV was significantly higher than in the SHAM mice at 30 s (Fig. 3E). The return to room air elicited a minor rise in TV after which TV gradually declined toward prehypoxic challenge values in SHAM and SCGX mice (Fig. 3D). As summarized in Table 2, both groups had similar total responses in TV with return to room air. Figure 3G shows that hypoxic challenge elicited an initial increase in MV in SHAM and SCGX mice that displayed a marked degree of roll-off. Figure 3H shows that the initial MV responses in the SCGX mice happened somewhat more quickly than in the SHAM mice and that this augmentation in MV was significantly higher in SCGX than SHAM mice at 45 s. Figure 3I shows that the exaggerated roll-off in the SCGX mice resulted in a significantly lower total increase in MV during epoch 106–300 s. As seen in Fig. 3G, there were substantial increases in MV in the SHAM mice and SCGX mice following the return to room air. As summarized in Table 2, the total MV responses following return to room air in the SHAM and SCGX groups were similar to one another.

Hypoxic Challenge and Return to Room-Air Phases: Inspiratory and Expiratory Times

Figure 4 summarizes the Ti and Te responses in SHAM and SCGX mice during the 5-min hypoxic challenge and following reintroduction of room air. As displayed in Fig. 4, A–C, exposure to hypoxic challenge elicited prompt decreases in Ti (Fig. 4A) that occurred at equal rates during the initial 60 s (Fig. 4B) in the SHAM and SCGX mice. Figure 4A shows that the decreases in Ti were subject to roll-off in the latter half of the hypoxic challenge and that Ti fell to baseline values more quickly in SCGX mice. Figure 4C shows that the total decreases in Ti were significantly smaller in SCGX mice than SHAM mice during the 106- to 300-s epoch due to the exaggerated roll-off seen in the SCGX mice. The decreases in Te during the hypoxic challenge initially occurred somewhat faster in the SCGX mice (Fig. 4, D and E) and were also subject to roll-off in both the SHAM and SCGX mice (Fig. 4D). This roll-off actually became an increase in Te during the latter stage of hypoxic challenge in SCGX mice (Fig. 4D). As a result, there was a significant difference in the total Te responses of SCGX mice (net increase) and SHAM mice (net decrease) during epoch 106–300 s (Fig. 4, D and F). As summarized in Table 2, the total Ti responses following return to room air in the SHAM and SCGX groups were similar to one another. In addition, the total Te responses following return to room air were equivalent in SCGX and SHAM mice (Table 2).

Hypoxic Challenge and Return to Room-Air Phases: Inspiratory Time/Expiratory Time and Inspiratory Quotient

Figure 5 summarizes the Ti/Te and inspiratory quotient [Ti/(Ti + Te)] responses in SHAM and SCGX mice during the 5-min hypoxic challenge and following re-exposure to room air. The Ti and Te responses observed during hypoxic challenge (Fig. 3) resulted in minor adjustments in Ti/Te and inspiratory quotient in SCGX and SHAM mice (Fig. 5, A, B, D, and E) that were of similar total magnitude (Fig. 5, C and F). The reductions in total Ti/Te and Ti/(Ti + Te) following reintroduction of room air were greater in the SCGX mice than SHAM mice (Table 2).

Hypoxic Challenge and Return to Room-Air Phases: End Inspiratory and Expiratory Pauses

Figure 6 summarizes the end inspiratory pause (EIP) and end expiratory pause (EEP) responses in SHAM and SCGX mice during the 5-min hypoxic challenge and following reintroduction to room air. Figure 6A shows that hypoxic challenge elicited an immediate reduction in EIP in SHAM and SCGX mice. Figure 6B shows that the initial changes were similar in the SHAM and SCGX mice, whereas Fig. 6C shows that the total EIP responses were significantly smaller in the SCGX mice during the 106- to 300-s epoch and the entire 0- to 300-s epoch. Figure 6D shows that hypoxic challenge prompted a decrease in EEP of ∼2 min in duration in both SHAM and SCGX mice. EEP went above baseline during the remaining 3 min of the hypoxic challenge in the SCGX mice whereas it returned to roughly baseline values in the SHAM mice. As seen in the Fig. 6E, the initial 60 s reduction in EEP occurred faster in SCGX mice as evident at 15 s. Figure 6F shows that the total increases in EEP recorded over the 106- to 300-s epoch and the entire 0- to 300-s epoch were significantly larger in the SCGX mice than SHAM mice. As shown in Figure 6, A and D, EIP and EEP gradually returned toward baseline values upon return to room air. Table 2 shows that the total EIP and EEP responses in the SHAM and SCGX mice were similar to one another.

Hypoxic Challenge and Return to Room-Air Phases: Inspiratory Drive and Expiratory Drive

Figure 7 summarizes the inspiratory drive (TV/Ti) and expiratory drive (TV/Te) responses in SHAM and SCGX mice during the 5-min hypoxic challenge and following reintroduction to room air. Figure 7, A and D, shows that hypoxic challenge elicited prompt increases in inspiratory and expiratory drives that displayed roll-off in both SHAM and SCGX mice. Figure 7, B and E, shows that the initial increases in inspiratory and expiratory drives occurred more quickly in SCGX mice compared with SHAM. Figure 7, C and F, shows that the roll-off was more pronounced in SCGX mice than in SHAM mice since the total inspiratory and expiratory drive responses were significantly smaller in the SCGX mice than SHAM mice during epoch 106–300 s. Figure 7A shows that there was an initial increase in inspiratory drive in the SHAM and SCGX mice upon return to room air that gradually subsided over time. Table 2 shows that the total changes in inspiratory drive in SHAM and SCGX mice were similar to one another. Figure 7D shows there was an initial increase in expiratory drive in the SHAM and SCGX mice upon return to room air that gradually subsided over time. Table 2 shows that the total changes in expiratory drive were similar in the SHAM and SCGX mice if all mice were considered. However, Table 2 shows that if the SHAM and SCGX mice were divided into two subgroups, the total changes in expiratory drive were significantly smaller (i.e., returned to baseline more quickly) in the SCGX mice when only the positive responses were considered.

Hypoxic Challenge and Return to Room-Air Phases: Peak Inspiratory and Expiratory Flows

Figure 8 summarizes the peak inspiratory flow (PIF), peak expiratory flow (PEF), and PIF/PEF responses in the SHAM and SCGX mice during the 5-min hypoxic challenge and upon reintroduction of room air. Figure 8A demonstrates that hypoxic challenge elicited immediate elevations in PIF in SHAM and SCGX mice that displayed a distinct roll-off. Figure 8B shows that the initial increase in PIF was similar in SHAM and SCGX mice, whereas Fig. 8C shows that roll-off was considerably faster in SCGX mice such that the total PIF responses during the 106- to 300-s epoch were significantly smaller in SCGX mice than SHAM mice. Figure 8D shows that the increases in PEF occurred faster in SCGX mice than SHAM mice, and Fig. 8E shows that the initial increase in PEF was significantly faster in the SCGX mice than SHAM mice at 45 s. Figure 8F shows that the total PEF responses were significantly smaller in SCGX mice than in SHAM mice during the 106- to 300-s epoch. Additionally, Figure 8, A, D, and G, shows that the return to room air caused initial increases in PIF and PEF in SHAM mice and SCGX mice and that the degree of these changes resulted in increases in PIF/PEF ratio. Table 2 shows that total changes in PIF, PEF, and PIF/PEF responses were equivalent in SHAM and SCGX mice when all of the mice were included in the analyses. However, Table 2 shows that the PIF/PEF ratio was substantially higher in SCGX mice if only the positive responses (+PIF/+PEF) were considered.

Hypoxic Challenge and Return to Room-Air Phases: EF₅₀, Rpef, and Relaxation Time

Figure 9 summarizes the EF₅₀, Rpef, and relaxation time (RT) responses in the SHAM and SCGX mice during the 5-min hypoxic challenge and following return to room air. As seen Fig. 9, A and B, hypoxic challenge caused a brief increase in EF₅₀ that happened more quickly in SCGX mice than in SHAM mice and Fig. 9B shows that the increase in EF₅₀ was significantly faster in the SCGX mice at 45 and 60 s. Figure 9C shows that the total EF₅₀ responses were significantly smaller in SCGX mice than SHAM mice during the 106- to 300-s epoch due to the more exaggerated roll-off in the SCGX mice compared with SHAM. Figure 9D shows that there were initial brief increases in Rpef in SHAM and SCGX mice on initiation of hypoxic challenge that were followed by sustained decreases in both groups. Figure 9E shows that the initial 60-s pattern of responses were similar in SHAM and SCGX mice although Fig. 9F shows that the reduction in Rpef from prevalues that occurred in the SHAM mice during the 106- to 300-s epoch was not observed in SCGX mice. Figure 9H shows that brief reductions in RT that occurred upon exposure to hypoxic challenge happened significantly more rapidly in SCGX mice than SHAM mice at 45 s. Figure 9G shows that the brief decreases in relaxation time during hypoxic challenge for both SHAM and SCGX mice were followed by substantial increases in RT during hypoxic challenge in SCGX mice such that the total RT responses the occurred in the 106- to 300-s epoch were significantly greater in SCGX mice than SHAM mice. Figure 9, A, D, and G, shows that restoration of room air was accompanied by the gradual return of EF₅₀, Rpef, and RT toward baseline values in SHAM and SCGX mice. Table 2 shows that there were no differences in EF₅₀ or Rpef responses between SHAM and SCGX mice following return to room air when all of the mice were included in the analyses, whereas RT was higher in the SCGX mice when all mice were included. The return to room-air responses were significantly greater in SCGX mice than SHAM mice if only positive responses for relaxation time (+RT) and Rpef (+Rpef) were considered (Table 2).

Hypoxic Challenge and Return to Room-Air Phases: Noneupneic Breathing Index, Noneupneic Breathing Index/Freq, and Apneic Pause

Figure 10 summarizes the noneupneic breathing index (NEBI), NEBI/Freq, and apneic pause (AP) responses that occurred in SHAM and SCGX mice during the 5-min hypoxic challenge and following restoration of room air. Figure 10A shows that hypoxic challenge elicited prompt elevations in NEBI of short duration for both SHAM and SCGX mice, and Fig. 10D shows variable changes in NEBI/Freq in both SHAM and SCGX mice. Figure 10, C and F, shows that the total NEBI and NEBI/Freq responses were similar in SHAM and SCGX mice, although changes in NEBI, but not noneupneic breathing index corrected for Freq (NEBI/Freq), was less in the SCGX mice between 106 and 300 s. Figure 10G shows that the degree of apneic pause decreased during hypoxic challenge in SHAM and SCGX mice, and Fig. 10I shows that the total reduction was more pronounced in SCGX mice. between 106 and 300 s. Figure 10, A, D, and G, shows that return to room air was associated with pronounced and rapid increases in NEBI, NEBI/Freq and the degree of apneic pause. As shown in Table 2, the total magnitude of these responses (NEBI, NEBI/Freq, and the degree of apneic pause) in SHAM and SCGX mice following return to room air were similar to each other when all mice were included in the analyses. Additionally, as seen in Table 2, when only negative apneic pause responses [(−Te/relaxation time)-1] were considered, it was evident that the SCGX mice showed a greater reduction in apneic pauses following return to room air (i.e., recovered more quickly) than SHAM mice.

DISCUSSION

The present study demonstrates that resting ventilatory parameters were equivalent in the SCGX and SHAM mice except for a greater incidence of apneic pauses in SCGX mice. Taken together, these data indicate that the absence of SCG projections to structures controlling breathing including brainstem nuclei, upper airway, and carotid bodies (42), does not dramatically affect baseline ventilatory timing and/or mechanics, whereas it can affect the quality of breathing by enhancing the incidence of noneupneic breaths (i.e., apneic pauses). The finding that resting noneupneic breathing index (NEBI) and NEBI/Freq in SCGX mice was not different from SHAM mice suggests that the incidence of central apneas and sighs (59) was not affected by the removal of SCG drive to central and/or peripheral structures. However, the enhanced degree of apneic pauses raises the possibility that the loss of SCG projections to the upper airway (e.g., nasopharynx, glottus, and epiglottus,) and tongue has subtle effects on the functional mechanics of these structures, although it is perhaps more likely to be due to the loss of glossopharyngeal sensory innervation of SIF cells (6). The increased incidence of apneic pauses in SCGX mice was not observed in CSCX mice (42), which suggests that the direct loss of SCG projections is not equivalent to cutting the preganglionic innervation to the SCG. It is expected that transection of the CSC would result in alterations in cellular processes within the SIF cells and postganglionic neurons of the SCG that, in turn, could lead to changes in firing properties of these interneurons and neurons, respectfully. However, we could find no published literature for the possibility that damage to the preganglionic motor neuron cell bodies within the T1−T4 spinal cord or their fibers within the CSC destined for the SCG would do anything to diminish the activities of postganglionic SCG neurons and SIF cells of the SCG. As such the question remains regarding why CSCX and SCGX have different effects on resting breathing. Several studies (71–74) provided evidence that the loss of preganglionic drive elicits sustained decreases in postganglionic nerve activity in sympathetic ganglia. In contrast, Qu and colleagues (72) found that firing of splenic and mesenteric postganglionic nerves continues unabated after damage to the spinal cord despite substantially diminished preganglionic outflow to splenic and inferior mesenteric ganglia. As such, the likelihood that postganglionic neurons and SIF cells in SCG continue to fire after transection of the CSC remains a possibility. It must be remembered that these SCGX studies were performed only 4 days after surgical removal of the SCG, and it is certainly a possibility that more exaggerated changes in resting breathing would occur at times greater than 4 days post-SCGX.

In agreement with Getsy et al (42), the present study shows that 1) hypoxic challenge elicited an initial increase in Freq in the SHAM C57BL6 mice that was associated with decreases in Ti, Te, EEP and relaxation time and an increase in Rpef, and all of these responses were subject to substantial roll-off; 2) the pronounced reductions in EIP were sustained throughout the hypoxic challenge; and 3) hypoxic challenge elicited substantial and sustained elevations in inspiratory and expiratory drives as well as TV, MV, PIF, PEF, and EF₅₀. The description of the C57BL6 mice with respect to ventilatory control processes and potential answers as to why some ventilatory parameters in C57BL6 mice express roll-off whereas other parameters do not was discussed in detail in Getsy et al. (42). While it is evident that the patterns and levels of noneupneic breathing in C57BL6 mice and their ventilatory responses to hypoxic challenge have a genetic component, there is no mechanistic evidence as to why ventilatory parameters in these mice do or do not display roll-off. We found that ventilatory mechanics, such as Rpef and relaxation time, and ventilatory timing parameters, such as Freq and EEP, display roll-off during hypoxic challenge, whereas mechanic parameters, such as PIF, PEF, and EF₅₀, and timing parameters, such as EIP, do not display roll-off. Coming to terms with how each of these ventilatory responses contributes to the overall hypoxic response will help to better define the physiological processes recruited by hypoxic challenge and perhaps help to identify the potential processes by which disease states elicit breathing disorders and abnormal responses to hypoxia.

In addition to studying resting ventilatory parameters, we show that freely moving C57BL6 male mice with SCGX displayed numerous altered responses to hypoxic challenge that also occurred in C57BL6 male mice with CSCX (e.g., more rapid occurrence of changes in frequency of breathing, minute ventilation, expiratory drive and EF₅₀), with the data for the CSCX study presented in Getsy et al (42). A qualitative comparison of the ventilatory responses that occurred during hypoxic challenge in CSCX mice and SCGX mice compared with their respective SHAM groups is provided in Table 3. Table 3, column 1, describes the actual response for each parameter (e.g., increase in frequency). The term “faster” in Table 3, columns 2 and 3, designated “Initial Response” denotes that the change in a particular parameter occurred more rapidly over the initial 60 s than in the SHAM group. The upward or downward directed arrows in Table 3, columns 4 and 5, designated “Epoch” indicates that the total responses recorded during the 106- to 300-s epoch of the 5-min hypoxic challenge were greater or lesser, respectively in the CSCX and SCGX mice than their respective SHAM controls. As can be seen, the changes in many parameters were altered in both CSCX and SCGX mice compared with their SHAM counterparts. Moreover, it is evident that the SCGX mice also displayed numerous changes in their responses to the hypoxic challenge that CSCX mice did not, including reductions in the total increases in Freq, MV, PIF, PEF, inspiratory and expiratory drives, and noneupneic breathing index (i.e., occurrence of noneupneic breaths) over the latter half of the hypoxic challenge (i.e., epoch 106–300 s). Therefore, from Table 3 and the findings of Getsy et al (42), it is evident that bilateral CSCX is not equivalent to bilateral SCGX, which suggests that hypoxia has a multiplicity of important effects on neural signaling within the CSC-SCG signaling pathways. Our data raise the possibility that hypoxic challenge directly activates subpopulations of cells in the SCG, such as SIF cells and postganglionic neurons, independently of increased CSC activity, and that this SCG drive to cardiorespiratory structures has dual roles of dampening the initial occurrence of the hypoxic ventilatory response, while promoting the overall magnitude of the response. These apparently divergent effects we see may be due to the loss of drive to peripheral and central structures with differential roles in the control of breathing in response to hypoxic challenge. The interpretation of our findings must include the compelling evidence that subpopulations of SCG cells and especially SIF cells are hypoxia sensitive (49, 50, 52, 54, 55) although there is equally compelling evidence that the SCG principal neurons themselves are not directly sensitive to hypoxia (46–51).

Table 3.

A qualitative assessment of the responses in C57BL6 mice with bilateral transection of the cervical sympathetic chain compared with their SHAM controls and in mice with bilateral removal of the superior cervical ganglia compared with their SHAM controls

Parameter	Initial Response		Epoch
	CSCX	SCGX	CSCX	SCGX
Increase in frequency	Faster			↓106-300
Increase in tidal volume		Faster	↑106-300
Increase in minute ventilation	Faster	Faster		↓106-300
Decrease in inspiratory time (Ti)	Faster			↓106-300
Decrease in expiratory time (Te)	Faster	Faster		↓106-300
Decrease in Ti/Te			↓106-300
Decrease in Ti/(Ti + Te)			↓106-300
Decrease in end inspiratory pause				↓106-300
Decrease in end expiratory pause	Faster	Faster
Increase in end expiratory pause				↑106-300
Increase in inspiratory drive	Faster	Faster		↓106-300
Increase in expiratory drive	Faster	Faster		↓106-300
Increase in peak inspiratory flow (PIF)				↓106-300
Increase in peak expiratory flow (PEF)	Faster	Faster	↑106-300	↓106-300
Decrease in PIF/PEF	Slower		↑106-300
Decrease in relaxation time (RT)	Faster	Faster
Increase in relaxation time				↑106-300
Decrease in rate of achieving PEF			↑106-300	↓106-300
Increase in EF50, ml/s	Faster	Faster		↓106-300
Increase in NEBI	Faster			↓106-300
Increase in NEBI/frequency	Faster	Slower		↓106-300
Decrease in apneic pause (Te/RT)-1				↑106-300

SHAM, sham operated; SCGX, superior cervical ganglionectomy; CSCX, bilateral transection of cervical sympathetic chain. NEVI, noneupneic breathing index; EF₅₀, expiratory flow at 50% exhaled tidal volume. ↑Enhancement compared with respective SHAM group. ↓Reduction compared with respective SHAM group.

The increase in noneupneic breathing index at the onset of hypoxic challenge and upon return to room air that we see in our study is a unique feature of C57BL6 mice, but not Swiss Webster mice or B6AF1 (C57BL6 dam × A/J sire) mice, which display minimal changes in noneupneic breathing index upon return to room air (59). We do not know the exact mechanisms by which noneupneic breathing index changes so remarkably in C57BL6 mice but conjecture that processes within carotid body glomus cells, carotid sinus nerve chemoafferents, and/or neurons within the NTS that process chemoafferent information allow for exaggerated expression of noneupneic breathing at the onset of hypoxic challenge and upon return to room air in the C57BL6 strain of mice.

Previous literature has shown that postganglionic cells in the SCG project to numerous structures that participate in the regulation of ventilation, including nuclei in the brainstem (e.g., NTS) and hypothalamus (25–34), and peripheral structures, such as the upper airway, tongue and carotid bodies (35–41). Additionally, there is substantial evidence that the SCG has important roles in the integration of the ventilatory and cardiovascular systems such that activation of the CSC in rats elicits profound decreases in upper airway resistance and arterial blood pressure (38, 39). As discussed by Prabhakar (75), there is no clear agreement regarding the ability of sympathetic nerve supply to the carotid bodies to influence the resting (normoxic) activity of glomus cells and/or chemoreceptor afferent nerve fibers and how this drive modulates the responses of these structures during hypoxic exposure. For instance, in cats, the CSC and ganglioglomerular nerve activity increases during exposure to hypoxic challenges (43–45, 76) and that activation of the ganglioglomerular nerve drive to the carotid bodies decreases the hypoxic responsiveness of the primary chemosensors (77). Moreover, diverse responses have been reported following application of sympathetic neurotransmitters, such as norepinephrine and dopamine, to in vitro and in vivo carotid body preparations, including 1) direct inhibitory effects on glomus cell and/or chemoreceptor afferent activity (78–88), 2) direct activation of glomus cells and/or chemoafferent fibers (89–93), 3) indirect excitation of glomus cells in the carotid body via constriction of microvascular blood vessels within the carotid body (76, 94), 4) a biphasic response pattern that consists of initial brief reduction in carotid sinus nerve activity followed by a longer-lasting period excitation (95), and 5) a biphasic response pattern that consists of initial brief burst in carotid sinus nerve activity followed by long-lasting inhibition (96). However, it must be remembered that the sympathetic neurotransmitters listed above are also present in populations of carotid body glomus cells and therefore can act in an autocrine-paracrine manner without postganglionic sympathetic involvement. In the context of sympathetic innervation of glomus cells, this is a confounding variable with respect to understanding the interplay between sympathetic nerves and glomus cells during a hypoxic challenge. At present, we do not know which postganglionic SCG nerves (i.e., internal carotid nerves, external carotid nerves and/or ganglioglomerular nerves) are involved in modulating the hypoxic challenge-induced ventilatory responses or what the exact targets are. We also do not know the exact nerve tracts and central projections of sensory afferents emanating from the SCG (6, 97). We are in the process of determining the effects resulting from transection of the above mentioned postganglionic sympathetic trunks to delineate the functional roles of postganglionic sympathetic innervation to the carotid bodies (i.e., studies involving transection of the ganglioglomerular nerve) as opposed to other central and peripheral structures (i.e., studies involving transections of the internal and/or external carotid nerves).

Ventilatory Responses That Occurred following Return to Room Air

Previous studies have found that the restoration of room air following hypoxic challenge can result in distinct respiratory patterns that can be categorized as posthypoxic frequency decline, when Freq falls below baseline levels (98) or short-term potentiation, when ventilation stays higher than baseline values for a considerable amount of time (59, 99). The C57BL6 mice used in this study and that of Getsy et al (42) showed substantially elevated Freq following reintroduction of room air that was associated with a prolonged phase of noneupneic (disordered) breathing. Potential mechanisms that underlie posthypoxic challenge disordered breathing include disturbances in brainstem signaling pathways (98, 100–102), but not altered signaling events within the carotid bodies (103, 104), although there is compelling evidence that disturbances in carotid body chemoreceptor afferent signaling have important roles in the etiology of sleep apneas (105). It should be noted that the posthypoxic challenge responses in the SHAM mice often showed considerable variability that prompted us to reanalyze the data and separate the mice into two subgroups: those that displayed net-positive responses (e.g., mice that had an overall increase in Freq during the 15-min recording period) and those that displayed net-negative responses (e.g., mice that had an overall decrease in Freq during the 15-min recording period). Since all mice appeared healthy, it would seem that this variability is related to the mixed genetics of the C57BL6 mice (106–111). When all 14 SHAM mice and 14 SCGX mice were compared following return to room air, we found that 1) the negative Ti/Te and Ti/(Ti + Te) responses were more pronounced in SCGX mice (largely because of differences in the changes in Te) (Table 2), and 2) relaxation time minimally changed in SHAM mice but significantly increased in SCGX mice (Table 2). When taking the dominant positive or negative responses into consideration, we found that 1) the total increases in Freq and expiratory drive were significantly smaller in SCGX mice, 2) the total increases in PIF/PEF ratios and Rpef were greater in SCGX mice, and 3) the total decreases in apneic pauses were greater in the SCGX mice. Therefore, it appears that the absence of SCG innervation to ventilatory control systems has significant consequences for parameters involved in ventilatory timing and mechanics.

Moreover, we have also determined how CSCX affects the relative number of dominant positive or negative posthypoxic challenge responses (see Table 2 in Ref. 42). For example, we see in apneic pause a shift from 5 mice in the SHAM group (45.5% of the 11 mice in the group) displaying a positive %change to 10 mice displaying a positive %change in the CSCX group (83.3% of the 12 mice in the group) and a shift from 6 mice displaying a negative %change in the SHAM group (54.5% of mice in the group) to only 2 showing a negative %change in the CSCX group (16.7% of mice in the group). Accordingly, it is evident that SCGX has a much greater influence on the relative populations of positive and negative responders during the posthypoxia phase than does CSCX.

Potential Roles of Small Intensely Fluorescent Cells in the SCG

Small intensely fluorescent (SIF) cells comprise ∼3% of the total SCG cell population and were initially described as dopaminergic interneurons that received innervation from preganglionic cholinergic fibers in the CSC (112, 113). Whereas principal SCG neurons are the main source of norepinephrine in the SCG, SIF cells are proportionately the richest source of dopamine and store ∼50% of total dopamine in the SCG (114, 115). Moreover, Libet and Owman (116) reported that dopamine released from SIF cells mediated slow inhibition of principal SCG neurons (i.e., dopamine generated slow inhibitory postsynaptic potential), thereby modulating postganglionic SCG nerve activity. SIF cells show morphological and biochemical similarities to hypoxia-sensitive carotid body glomus cells (117) and reside in clusters near fenestrated capillaries similar to carotid body glomus cells being located in clusters next to fenestrated capillaries (118). Unlike principal postganglionic SCG neurons, SIF cells present in the SCG possess O₂-sensing properties (53). Although direct evidence is lacking, there is strong support in favor of SIF cells being the hypoxia-sensitive cell type in SCG, including 1) the morphological plasticity following prolonged hypoxia is similar in dopaminergic carotid body glomus cells and SIF cells of the SCG (115, 117); 2) similar to carotid body glomus cells, SIF cells receive afferent and efferent innervation (6–9); 3) a combination of immunohistochemistry and retrograde labeling has allowed identification of three types of SIF cells in the SCG and similar to carotid body glomus cells, most afferent endings on SIF cells originated from the petrosal ganglion (6); and 4) afferent endings on SIF cells express purinergic P2X3 receptors (6), thus revealing a sensory pathway analogous to carotid body chemoreceptors where ATP acting on P2X3-containing postsynaptic receptors on chemosensory afferent terminals is a major mechanism for hypoxic chemo-transmission (119). Taken together, the possibility arises that the ability of SCGX (removal of all input/output signals) to affect the hypoxic ventilatory responses in a fashion clearly different from CSCX (removal of preganglionic innervation) may arise from 1) the ability of SIF cells to be directly activated by hypoxia independently of CSC activity, and 2) the ability of SCGX to remove the SIF cell-sensory neuron interactions within the SCG. It remains to be determined whether our findings are relevant to other species given the broad species-dependent variability in the sympathetic supply to the carotid body vasculature and the resulting control of carotid body sensitivity (97).

Study Limitations

A clear limitation of the present study and that of Getsy et al (42) is that mice were used 4 days after SCGX and CSCX (42), which only provides a relatively early view of the changes in ventilatory responses that occur in response to hypoxic challenge following these surgeries. The 4-day time point was selected because it is the earliest time that SCGX results in a substantial decrease in the number of tyrosine hydroxylase-positive sympathetic terminals in the carotid bodies (14, 61–63). It is important to note that we have not yet determined whether this substantial loss of sympathetic innervation of the carotid bodies occurred in our C57BL6 mice or whether there is a major decrease in sympathetic innervation of other vital structures, such as those within the brainstem and upper airway at this 4-day time point. This presence of sympathetic nerve terminals that are disconnected from their axons is an important consideration since hypoxia can produce action potential/extracellular Ca²⁺-independent release of catecholamines from these terminals (120–124). Therefore, it would seem vital to establish what the patterns of ventilatory responses to hypoxic challenge look like at longer time points after SCGX or CSCX. Moreover, the present SCGX study and companion CSCX study (42) raise the issue of the heterogeneity of responses that occur in the C57BL6 mice within the SHAM groups upon return to room air after posthypoxic challenge (i.e., some mice display positive changes in a particular parameter whereas others in the group display negative changes). Studies in males and females of other mouse strains, such as the A/J and Swiss-Webster, may help to define whether this heterogeneity is determined by sex and/or genetic factors.

With respect to the effects of hypoxic challenge on metabolism, it is known that a 5-min hypoxic challenge is associated with minor falls in body temperature in C57BL6 mice (56–60). As such, it could be expected that the decrease in oxygen availability would affect metabolic rate and perhaps explain why mice become progressively more still during the hypoxic challenge in this study. Future studies designed to address relationships between the changes in ventilatory performance and metabolic rate may greatly expand our understanding of how CSCX and SCGX affect the responses to hypoxic gas challenge. In addition, we did not perform plethysmography on the mice before surgery, which would have established resting ventilatory parameters in the absence of surgical stress and to allow for comparison to the values recorded 4 days postsurgery. However, we have established that the peak changes in Freq, TV, and MV recorded in the present studies (+116 ± 10 breaths/min, +70 ± 6 mL and +273 ± 23 mL/min, respectively) were very similar (+121 ± 12 breaths/min, +73 ± 6 mL and +294 ± 28 mL/min, respectively; P > 0.05, for all between-parameter comparisons) to those obtained in a group of naïve C57BL6 mice (n = 12) of similar age (96.3 ± 0.9 days) and body weight (27.1 ± 0.4 g).

Conclusions and Contributions to the Field

The data presented in Getsy et al (42) provide compelling evidence that in C57BL6 mice the CSC-SCG complex may provide inhibitory drive to central and peripheral structures that participate in the expression of the initial ventilatory responses to hypoxic challenge. However, the present study demonstrates that bilateral SCGX has greater effects on the overall hypoxic responses than the CSCX (42). As discussed above, this raises the possibility that hypoxic challenge may directly alter the activity of postganglionic neurons in the SCG independently of projections from the ipsilateral CSC. If the SIF cells rather than principal SCG neurons are indeed the direct hypoxia-sensitive targets (as would seem more likely), then their afferent connections via the glossopharyngeal nerve (6) may be sufficient to drive the chemoreflex response without principal SCG neuron involvement or CSC innervation. Moreover, the dopaminergic inhibitory pathway from SIF cells to principal SCG neurons could still be active during direct hypoxic stimulation of SIF cells, and this could further modulate postganglionic sympathetic drive to the carotid bodies (125, 126). A related key question is whether or not SIF cells in mouse SCG also receive a glossopharyngeal P2X3-mediated sensory innervation as reported for rat SCG (6) and whether this pathway is intact and functional following CSCX. Assuming that some SIF cells in mouse SCG receive cholinergic CSC and glossopharyngeal P2X3-afferent innervation as seen in rat SCG (6), the presence of this parallel chemoafferent reflex circuit may be sufficient to unify the present findings with those of Getsy et al (42).

Figure 11 shows a schematic diagram of our working model. Our data suggest that under physiologically normal conditions the preganglionic sympathetic fibers destined for the SCG course through the CSC and activate principal postganglionic SCG neurons (in blue) and inhibit small intensely fluorescent (SIF) cells (in pink) in the SCG. Postganglionic SCG fibers destined for structures, such as the upper airways, exit the SCG via the internal carotid nerve, and other postganglionic fibers exit via the external carotid nerve (ECN). There is also a small branch off the ECN called the ganglioglomerular nerve (GGN) that provide postganglionic SCG innervation to structures in the carotid body (CB), such as glomus cells and vasculature. The CB glomus cells are also innervated by chemosensory afferent fibers of the carotid sinus nerve (CSN), which branches off the glossopharyngeal (IX) nerve, and has cells bodies located in the petrosal ganglion (PG). These chemosensory afferents send projections to the commissural nucleus tractus solitarius (c-NTS), which signals the rostral ventral lateral medulla (RVLM) to alter ventilation. The SIF cells in the SCG are also innervated by sensory afferent fibers of the glossopharyngeal nerve with cell bodies in the PG and projections to the c-NTS. Following transection of the CSC, the preganglionic sympathetic innervation is removed from the principal postganglionic SCG neurons and the SIF cells. Thus, during hypoxia, populations of principal postganglionic SCG neurons can either sense hypoxia and become activated (via projections to the upper airway) or receive inhibitory dopaminergic input from hypoxia-sensitive SIF cells because of the disinhibition of SIF cells by cutting the CSC. Moreover, the hypoxia-sensing glomus cells activate the CSN and sends excitatory signaling to the c-NTS and the hypoxia-sensing SIF cells can now activate afferent sensory nerve endings in the glossopharyngeal nerve and increase overall excitatory discharge signaling (++) to the brainstem respiratory neurons in response to hypoxia. We propose from our data that this increase in ventilatory output is brief because hypoxia activation of SIF cells is transient and therefore after ∼60 s of hypoxia exposure SIF cells become desensitized and the magnitude of respiratory output is similar to SHAM control for the remainder of the 5-min hypoxic exposure. Removal of the SCG eliminated CSC drive to principal postganglionic SCG neurons and SIF cells, in addition to eliminating the cell bodies of the postganglionic SCG neurons and SIF cells, along with eliminating SIF cell-glossopharyngeal sensory afferent fibers. Therefore, during hypoxia, the increase in ventilatory output is due primarily from an increase in excitatory signaling from CB glomus cells to brainstem respiratory neurons. We propose from our data that sympathetic input is necessary for overall ventilatory responses to hypoxia exposure. In addition, without it we see reduced total ventilatory responses to hypoxia. Therefore, the activity of the CSC-SIF cell-glossopharyngeal circuit alone may account for the observed differences in respiratory parameters following CSCX (42) and SCGX. The SCG projects widely to both central and peripheral structures that control breathing, and we are performing studies to determine the effects of transection of the major postganglionic branches of the SCG (i.e., internal carotid nerve, external carotid nerve, and ganglioglomerular nerve) on ventilatory function in mice and their responses to hypoxic challenge. In addition, to determine the genetic and temporal aspects of SCGX in the control of ventilation and the responses to hypoxic challenge we are performing experiments using C57BL6, Swiss-Webster and A/J mice studied 4, 14, 28, and 56 days post-SCGX.

Figure 11.

Diagram of superior cervical ganglionectomy (SCGX) and cervical sympathetic chain transection (CSCX). Under physiologically normal conditions the preganglionic sympathetic fibers destined for the SCG course through the CSC and activate principal postganglionic SCG neurons (in blue) and inhibit small intensely fluorescent (SIF) cells (in pink) in the SCG. Following hypoxia (drop in Po₂), CSCX, which eliminates the cholinergic innervation of postganglionic SCG neurons and O₂-sensitive SIF cells, causes an increase in the initial hypoxic ventilatory response (HVR) by activating small intensely fluorescent (SIF) cells, which triggers a transient activation of glossopharyngeal afferent fibers to cause an increase (++) in overall excitatory signaling to brainstem respiratory neurons in response to hypoxia. On the other hand, following hypoxia, SCGX, which eliminates 1) the cholinergic innervation to postganglionic SCG neurons and O₂-sensitive SIF cells, 2) the cell bodies of postganglionic SCG neurons and O₂-sensitive SIF cells, and 3) SIF cell-glossopharyngeal sensory afferent fibers, blunts the total HVR and the expression of ventilation relies solely on the activation of O₂-sensitive carotid body glomus cells without modulation from SCG postganglionic fibers. ECN, external carotid nerve; CB, carotid body; ICN, internal carotid nerve; GGN, ganglioglomerular nerve; RVLM, rostral ventral lateral medulla; IX, glossopharyngeal nerve; c-NTS, commissural nucleus tractus solitarius. See Conclusions and Contributions to the Field for a more descriptive explanation of the schematic diagram.

Our study, which investigated the effects of CSCX in C57BL6 mice (42), showed that the CSC-SCG complex provides inhibitory drive to central structures (e.g., nuclei in the brainstem) and peripheral structures (e.g., carotid bodies and upper airway) that are responsible for the initial changes in breathing in response to hypoxic challenge. The present article extends these findings by providing evidence that the response to a hypoxic challenge in C57BL6 mice, which had undergone bilateral SCGX is not equivalent to those that had undergone bilateral CSCX (42). Thus, these data suggest that hypoxia may directly activate subpopulations of SCG cells, including SIF cells and/or principal SCG neurons, independently of innervation from the CSC. This knowledge of heterogeneity present within the SCG can potentially be used to target specific areas of the SCG when looking at various diseases that may involve disturbances in the CSC-SCG pathway, such as sleep-disordered breathing.

ETHICAL APPROVAL

The Institutional Animal Care and Use Committee of Case Western Reserve University reviewed and provided official approval for the studies presented in this manuscript.

DATA AVAILABILITY

The data that support this study are available upon e-mail request at sjl78@case.edu.

GRANTS

The experiments described in this manuscript were funded in part by National Institutes of Health-Stimulating Peripheral Activity to Relieve Conditions Award 10T20D023860 (Functional Mapping of the Afferent and Efferent Projections of the Superior Cervical Ganglion Interactome) to S.J.L.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.