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. 2014 May 16;592(Pt 13):2799–2811. doi: 10.1113/jphysiol.2014.273482

Chronic intermittent hypoxia–hypercapnia blunts heart rate responses and alters neurotransmission to cardiac vagal neurons

Jhansi Dyavanapalli 1, Heather Jameson 1, Olga Dergacheva 1, Vivek Jain 1,2, Mona Alhusayyen 1, David Mendelowitz 1
PMCID: PMC4221821  PMID: 24835174

Abstract

Patients with obstructive sleep apnoea experience chronic intermittent hypoxia–hypercapnia (CIHH) during sleep that elicit sympathetic overactivity and diminished parasympathetic activity to the heart, leading to hypertension and depressed baroreflex sensitivity. The parasympathetic control of heart rate arises from pre-motor cardiac vagal neurons (CVNs) located in nucleus ambiguus (NA) and dorsal motor nucleus of the vagus (DMNX). The mechanisms underlying diminished vagal control of heart rate were investigated by studying the changes in blood pressure, heart rate, and neurotransmission to CVNs evoked by acute hypoxia–hypercapnia (H–H) and CIHH. In vivo telemetry recordings of blood pressure and heart rate were obtained in adult rats during 4 weeks of CIHH exposure. Retrogradely labelled CVNs were identified in an in vitro brainstem slice preparation obtained from adult rats exposed either to air or CIHH for 4 weeks. Postsynaptic inhibitory or excitatory currents were recorded using whole cell voltage clamp techniques. Rats exposed to CIHH had increases in blood pressure, leading to hypertension, and blunted heart rate responses to acute H–H. CIHH induced an increase in GABAergic and glycinergic neurotransmission to CVNs in NA and DMNX, respectively; and a reduction in glutamatergic neurotransmission to CVNs in both nuclei. CIHH blunted the bradycardia evoked by acute H–H and abolished the acute H–H evoked inhibition of GABAergic transmission while enhancing glycinergic neurotransmission to CVNs in NA. These changes with CIHH inhibit CVNs and vagal outflow to the heart, both in acute and chronic exposures to H–H, resulting in diminished levels of cardioprotective parasympathetic activity to the heart as seen in OSA patients.

Key points

  • Chronic intermittent hypoxia–hypercapnia (CIHH) in adult rats evoked hypertension and blunted the heart rate responses to acute hypoxia–hypercapnia (H–H).

  • CIHH induced an increase in spontaneous inhibitory and decreased excitatory neurotransmission to cardiac vagal neurons.

  • CIHH completely abolished acute H–H evoked inhibition of GABAergic while facilitating glycinergic neurotransmission to cardiac vagal neurons of nucleus ambiguus.

  • These changes with CIHH inhibit cardiac vagal neurons to result in diminished cardioprotective vagal activity to the heart, characteristic of obstructive sleep apnoea.

Introduction

Individuals with obstructive sleep apnoea (OSA) have repetitive intermittent periods of hypoxia–hypercapnia (H–H) during sleep that is accompanied by arterial oxygen desaturations and increases in arterial carbon dioxide levels. OSA is an independent risk factor for the development of hypertension, coronary artery disease, and arrhythmias (Sanchez-de-la-Torre et al. 2013). Patients suffering from OSA have increases in blood pressure, lower heart rate variability, and reduced baroreflex sensitivity (Carlson et al. 1996; Trimer et al. 2013; Konecny et al. 2014), with chronic impairment in cardiac autonomic function i.e. sympathetic hyperactivity and diminished parasympathetic activity (Trimer et al. 2013). Animal models of OSA, using chronic intermittent hypoxia (CIH) closely mimic OSA in humans (Fletcher et al. 1992; Campen et al. 2005; Kline et al. 2007). While identification of the mechanisms underlying the elevations in sympathetic nerve activity in CIH and OSA has been the subject of numerous studies (Fletcher et al. 1999, 2002; Kc et al. 2010; Zoccal et al. 2011), in contrast, studies identifying the characteristics and mechanisms underlying depressed cardiac parasympathetic activity are scarce, hindering potential therapeutic interventions to restore cardioprotective parasympathetic activity to the heart in OSA.

The parasympathetic activity to heart arises from cardiac vagal neurons (CVNs) located in the nucleus ambiguus (NA) and dorsal motor nucleus of the vagus (DMNX) that dominate the control of heart rate (Mendelowitz, 1999). The preganglionic vagal efferent nerve terminals of the CVNs synapse with the postganglionic intracardiac ganglia neurons located within the connective and fat tissue surrounding sinoatrial and atrioventricular nodes (Armour, 2008). CVNs are typically intrinsically silent and thus depend on synaptic inputs (glutamatergic, GABAergic and glycinergic) to dictate their activity (Mendelowitz, 1996; Willis et al. 1996; Neff et al. 1998; Wang et al. 2001, 2003).

While it is known that CIH decreases the baroreflex control of heart rate and diminishes parasympathetic activity to the heart, these changes are not due to changes in parasympathetic innervation of the sinoatrial node or function within the cardiac ganglia (Gu et al. 2007; Lin et al. 2007; Yan et al. 2008). Although anatomical work has shown a decrease in efferent cardiac vagal innervation, ganglia size and density of axonal terminals within the cardiac ganglia after CIH (Soukhova-O'Hare et al. 2006; Lin et al. 2008), heart rate responses to vagal efferent stimulation are not diminished but rather enhanced (Gu et al. 2007; Lin et al. 2007; Yan et al. 2008). These results indicate that a central brainstem dysregulation of premotor CVN activity, but not cardiac ganglia function or vagal cardiac innervation, is responsible for the impaired parasympathetic control of heart rate that occurs with CIH. Supporting the hypothesis that CIH impairs CVN function in the brainstem, CIH diminished the bradycardia evoked upon microinjection of glutamate (Yan et al. 2008), as well as NMDA and AMPA (Yan et al. 2009), into the NA. However, beyond alterations in glutamate receptor density, little is known about how CIH impairs CVN function.

Acute exposure to hypoxia has been shown to evoke a biphasic change in heart rate, i.e. transient tachycardia followed by parasympathetically mediated bradycardia (Taylor & Butler 1982; Schuen et al. 1997), and these alterations were paralleled by an initial increase and a subsequent decrease in inhibitory (GABAergic and glycinergic) neurotransmission to CVNs (Neff et al. 2004). Also, acute intermittent hypoxia augmented respiratory related glutamatergic neurotransmission to CVNs (Griffioen et al. 2007). We hypothesized that rats exposed to repetitive episodic mild H–H 8 h day−1 for 4 weeks, as a model of OSA, would show increases in blood pressure to hypertensive levels and a blunted response to an acute H–H exposure. In this study we also identify the mechanisms responsible for diminished parasympathetic control of heart rate by studying the responses to an acute H–H challenge that mimic the cardiovascular challenge to a single bout of apnoea during OSA, as well as the long term changes in synaptic transmission to CVNs in NA and DMNX in response to 4 weeks of CIHH.

Methods

Ethical approval

All animal procedures carried out were in accordance with The George Washington University Institutional Guidelines and in compliance with the recommendations of the panel of Euthanasia of the American Veterinary medical association and the NIH publication (85–23, revised 1996) Guide for the Care and Use of Laboratory Animals. The minimal number of animals was used and care was taken to reduce any possible discomfort.

Labelling of CVNs

For animals in which electrophysiological recordings from CVNs were obtained neonatal Sprague–Dawley rats (postnatal days 2–5, Hilltop Laboratory Animals Inc, Scottdale, PA, USA) were anaesthetized using hypothermia by cooling to approximately 4°C. A right thoracotomy was performed and retrograde tracer X-Rhodamine-5-(and-6)-isothiocyanate (Invitrogen, USA) was then injected into the fat pads at the base of the heart to retrogradely label CVNs (Mendelowitz & Kunze 1991). The animals were then allowed to recover until they were 3–4 weeks old.

Telemetry implantation

For animals in which blood pressure and heart rate were obtained, male Sprague–Dawley rats, 3–4 weeks of age, were anaesthetized using isoflurane (2–4%) and a HD-X11 pressure transmitter was implanted (Data Sciences International, St Paul, MN, USA) with its cathether inserted into the abdominal aorta to record pressure and EKG leads were attached subcutaneously to obtain EKG recordings and heart rate. All rats with telemetry devices were allowed 7–14 days to recover from transmitter implantation surgery before any measurements were recorded. Blood pressure and heart rate were recorded via radio-frequency signals obtained through the Ponemah data acquisition system (Data Sciences International). Baseline recordings of blood pressure and heart rate were obtained for 3 days prior to CIHH exposure. Prior to, and during, the 28 day CIHH exposure period daily baseline recordings of blood pressure and heart rate were recorded.

Air or CIHH exposure

Adult rats (4 weeks old) that previously underwent surgery to label CVNs or for telemetry recordings were transferred to commercial atmosphere controlled chambers which contained their cages (with free access to food and water). This system (Oxycycler model A84, Biospherix, NY, USA) allowed for the monitoring and independent control of oxygen, carbon dioxide, and nitrogen. The animals inside the chambers were exposed to repetitive cycles of 3 min of mild H–H (6% O2 + 5% CO2 + 89% N2) followed by 3 min of normoxia (21% O2 + 79 % N2), repeated 10 times per hour, 8 h day−1, for 4 weeks. The animals were exposed to CIHH for 8 h during the light phase and to normal air during the remaining 16 h. ‘Unexposed’ animals that were exposed to normal air (21% O2 + 79% N2), were placed adjacent to the chambers during the exposure period to undergo similar handling, general lab conditions and background noise to the CIHH group. Animals for electrophysiological experiments (with cardiac labelling) were randomly divided into two groups, unexposed and CIHH exposed.

In vitro brainstem slice preparation

To obtain viable brainstem slices from mature animals, we adopted the methodology of Ye and colleagues (Ye et al. 2006). According to this method, glycerol based artificial cerebrospinal fluid (aCSF) was used for cardiac perfusion and brainstem slicing. Glycerol based aCSF contained (in mm): 252 glycerol, 1.6 KCl, 1.2 NaH2PO4, 1.2 MgCl, 2.4 CaCl2, 26 NaHCO3, and 11 glucose. Immediately following air or CIHH exposure for 4 weeks, rats were anaesthetized using isoflurane and placed on ice. Glycerol aCSF (4°C, pH 7.4, bubbled with 95% O2–5%CO2) was perfused transcardially at a speed of ∼10 ml min−1, after which the brain was quickly removed, glued on to a stage using 2% low melt agarose and placed in a vibrotome containing glycerol aCSF. Brainstem slices (330 μm thickness), containing either DMNX or NA, were obtained and briefly placed in a solution with the following composition (in mm): 110 N-methyl-d-glucamine (NMDG), 2.5 KCl, 1.2 NaH2PO4, 25 NaHCO3, 25 glucose, 110 HCl, 0.5 CaCl2, and 10 MgSO4 equilibrated with 95% O2–5% CO2 (pH 7.4) at 34°C for 15 min. NMDG based aCSF was used to help slices recover and to maintain viable brainstem slices for electrophysiological recordings (Zhao et al. 2011). The slices were then mounted in a recording chamber constantly perfused with normal aCSF comprising (in mm): 125 NaCl, 3 KCl, 2 CaCl2, 26 NaHCO3, 5 glucose and 5 Hepes; oxygenated with 95% O2–5% CO2 (pH 7.4) and allowed to recover for at least 30 min before an experiment was performed.

Electrophysiological recordings

CVNs in NA and DMNX were identified by the presence of the fluorescent tracer rhodamine and imaged using differential interference contrast optics and infrared illumination. Whole cell voltage clamp recordings from CVNs were done using Axopatch 200B and pClamp 8 software (Axon Instruments, Union City, NJ, USA), at a holding voltage of −80 mV at room temperature. The patch pipettes (2.5–5 MΩ) were filled with a solution consisting (in mm) of 150 KCl, 4 MgCl2, 10 EGTA, 2 Na-ATP and 10 Hepes or 150 K-gluconic acid, 10 Hepes, 10 EGTA, 1 MgCl2 and 1 CaCl2 at a pH of 7.3 for recording inhibitory or excitatory events, respectively.

Drugs were focally applied to CVNs using a pneumatic picopump pressure delivery system (WPI, Sarasota, FL). GABAergic inhibitory post synaptic currents (IPSCs) were isolated by focal application of solution containing strychnine (1 μm, glycine receptor antagonist), 6-cyano-7-nitroquinoxaline- 2,3-dione (CNQX, 50 μm, non-NMDA receptor antagonist) and d-2-amino-5-phosphonovalerate (AP5, 50 μm, NMDA receptor antagonist), with the puffer pipette positioned near the patched neuron. Glycinergic IPSCs were isolated by including gabazine (25 μm, GABA-A receptor antagonist), CNQX, and AP5 in the puffer pipette. The puffer pipette was filled with gabazine and strychnine to isolate glutamatergic excitatory postsynaptic currents (EPSCs).

Acute H–H

The respective EPSCs or IPSCs were recorded in control conditions for 5 min in the presence of aCSF equilibrated with 95% O2–5% CO2. Brainstem slices containing CVNs were exposed to H–H by superfusing the aCSF equilibrated with 85% N2–6% O2–9% CO2 for 10 min. Gabazine, strychnine, or CNQX, and AP5 were applied at the end of each experiment to confirm the targeted isolation of GABAergic, glycinergic, or glutamatergic activity, respectively. Each slice was exposed to hypoxia only once, limiting the experiments to only one CVN per slice of tissue. Gabazine, strychnine, CNQX, and AP5 were obtained from Sigma Aldrich (St Louis, MO, USA).

Data analysis

Synaptosoft software (version 6.0.3; Synaptosoft, Decatur, GA, USA) was used to analyse the synaptic events recorded from CVNs. Threshold value was set to the root mean square of noise levels multiplied by 5. The frequency and amplitudes of synaptic currents were grouped in 10 s bins and averaged for 2 min at the end of control and H–H. The data were presented as means ± SEM. To examine the chronic changes in blood pressure and heart rate over the 28 day CIHH exposure, daily values recorded before each CIHH exposure were statistically analysed by One-way repeated-measures analysis of variance (one-way ANOVA) followed by Bonferroni's multiple comparison test. Students unpaired t test was used to compare statistical significance between unexposed and CIHH exposed groups. For acute H–H evoked blood pressure and heart rate responses during CIHH exposure and in vitro experiments utilizing different conditions in the same CVN, Student's paired t test was used to test the significance using Graphpad Prism 5 software (La Jolla, CA, USA). Data with P < 0.05 was considered significant; in the figures, * denotes P < 0.05, ** denotes P < 0.01, *** denotes P < 0.001.

Results

Effect of CIHH on blood pressure

Blood pressure and heart rate were examined before and throughout 28 days of CIHH exposure. After 4 weeks of CIHH, systolic and diastolic pressure increased to hypertensive levels (from a systolic pressure of 105 ± 4.0 mmHg at the onset of CIHH to 144 ± 3.0 mmHg after 28 days of CIHH, n = 6; P < 0.05; one-way ANOVA) and diastolic pressure increased from 77 ± 1.0 mmHg to 110 ± 5.0 mmHg after 28 days of CIHH (n = 6; P < 0.05, one-way ANOVA; see Fig. 1A).

Figure 1. CIHH evokes hypertension, blunts bradycardic response and increases blood pressure in response to acute H–H.

Figure 1

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A, changes in systolic and diastolic blood pressure from day 1 (control) to day 28 of CIHH exposure. Systolic (continuous line) and diastolic (dashed line) blood pressures significantly increased from day 6 and day 16, respectively, compared to day 1 control and reached hypertensive levels by day 28 (n = 6; *P < 0.05, one-way ANOVA). The values represent average blood pressure recorded for 20 min during exposure to air in days prior to and during CIHH exposures. B, changes in heart rate in response to normoxia and an acute bout of H–H (3 min) at the onset and after 4 weeks of CIHH exposure (n = 6; *P < 0.05, Student's paired t test). C, changes in mean arterial blood pressure in response to normoxia and an acute bout of H–H (3 min) at the onset and after 4 weeks of CIHH exposure (n = 6; **P < 0.05, Student's paired t test).

Acute H–H evoked blood pressure and heart rate responses at the start and end of CIHH exposure

At the beginning of the 28 days of CIHH exposures, during a single exposure to H–H, heart rate decreased by 25% (438 ± 15 beats min−1 in normoxia and 325 ± 21 beats min−1 in acute H–H; n = 6; P < 0.05, paired t test), and this decrease in heart rate occurred without significant changes in blood pressure (99 ± 2 mmHg in normoxia and 97 ± 3 mmHg in acute H–H; n = 6; P > 0.05, paired t test). However, at the end of 4 weeks of CIHH exposure, acute H–H evoked a significant increase in blood pressure (112 ± 7 mmHg in normoxia and 123 ± 5 mmHg in acute H–H; n = 6; P < 0,01, paired t test) while there were no significant changes in heart rate (389 ± 23 beats min−1 in normoxia and 353 ± 27 beats min−1 in acute H–H; n = 6; P > 0.05, paired t test; see Fig. 1B and C).

Actions of CIHH on inhibitory neurotransmission to CVNs

GABAergic and glycinergic IPSCs were examined from CVNs both in the NA and DMNX of the brainstem from unexposed and CIHH exposed animals. In unexposed animals, the frequency of both GABAergic (7.9 ± 1.2 Hz, n = 48 in NA and 3.5 ± 0.3 Hz, n = 20 in DMNX; P < 0.05, unpaired t test) and glycinergic (4.4 ± 0.6 Hz, n = 29 in NA and 1.8 ± 0.2 Hz, n = 27 in DMNX; P < 0.001, unpaired t test) IPSCs in NA CVNs was greater than that in DMNX CVNs (see Fig. 2A and B). In addition, the amplitude of glycinergic IPSCs in CVNs of DMNX was significantly less than that of NA (58.6 ± 9.8 pA, n = 29 in NA and 23.6 ± 1.5 pA, n = 27 in DMNX; P < 0.01, unpaired t test). The amplitudes of GABAergic IPSCs in NA and DMNX CVNs were not different (44.0 ± 2.5 pA, n = 48 in NA CVNs and 46.6 ± 4.3 pA, n = 20 in DMNX CVNs; P > 0.05).

Figure 2. Effect of CIHH on GABAergic IPSCs in CVNs.

Figure 2

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Representative traces in control conditions showing GABAergic spontaneous IPSCs recorded from CVNs in NA (A) and DMNX (B) of unexposed (a) and CIHH exposed (b) animals while applying glycinergic and glutamatergic blockers. Quantitative bar charts depict the frequency (Ac and Bc) of GABAergic IPSCs in CVNs in NA (A) and DMNX (B), in unexposed and CIHH exposed animals. Number in parentheses represents n value. *P < 0.05, unpaired t test.

CIHH exaggerated the frequency of GABAergic (but not glycinergic) IPSCs in NA CVNs, whereas glycinergic (but not GABAergic) IPSC frequency was increased in DMNX CVNs following CIHH. The frequency of GABAergic IPSCs recorded from NA CVNs of CIHH exposed animals was 49% greater than that in unexposed animals (7.9 ± 1.2 Hz, n = 48 in unexposed and 11.8 ± 1.3 Hz, n = 51 in CIHH exposed; P < 0.05, unpaired t test; Fig. 2A). In DMNX, no change in GABAergic IPSC frequency to CVNs was observed between unexposed and CIHH exposed animal groups (3.5 ± 0.3 Hz, n = 20 in unexposed and 4.5 ± 0.7 Hz, n = 25 in CIHH exposed; P > 0.05, unpaired t test; Fig. 2B). The amplitude of GABAergic IPSCs to CVNs of NA and DMNX in unexposed group was not different from that of CIHH exposed group.

With respect to glycinergic IPSCs in CVNs, their frequency and amplitudes in CIHH and unexposed groups were not different in NA CVNs. However, in DMNX CVNs, the frequency of glycinergic IPSCs from the CIHH group was 50% greater compared to the unexposed group (1.8 ± 0.2 Hz, n = 27 in unexposed and 2.7 ± 0.4 Hz, n = 23 in CIHH exposed; P < 0.05, unpaired t test; see Fig. 3).

Figure 3. Effect of CIHH on glycinergic IPSCs in CVNs.

Figure 3

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Representative traces in control conditions showing glycinergic IPSCs recorded from CVNs in NA (A) and DMNX (B) of unexposed (a) and CIHH exposed (b) animals while applying GABAergic and glutamatergic blockers. Bar charts depict the frequency (Ac and Bc) of glycinergic IPSCs in CVNs in NA (A) and DMNX (B), in unexposed and CIHH exposed animals. Number in parentheses represents n value. *P < 0.05, unpaired t test.

Actions of CIHH on excitatory glutamatergic neurotransmission to CVNs

The amplitude of EPSCs in NA CVNs was significantly less than the amplitude of EPSCs in DMNX CVNs (18.0 ± 1.8 pA, n = 28 in NA and 34.1 ± 1.9 pA, n = 19 in DMNX; P < 0.001, unpaired t test). CIHH significantly reduced the frequency of glutamatergic EPSCs in CVNs in both NA (4.0 ± 0.4 Hz, n = 28 in unexposed and 2.7 ± 0.3 Hz, n = 24 in CIHH exposed; P < 0.01, unpaired t test) and DMNX (4.1 ± 0.3 Hz, n = 17 in unexposed and 2.3 ± 0.3 Hz, n = 18 in CIHH exposed; P < 0.001; unpaired t test) compared to unexposed group (see Fig. 4). CIHH also reduced the amplitude of EPSCs in DMNX, but not NA, CVNs (34.1 ± 3.2 pA, n = 17 in unexposed and 25.8 ± 2.5 pA, n = 18 in CIHH exposed; P < 0.05, unpaired t test; Fig. 4B).

Figure 4. CIHH inhibits glutamatergic EPSCs in CVNs.

Figure 4

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Representative traces in control conditions showing glutamatergic IPSCs recorded from CVNs in NA (A) and DMNX (B) of unexposed (a) and CIHH (b) exposed animals while applying GABAergic and glycinergic blockers. Bar graphs depict the frequency (Ac and Bc) and amplitude (Ad and Bd) of glutamatergic EPSCs in CVNs in NA (A) and DMNX (B), in unexposed and CIHH exposed animals. Number in parentheses represents n value. *P < 0.05; **P < 0.01; ***P < 0.001, unpaired t test.

Effect of acute H–H on inhibitory neurotransmission to CVNs in unexposed animals

GABA

In unexposed animals acute exposure to H–H inhibited the frequency of GABAergic IPSCs by 40% and 60% in the NA and DMNX CVNs, respectively (NA CVNs: 6.3 ± 1.0 Hz in control and 3.7 ± 0.5 Hz in H–H; n = 14; P < 0.05, paired t test; DMNX CVNs: 3.4 ± 0.5 Hz in control and 1.3 ± 0.3 Hz in H–H; n = 9; P < 0.001, paired t test; see Fig. 5A). In addition, H–H inhibited the amplitude of GABAergic IPSCs in DMNX CVNs (52.5 ± 3.7 pA in control and 42.4 ± 3.5 pA in H–H; n = 9; P < 0.05, paired t test) but not in CVNs within the NA (see Fig. 5A).

Figure 5. Actions of acute H–H on GABAergic IPSCs in CVNs of unexposed and CIHH exposed animals.

Figure 5

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A, traces represent GABAergic IPSCs recorded from CVNs in NA (a) and DMNX (c) of unexposed animals in control conditions and following H–H exposure for 10 min. Bar charts depict the frequency and amplitude of GABAergic IPSCs recorded from CVNs in NA (b) and DMNX (d) of unexposed animals in control and H–H (10 min) conditions. B, representative traces showing GABAergic IPSCs recorded from CVNs in NA (a) and DMNX (c) of CIHH exposed animals in control conditions and following H–H exposure for 10 min. Bar charts depict the frequency and amplitude of GABAergic IPSCs recorded from CVNs in NA (b) and DMNX (d) of CIHH exposed animals in control and H–H (10 min) conditions. *P < 0.05; ***P < 0.001, paired t test.

Glycine

Acute H–H inhibited the frequency of glycinergic IPSCs in DMNX CVNs by 50% (2.0 ± 0.3 Hz in control and 1.0 ± 0.2 Hz in H–H; n = 12; P < 0.01, paired t test). However the frequency and amplitude of glycinergic IPSCs in NA CVNs were unaltered by acute H–H (see Fig. 6A).

Figure 6. Actions of acute H–H on glycinergic IPSCs in CVNs of unexposed and CIHH exposed animals.

Figure 6

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A, traces represent glycinergic IPSCs recorded from CVNs in NA (a) and DMNX (c) of unexposed animals in control conditions and following H–H exposure for 10 min. Bar charts depict the frequency and amplitude of glycinergic IPSCs recorded from CVNs in NA (b) and DMNX (d) of unexposed animals in control and H–H (10 min) conditions. B, representative traces showing glycinergic IPSCs recorded from CVNs in NA (a) and DMNX (c) of CIHH exposed animals in control conditions and following H–H exposure for 10 min. Bar charts depict the frequency and amplitude of glycinergic IPSCs recorded from CVNs in NA (b) and DMNX (d) of CIHH exposed animals in control H–H (10 min) conditions. *P < 0.05; **P < 0.01, paired t test.

Effect of acute H–H on inhibitory neurotransmission to CVNs in CIHH exposed animals

GABA

Similar to the responses in the unexposed group, in animals exposed to CIHH acute H–H inhibited the frequency of GABA IPSCs in DMNX CVNs by 60% (4.5 ± 1.6 Hz in control and 1.3 ± 0.3 Hz in H–H; n = 11; P < 0.05, paired t test; see Fig. 5Bc and d). In contrast, in CIHH exposed animals the GABAergic responses to acute H–H on NA CVNs was abolished (7.1 ±1.2 Hz in control and 7.2 ± 1.5 Hz in H–H; n = 13; P > 0.05, paired t test; see Fig. 5Ba and b).

Similar to the responses in the unexposed animal group, in animals exposed to CIHH acute H–H reduced the amplitude of GABA IPSCs in DMNX CVNs (42.3 ± 4.5 pA in control and 33.8 ± 3.1 pA in H–H; n = 11; P < 0.05, paired t test) but not in NA CVNs (see Fig. 5B).

Glycine

Unlike the unexposed animals, in animals exposed to CIHH acute H–H significantly increased the frequency of glycinergic IPSCs in NA CVNs by 40%, without any significant changes in glycinergic IPSC amplitude (5.5 ± 0.9 Hz in control and 7.8 ± 0.9 Hz in H–H; n = 12; P < 0.05, paired t test; see Fig. 6). In animals exposed to CIHH, acute H–H inhibited the frequency of glycinergic IPSCs in DMNX CVNs by 25% (see Fig. 6B).

Effect of acute H–H on glutamatergic neurotransmission to CVNs of unexposed and CIHH exposed animals

Acute H–H had no effect on the frequency or amplitude of glutamatergic EPSCs to CVNs in NA and DMNX in both unexposed and CIHH exposed animals.

Discussion

The three key findings of this study are (1) rats exposed to CIHH for 28 days show significant increases in blood pressure, leading to hypertension, and a blunted heart rate response to acute H–H; (2) CIHH induced an increase in spontaneous inhibitory GABAergic IPSC and glycinergic IPSC frequency to CVNs in NA and DMNX, respectively; and a reduction in the excitatory glutamatergic EPSC frequency in both NA and DMNX CVNs, mechanisms likely to be responsible for the reduced vagal activity to the heart and reduced baroreflex sensitivity in OSA patients; and (3) CIHH blunted the bradycardia evoked by an acute bout of H–H and completely abolished acute H–H evoked inhibition of GABAergic IPSC frequency, and enhanced rather than inhibited glycinergic neurotransmission to CVNs in NA.

CIHH induces hypertension

OSA is a very common disease, occurring in ∼24% of adult males and ∼9% of adult females (Young et al. 1993), and increases the risk of hypertension, tachycardia, reduced heart rate variability and depressed baroreflex sensitivity. Other work has shown CIH evoked hypertension is mediated by increased peripheral chemoreflex sensitivity to hypoxia (Peng et al. 2001; Peng & Prabhakar 2004) and depressed baroreflex control of blood pressure (Lai et al. 2006). For instance, CIH augmented spontaneous glutamatergic transmission from chemoreceptor afferents, resulting in excitation of nucleus of solitary tract (NTS) second order neurons (Kline et al. 2007; Kline 2010). A direct activation of central chemoreceptors and an increase in ongoing paraventricular nucleus (Sharpe et al. 2013) and presympathetic rostro-ventrolateral medulla (Kc et al. 2010; Zoccal et al. 2011; Boychuk et al. 2012) neuronal activity is likely to contribute to increased sympathoexcitation in CIH related hypertension.

CIHH depresses baroreflex control of heart rate by modulating neurotransmission to CVNs

In contrast to the CIHH activation of the sympathetic side of the autonomic nervous system, CIHH impairs baroreflex sensitivity and diminishes cardiac parasympathetic activity (Lai et al. 2006; Soukhova-O'Hare et al. 2006; Lin et al. 2007; Yan et al. 2009). The parasympathetic cardiac vagal neurons located in NA and DMNX dominate the control of heart rate. GABA is the major inhibitory input to CVNs that regulates tonic and reflex control of heart rate (Wang et al. 2001). Previous studies using photo-uncaging and electrical stimulation approaches showed that GABA neurons projecting to NA CVNs are located medial and ventral to the nucleus ambiguus as well as close to the NTS (Wang et al. 2001; Frank et al. 2009). CVNs in NA and DMNX receive a major glutamatergic synaptic pathway from NTS that acts as a crucial link between increases in blood pressure and afferent baroreceptor activity and parasympathetic CVNs (Willis et al. 1996; Neff et al. 1998). Previous anatomical and microinjection studies (Chitravanshi et al. 1991; Batten 1995) have identified glycinergic fibres and receptors surrounding CVNs, however, the origin of these fibres to CVNs remains unknown.

A functional deficit in central nervous system baroreflex circuitry (Gu et al. 2007) rather than changes in peripheral or ganglionic vagal nerve activity or function (Lin et al. 2007) was identified to be responsible for CIH induced changes in baroreflex sensitivity. More specifically, CIH diminished the bradycardia evoked upon microinjection of glutamate (Yan et al. 2008), as well as NMDA and AMPA (Yan et al. 2009), into the NA, suggesting that CIH diminishes glutamate receptor density in CVNs. Interestingly, in support of this hypothesis, our data demonstrated that CIHH attenuated glutamatergic EPSC frequency to CVNs in both NA and DMNX; diminished EPSC amplitude in DMNX CVNs, and increased GABAergic and glycinergic IPSC frequency to CVNs of NA and DMNX, respectively. All of these changes in synaptic transmission would work together to inhibit CVNs, resulting in diminished vagal outflow to the heart and depressed baroreflex control of heart rate, hallmarks of OSA.

The amplitudes of GABA and glycine postsynaptic currents were unaltered by CIHH, indicating that CIHH had no effect on the density or responses upon activation of inhibitory postsynaptic GABA and glycine receptors, but CIHH acts on the function of the preceding GABA and glycine neurons that synapse on CVNs. With respect to glutamatergic transmission to DMNX CVNs, CIHH reduced both its EPSC frequency and amplitude, indicating that CIHH modulates the activity of presynaptic glutamatergic neuronal activity and postsynaptic glutamatergic receptor density and/or responses in CVNs. Electrophysiological and fluorescence imaging studies have indicated that acute intermittent hypoxia evoked an increase in reactive oxygen species within glutamatergic neurons of ventrolateral medulla and inspiratory related glutamatergic neurotransmission to CVNs which was attenuated by ROS scavengers (Griffioen et al. 2007), suggesting a role of ROS in mediating CIHH induced changes in glutamatergic transmission to CVNs.

Responses to acute hypoxia–hypercapnia

In response to changes in pH and gas concentrations induced by hypoxia, the chemoreflex evokes increases in blood pressure and a compensatory reduction in heart rate via modulating sympathetic and parasympathetic activities, respectively (Braga et al. 2008). In this study acute hypoxia–hypercapnia exposure in adult rats evoked a bradycardic response, and inhibited GABAergic and glycinergic neurotransmission to CVNs in NA and DMNX. These findings are consistent with previous in vivo studies showing biphasic changes in heart rate evoked by hypoxia, i.e. initial transient tachycardia followed by a sustained vagally mediated bradycardia (Taylor & Butler 1982; Schuen et al. 1997). In addition, previous studies in neonatal rats demonstrated that these biphasic changes in heart rate evoked by hypoxia were due to a transient increase followed by a sustained reduction in inhibitory GABA and glycinergic neurotransmission to CVNs in NA (Neff et al. 2004). In contrast, these biphasic changes in inhibitory neurotransmission to CVNs were replaced by a gradual reduction in GABA transmission with an increase in postnatal age (Dergacheva et al. 2013). In accordance with those findings, recordings from in vitro brainstem slices from adult animals in our study show only GABAergic and glycinergic inhibition to CVNs in response to acute H–H. Attenuation of GABAergic (in both NA and DMNX) and glycinergic (in DMNX only) transmission due to hypoxia relieves the inhibition on CVNs and increases parasympathetic activity to the heart, resulting in bradycardia.

CIH impairs the interaction between chemo- and baroreceptor autonomic reflexes due to desensitization of baroreflex sensitivity (Freet et al. 2013). An increase in blood pressure in response to acute H–H after 4 weeks of CIHH exposure can be accounted for increased chemoreflex induced sympathetic response. Previous studies have shown that CIH decreases heart rate responses to aortic depressor nerve stimulation (Gu et al. 2007). Our data confirm that the bradycardic response mediated by acute H–H was significantly blunted by exposure to CIHH. Indicating the mechanisms likely to be responsible for our in vivo data, CIHH completely abolished inhibition of GABAergic IPSC frequency in NA CVNs evoked by acute H–H, while significantly enhancing the glycinergic neurotransmission. A significant increase in glycinergic transmission and attenuation of H–H induced GABAergic inhibition to CVNs by CIHH would inhibit CVN activity and vagal outflow to the heart, resulting in a blunted bradycardic response to acute H–H. Consistent with these data, the magnitude of glycinergic inhibition to DMNX CVNs induced by acute H–H in the CIHH group was smaller than that in the unexposed group which would also contribute to a blunted bradycardic response. Taken together, data from this study indicate that changes in inhibitory neurotransmission to CVNs in response to acute H–H contribute to the blunted bradycardic response seen in CIHH exposed animals.

Conclusions

OSA, associated with CIHH, is characterized by sympathetic hyperactivity and diminished parasympathetic activity, leading to hypertension, tachycardia, decreased heart rate variability and depressed baroreflex sensitivity. The parasympathetic control of heart rate is dominated by CVNs located in NA and DMNX. The CIHH induced increase in basal spontaneous inhibitory glutamatergic synaptic input, and a reduction in excitatory glutamatergic synaptic input, to CVNs would inhibit parasympathetic vagal outflow to the heart, leading to tachycardia and increased blood pressure. In addition, acute H–H mediated potentiation of glycinergic neurotransmission to CVNs by CIHH also inhibits CVNs and parasympathetic control of the heart rate, ultimately resulting in the blunted baroreflex mediated bradycardic responses as seen in OSA patients.

Acknowledgments

We would like to thank Peter Byrne for his technical support.

Glossary

aCSF

artificial cerebrospinal fluid

AP5

d-2-amino-5-phosphonovalerate

CIH

chronic intermittent hypoxia

CIHH

chronic intermittent hypoxia–hypercapnia

CNQX

6-cyano-7-nitroquinoxaline-2,3-dione

CVN

cardiac vagal neuron

DMNX

dorsal motor nucleus of the vagus

EPSC

excitatory postsynaptic current

IPSC

inhibitory postsynaptic current

NA

nucleus ambiguus

NMDG

N-methyl d-glucamine

NTS

nucleus of the solitary tract

OSA

obstructive sleep apnoea

ROS

reactive oxygen species

RVLM

rostro-ventrolateral medulla

Additional information

Competing interests

None of the authors have any conflicts of interests.

Author contributions

Conception and design of the experiments: J.D., H.J., O.D., V.J. and D.M. Collection, analysis and interpretation of data: J.D., O.D., H.J., M.H. and D.M. Drafting the article or revising it critically for important intellectual content: J.D., H.J. and D.M. All authors read and approved this submission.

Funding

This study was supported by NIH Grants HL49965, HL59895 and HL72006 awarded to D.M.

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