Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2021 Dec 1.
Published in final edited form as: Auton Neurosci. 2020 Sep 28;229:102735. doi: 10.1016/j.autneu.2020.102735

Exaggerated potassium current reduction by oxytocin in visceral sensory neurons following chronic intermittent hypoxia

Heather A Dantzler 1, David D Kline 1
PMCID: PMC7704630  NIHMSID: NIHMS1635282  PMID: 33032244

Abstract

Oxytocin (OT) from the hypothalamus is increased in several cardiorespiratory nuclei and systemically in response to a variety of stimuli and stressors, including hypoxia. Within the nucleus tractus solitarii (nTS), the first integration site for cardiorespiratory reflexes, OT enhances synaptic transmission, action potential (AP) discharge, and cardiac baroreflex gain. The hypoxic stressor obstructive sleep apnea, and its CIH animal model, elevates blood pressure and alters heart rate variability. The nTS receives sensory input from baroafferent neurons that originate in the nodose ganglia. Nodose neurons express the OT receptor (OTR) whose activation elevates intracellular calcium. However, the influence of OT on other ion channels, especially potassium channels important for neuronal activity during CIH, is less known. This study sought to determine the mechanism(s) by which OT modulates sensory afferent-nTS mediated reflexes normally and after CIH. Nodose ganglia neurons from male Sprague-Dawley rats were examined after 10d CIH (6% O2 every 3 min) or their normoxic (21% O2) control. OTR mRNA and protein were identified in Norm and CIH ganglia and was similar between groups. To examine OTR function, APs and potassium currents (IK) were recorded in dissociated neurons. Compared to Norm, after CIH OT depolarized neurons and reduced current-induced AP discharge. After CIH OT also produced a greater reduction in IK that where tetraethylammonium-sensitive. These data demonstrate after CIH OT alters ionic currents in nodose ganglia cells to likely influence cardiorespiratory reflexes and overall function.

Keywords: afferent signaling, baroreflex, neuropeptides, ion channels, sleep apnea

1. INTRODUCTION

Oxytocin (OT) is a neuropeptide produced in the hypothalamus that induces a variety of physiological responses. OT release within and from the hypothalamus is multifold, including via neuronal dendrites where it may act as a volume transmitter to coordinate circuits within a nucleus (Son et al., 2013), into the cerebrospinal fluid to influence distant neuronal networks, and through direct axonal projections to number of nuclei (Veening et al., 2010). OT is released into the systemic circulation in response to periods of acute and chronic hypoxia (Goraca, 2004; Kelestimur et al., 1997; Stegner et al., 1984). In addition, OT is released from OTergic hypothalamic-originating projections into a variety of central nuclei, including the brainstem nucleus tractus solitarii (nTS) and the dorsal motor nucleus of the vagus (DMV) (Blevins et al., 2003; Maley, 1996; Ruyle et al., 2018; Sofroniew et al., 1980; Voorn et al., 1983). These regions constitute the primary integrating nuclei in the brainstem for vagal sensory afferent integration and its efferent output, respectively. The nTS expresses the OT receptor [OTR, (Gimpl et al., 2001)] and its activation by OT alters neuronal and autonomic function. For instance, OT in the nTS enhances nTS neuronal discharge (Henry et al., 1989; Raggenbass et al., 1989), baroreflex sensitivity, and lowers blood pressure and/or heart rate (Higa et al., 2002; Michelini, 2001); many of these effects are blocked by OTR antagonism (Michelini, 2007). OT also enhances afferent synaptic neurotransmission in the nTS, in part, via its increase in afferent glutamate release (Peters et al., 2008). Together, these data suggest OT modulates afferent activity that likely originate from vagal sensory neurons whose cell bodies lie in the nodose ganglia. In support of this concept, OTRs have been identified in nodose neurons (Welch et al., 2009) and their activation increases intracellular Ca++ and induces depolarization and action potential discharge (Iwasaki et al., 2015). Thus, OT may modulate not only afferent activity but visceral reflexes in health and disease.

Chronic intermittent hypoxia (CIH) is a model of the hypoxic episodes observed during obstructive sleep apnea and their autonomic and respiratory alterations (Prabhakar et al., 2002). In this model, days of CIH leads to desensitization of baroreflex sensitivity (Gu et al., 2007; Lin et al., 2007) and hypertension (Kunze et al., 2007; Weiss et al., 2007; Zoccal et al., 2007), mediated, in part by reduced sensory afferent-nTS neurotransmission (Almado et al., 2012; Kline et al., 2007). CIH also increases circulating OT (Wilson et al., 2018) which may influence nodose sensory neurons (Lacolley et al., 2006) as well as their central terminals in the nTS (Gross et al., 1990; Maolood et al., 2009). Neuronal release of OT into the DMV from hypothalamic neurons decreases heart rate, and when this occurs chronically, ablates CIH-induced hypertension (Jameson et al., 2016). While evidence supports a role for OT in cardiorespiratory synaptic and neuronal activity, and physiological reflexes during normoxia, its role during CIH is relatively unknown.

The goal of this study is to determine the mechanism(s) by which OT and OTR modulates sensory afferent activity normally and after CIH. We hypothesized CIH enhances OTR expression and function in nodose ganglia neurons. To test this hypothesis, we used expression analysis and whole cell patch-clamp electrophysiology. We demonstrate that after CIH OT activation of OTR depolarizes neurons, and decreases action potential discharge and outward voltage-gated potassium currents.

2. METHODS

2.1. Animals and ethical approval.

Electrophysiological, molecular and ELISA experiments were conducted on male Sprague-Dawley rats (Envigo, IN, USA) aged 5–6 weeks. In brief, 3 wk old rats were allowed to acclimate to the vivarium from ~1 week prior to placing them in the hypoxic chamber for an additional 3 days of acclimation, after which animals were exposed to 10 days intermittent hypoxia or normoxia. Rats were housed in standard rat cages on a 12:12-h light-dark cycle with food and water available ad libitum in an Association for Assessment of Laboratory Animal Care-accredited vivarium of the Dalton Cardiovascular Research Center. All experiments were conducted in accordance with National Institutes of Health guidelines and approved by the Animal Care and Use Committee of the University of Missouri. Male rats were used to extend our previous studies examining the neural networks and neuropeptides responsible for CIH and hypoxic responses (Kline et al., 2007; Martinez et al., 2020; Ruyle et al., 2018).

2.2. CIH exposure.

Rats were exposed to CIH for 10 days similar to our previous work (Kline et al., 2019). Briefly, freely moving rats in standard cages inspired alternating 3 min cycles of oxygen and nitrogen to produce periods of normoxia (21% O2, 3 min) and hypoxia (6% O2, ~45 sec). Cyclic hypoxia occurred for 10 episodes per hour between 9:00 a.m. to 5:00 p.m., with 21% O2 between 5 p.m. and 9 a.m. CIH exposure occurred via a commercially available system (Biospherix LTD, Parish NY). Normoxic control animals were exposed only to 21% O2, with intermittent air flow from 9:00 a.m. to 5:00 p.m. Experiments and tissue/blood collection (except 1 hr post-CIH blood collection) were conducted the morning after the final CIH exposure or its appropriate normoxic control period.

2.3. Measurement of arterial blood pressure, heart rate and baroreflex.

Four to five-week old rats were instrumented with a radiotelemetry transmitter (HD-X11, Data Sciences International, St. Paul, MN) in the abdominal aorta. Implants occurred via Envigo surgical services prior to arrival at the Dalton Cardiovascular Research Center vivarium. After a 1-wk recovery, rats were placed in the hypoxic chamber for acclimation. Blood pressure was acquired for 5 minutes every hour (sampling rate 1000 Hz) for 2 days in room air prior to initiating either 10d CIH or normoxic exposure. Mean arterial pressure (MAP) and heart rate (HR, determined from the systolic pressure) values were acquired for 5 minutes each hour. MAP and HR was averaged for each 5 minute period, and then averaged during the hypoxic light period (9:00 a.m. – 5:00 p.m.) and dark period (7:00 p.m. – 7:00 a.m.).

Telemetry data was analyzed for spontaneous baroreflex sensitivity (sBRS) using the sequence method with the aid of HemoLab software (http://www.haraldstauss.com/HaraldStaussScientific/hemolab/). Analysis was performed on 5 min of data during the 12-hr dark (non-hypoxic) cycles at 0, 1 and 10d CIH (or their normoxic controls). This time period was chosen to avoid direct influence of hypoxic episodes (or their transition) on sBRS that occurred during the day. Baroreflex sequences were identified as ramps of four or more consecutive beats in which both systolic BP and interpulse interval (PI) increased or decreased simultaneously, and sBRS was calculated via linear regression (R for inclusion > 0.8) as the slope (in ms/mmHg) of that relationship.

2.4. Enzyme-linked immunosorbent assay (ELISA) of OT.

Circulating plasma OT was measured (Leng et al., 2016) in normoxic and CIH-exposed rats. Rats were anesthetized with isoflurane and following decapitation trunk blood was collected, placed in K2 EDTA tubes (BD Vacutainer, cat 36643) and spun at 1600 x g at 4°C for 15 minutes. The supernatant was collected and following addition of protease inhibitor cocktail (0.5 μL/mL, Sigma, cat P1860) the plasma was stored at −80°C. Oxytocin concentration was measured with DetectX Oxytocin Enzyme Immunoassay kit (Arbor Assays, cat K048) according to manufacturer’s instructions. All samples were ran in duplicate. The optical density of the circulating OT was measured against a standard curve using a Multi-Detection Microplate reader (BIO-TEK, Synergy HT).

2.5. Reverse transcription real-time polymerase chain reaction (RT-PCR) of OTR.

Experiments were performed similar to our previous work (Matott et al., 2020). Normoxic and 10d CIH isoflurane anesthetized rats were decapitated and the nodose ganglia was removed, flash frozen in liquid nitrogen and stored at −80°C until use. mRNA was isolated via RNaquoeus-micro kit (Invitrogen, cat AM1931), quantified, and 100 ng of mRNA was used to generate cDNA via SuperScript III First Strand kit (Invitrogen, cat 18080–051). Relative RT-PCR was performed with 2 μL of cDNA using PowerUp SYBR Green master mix (Applied Biosystems), SmartCycler System (Cepheid) and primers for oxytocin receptor (Oxtr, NM_012871.3, forward: AGC GTT TGG GAC GTC AAT, reverse: GTT GAG GCT GGC CAA GAG, 10 μM) and the housekeeping gene Beta-2-microglobulin (B2m, NM_012512.2, FOWARD: AGC AGG TTC CTC AAA CAA GG, reverse: TTC TGC CTT GGA GTC CTT TC, 10 μM). Controls included no template and no primer. All products were visualized on 1.5% agarose gel. The amount of Oxtr mRNA was normalized to B2m using the 2−ΔΔCT method (Livak and Schmittgen 2001).

2.6. Immunohistochemistry (IHC) of OTR.

As previously reported (Ramirez-Navarro et al., 2011), nodose ganglia were collected from isoflurane anesthetized normoxic and 10d CIH rats and flash frozen in liquid nitrogen. Tissue was cryoprotected in 30% sucrose, sectioned at 10 μm on a cryostat (Leica 1850) and briefly fixed in ice-cold 4% paraformaldehyde. Tissue sections were stored in 0.01 M phosphate buffered saline (PBS, pH 7.4) at 4°C until processed. All IHC steps were carried out in a humidified chamber. Nodose ganglia sections were pre-blocked in PBS-0.3% Triton X100 (PBS-T) containing 10% normal donkey serum (Millipore, cat S30) for 30 minutes at room temperature. Sections were subsequently incubated in goat anti-oxytocin receptor antibody (OTR, 4 μg/mL, Abcam) in PBS-T with 1% normal donkey serum for 12 hours at 4°C. Antibody specificity has been previously characterized (Gonzalez-Iglesias et al., 2015). Following PBS wash (2X 15 min), sections were incubated for 2 hours in donkey anti-goat antibody conjugated to Cy2 (1:200, Jackson ImmunoResearch) then coverslipped with ProLong Diamond antifade with DAPI (Invitrogen, cat P36962). Control sections were incubated without primary antibody. An Olympus epifluorescent spinning disk confocal system equipped with an Orca Hammamatsu CCD camera was used to examine sections.

2.7. Immunoblot of OTR.

Norm and CIH nodose ganglia were collected and flash frozen in liquid nitrogen and stored in −80°C until used. Ganglia were homogenized in buffer containing [(in mM, pH 7.4) 250 sucrose, 10 Tris, 1 EDTA, and protease inhibitors (Complete Mini, EDTA-free tablets, Roche)] followed by incubation on ice for an additional 30 min. The homogenate was then centrifuged at 13,300 RPM for 10 min at 4°C and the supernatant collected. Protein concertation of the supernatant was determined by the BCA method. Protein (80 μg) was mixed with 4x loading dye (Laemmli Sample Buffer, Bio-Rad), boiled at 90°C (5 min), separated on a Tris-HCl gel and transferred to a polyvinylidene difluoride membrane (Bio-Rad). The membrane was pre-blocked in Tris Buffered Saline with 0.1% Tween 20 (TBS-T) containing 5% powdered milk for 1 hr on a shaker at room temperature. The membrane was rinsed 3X 5 min in TBS-T then incubated overnight in rabbit anti-OTR antibody in TBS-T with 5% milk (1:5000, Abcam, cat ab181077) at 4°C overnight on a shaker. Following a TBS-T wash (3X 5 min) the membrane was incubated in donkey anti-rabbit antibody conjugated to peroxidase (1:10:000, Jackson ImmunoResearch) for 2 hr on shaker at room temperature. After a TBS-T wash (3X 5 min) the membrane was incubated in Clarity Western ECL (Bio-Rad, cat 1705061) for 5 min and imaged on the Bio-Rad ChemiDoc. Mouse anti-tubulin (1:1000, Abcam) was used as a loading control. All blots were normalized to tubulin. Quantification was performed using the ImageJ Gel plugin.

2.8. Nodose neuron isolation and electrophysiology.

As previously (Kline et al., 2009), normoxic and CIH-exposed rats were anesthetized with isoflurane and decapitated. The nodose ganglia was quickly removed and placed in ice-cold Earle’s balanced salt solution (EBBS, Gibco, cat 14155063). After resting for 5–10 min, tissue was transferred to EBBS containing 335 U/mL Collagenase Type 2 (Worthington Biochemical, cat LS004176) and 0.5 U/mL Dispase (Worthington Biochemical, cat LS020109) and incubated for 1 hr at 37°C in a shaking (110 RPM) water bath. The enzyme solution was replaced with trituration solution containing DMEM/F12 media (Gibco, cat 11320–033), 5% fetal bovine serum (Gibco, cat 16140063), 1x Penicillin-Streptomycin-Neomycin (Gibco, cat 1564055), Mito+ Serum Extender (Corning, cat 355006) and 0.1% Bovine Serum Albumin (Sigma Aldrich, cat A4378). Ganglia were manually disassociated by repeatedly passing tissue through a fire-polished glass Pasteur pipette of gradually decreasing tip size. The cells were concentrated at 70 RPM for 4 min, the supernatant discarded, and the pellet re-suspended in the above media with the addition of Nerve Growth Factor 7.0s (Chemicon, cat NC010). Cells were plated on 15 mm poly-D-lysine (100 μg/ml) treated glass coverslips and allowed to settle for 2 hr in an incubator (5% CO2, 37°C) before recording.

Neurons were recorded no later than 5 hrs following dissociation. Glass coverslips covered with neurons were placed in Series 40 Quick Change Imaging Chamber (Warner RC-40-LP) and immersed in room temperature physiological recording saline [(in mM, pH 7.4, 300 mosM) 137 NaCl, 3 KCl, 1 MgCl2+6 H2O, 2 CaCl2, 10 Glucose, 10 Hepes]. Recording electrodes (4.0–6.0 MΩ) contained (in mM, pH 7.3, 295–300 mosM) 130 K-gluconate, 10 NaCl, 10 Hepes, 11 EGTA, 1 MgCl2, 2 MgATP, 1 CaCl2 and 0.2 NaGTP. Outward potassium currents (IK) were recorded in voltage clamp mode (Vhold at −60 mV) and evoked by depolarizing voltage steps from −70 to +60 mv (10 mV steps, 140 ms). To examine action potential (AP) discharge, neurons were recorded in current clamp mode and evoked via current injection (−20 to 50 pA current steps, 516 ms, 50 pA steps). In some voltage clamp experiments, to eliminate the influence of sodium and calcium on the analyzed currents, cells were perfused with N-Methyl-D-glucamine based solution [NMDG, (in mM, pH 7.4, 300 mosM) composed of 137 NMDG, 3 KCl, 1 MgCl2+6 H2O, 0.02 CaCl2, 10 Glucose, 10 Hepes] while recording electrodes contained (in mM, pH 7.4, 295–300 mosM) 145 K-Aspartate, 0.3 CaCl2, 10 Glucose, 5 HEPES, 1 MgCl2+6 H2O, 2.2 EGTA.

Currents and discharge were examined in the presence of OT (300, 600, 1000 nM, Tocris), the OT agonist Thr4, Gly7 -Oxytocin (TGOT, 600 nM, Bachem) and/or OT receptor antagonist L-368,899 HCl (1 μM, Tocris). OT concentrations were based on previous reports (Gong et al., 2015; Han et al., 2018; Iwasaki et al., 2015). The K+ channel blockers Tetraethylammonium (TEA, 10 mM) or 4-Aminopyridine (4-AP, 5 mM) were applied in some protocols (Glazebrook et al., 2002; Moak et al., 1993). Following 5 minutes of baseline, pharmacological agents were bath applied for three minutes followed by three minutes washout. Each cell was only exposed to a single concentration of OT.

2.9. Data analysis and statistics.

Recordings were analyzed in Clampfit 10 and Excel 2016. Statistical significance was tested in GraphPad Prism 7. Reported electrophysiological parameters were evaluated at the end of baseline and OT receptor manipulation. Action potentials during OT were normalized (i.e., “1”) to the peak discharge during baseline. Potassium currents were examined during the immediate 5 ms (i.e, early currents, excluding initial sodium and capacitive currents) and during the last 3 ms (i.e., steady state currents). OT-sensitive currents were determined by subtracting the currents during OT from those of its proceeding baseline. All currents were normalized to the cell’s initial capacitance (i.e, pA/pF). Current-voltage plots for the early and steady state currents and the effects of OT were evaluated via two-way repeated measures (RM) ANOVA followed by Holm-Sidak post hoc test. OT-sensitive currents, membrane potential and action potential discharge were evaluated by two-way ANOVA followed by Fisher’s least significant difference (LSD) post hoc test. Blood pressure prior to CIH (0d) and following 1 and 10d CIH (or normoxic controls) was compared within groups via one-way repeated measures ANOVA. Comparing normoxic versus CIH mRNA, immunoblot, plasma OT, sBRS and changes in MAP and HR were compared via t-test. P values < 0.05 were considered significant. Data are shown as mean ± SEM. “N” denotes number of animals, “n” denotes number of cells.

3. RESULTS

3.1. CIH increases blood pressure.

Rats were exposed to 10 days of chronic intermittent hypoxia (10d CIH) or 10-day normoxic control. Figure 1A shows mean arterial pressure (MAP) in 4 rats prior to (0d) and after 1 and 10 days CIH. As shown, MAP was greater after 10d CIH (one way ANOVA) in both the hypoxic light (9 a.m. – 5 p.m.) and dark (7 p.m. – 7 a.m.) periods. The increase in MAP was greater than that of a comparable normoxic exposure (Fig 1B). Heart rate increased in the light (hypoxic) period after 10d CIH but was unaltered in the dark period (Fig 1C). The increase in HR during the light period was greater than normoxia (Fig 1D). Spontaneous baroreflex sensitivity (sBRS, i.e., gain) was not altered in the up, down or total sequences after 10d CIH (Fig 1E).

Figure 1. Cardiovascular parameters following CIH.

Figure 1.

Open in a new tab

A. Mean arterial pressure (MAP, mmHg) in CIH rats prior to (0d) and after 1 and 10 days CIH (1 & 10d). Note the increase in MAP after 10d CIH (one-way RM ANOVA). B. Comparison of the changes in MAP after 10d of CIH exposure (n=4) or normoxia controls (n=2). C, D., Heart rate (beats per minute, bpm) after 0,1 & 10d CIH (C) and its change compared to normoxia controls (D). E. Baroreflex sensitivity (BRS) in CIH rats during the night (dark) period. Shown is the total, up and down sequences. A,C, E, one way RM ANOVA; B,D, t-test within light or dark periods.

3.2. Expression of oxytocin receptors (OTR) in the nodose ganglia and circulating OT after CIH.

Twelve hours following the last hypoxic exposure on day 10 of CIH, rats were anesthetized and decapitated for nodose ganglia removal and capturing of core blood. Within the nodose ganglia, mRNA expression of Oxtr did not change after 10d CIH (Norm and CIH, N=3 each, Fig 2A, p=0.51, t-test). Quantitative assessment of OTR protein expression via immunoblot demonstrated similar OTR after 10d CIH (Norm and CIH, N=11 each, Fig 2B&C, p= 0.08, unpaired t-test). OTR-immunoreactivity was localized to somas and fibers within CIH and Norm nodose ganglia (Fig 2D). ELISA examination of circulating oxytocin (OT) was comparable between Norm and 10d CIH animals (Norm and CIH, N=5–6 each, Fig 2E). To examine if the comparable plasma OT in Norm and CIH was due to the 12hrs between hypoxia exposure and tissue collection, in a separate group of rats we measured plasma OT immediately (< 1 hr) following the last hypoxic exposure on the 10th day of exposure. Plasma OT was greater in CIH than Norm in these rats (N=4 each, p = 0.03, Fig 2F). These data suggest CIH does not alter OTR expression; the functional influence of OTR was studied further.

Figure 2. Oxytocin receptors (OTR) are present in CIH nodose ganglia neurons.

Figure 2.

Open in a new tab

A. Oxtr mRNA expression in nodose ganglia after Norm and CIH (N = 3 each), determined by the 2−ΔΔCT method. Oxtr was normalized to the housekeeping gene B2m. CIH did not alter Oxtr mRNA expression. B. Example of immunoblot with Norm and CIH nodose protein (80 μg, n=3 each) showing a band for OTR at ~43 kDa (arrowhead). Molecular markers shown in left most lane. Loading control tubulin shown below. C. Immunoblot quantification of OTR protein expression normalized to tubulin (N = 11 each) D. OTR immunoreactivity in fibers and somas of nodose neurons after Norm and CIH. Scale = 200 μm. E, F. Circulating plasma OT levels (pg/ml) 12 hrs (E., Norm and CIH, N = 5 each) and within 1hr (F., Norm and CIH, N = 4 each) after the last hypoxic challenge. A, C, E, F; unpaired t-test, p < 0.05.

3.3. OTR activation alters nodose neuron membrane potential and action potential properties after CIH.

The contribution of circulating OT and CIH was evaluated via examination of the neuronal membrane potential (Vm), action potential (AP) discharge, and AP properties. OT was studied in isolated nodose neurons at 300, 600 and 1000 nM for 3 minutes within 5 hours of isolation. Control cells consisted of only those exposed to OT vehicle (Veh, physiological saline) for a similar amount of time. In Norm-exposed neurons, Vm was not altered by Veh (N=3, n=5) or any concentration of OT (300 nM, N=4, n=7; 600 M, N=4, n=11, 1000nM, N=2, n=5) compared to its initial baseline (Bsl, Fig 3A only Veh and 600 nM OT shown). Following CIH, 300 (N=4, n=9) and 600 (N=3, n=5) nM OT depolarized nodose ganglia neurons (600 nM shown in Fig 3B) whereas Veh (N=3, n=7) and 1000 (N=4, n=5) nM OT was without effect. Across all neurons examined under baseline conditions and independent of subsequent OT concentration application, the membrane potential of Norm and CIH neurons was comparable (Norm, n=30, −51.9 ± 1.9 mV vs CIH, n=26, −56.3 ± 1.9 mV, p=0.119, t-test).

Figure 3. Oxytocin (OT) depolarizes membrane potential (Vm) and alters action potentials (AP) after CIH.

Figure 3.

Open in a new tab

A. Vm is not altered in Norm neurons during vehicle (n=5) or 600 nM OT (n=11) exposure. B. After CIH, 600 nM OT depolarized neurons (n=5). There was no effect in vehicle (n=7). C. After Norm AP discharge is not altered during vehicle (n=7) and 600 nM OT (n=11). D. Following CIH, 600 nM OT (n=5) but not vehicle (n=8) reduced AP discharge. Inset, example of the initial action potential in CIH-exposed neurons (black), and the reduction in AP threshold by OT (red). Scale = 100 mV and 50 ms. *, p < 0.05 vs Bsl, Two-way RM ANOVA with Holm-Sidak. Bsl was physiological solution.

Depolarizing current steps induced AP discharge in nodose ganglia neurons. After Norm exposure, Veh (N=3, n=7) and 300 nM OT (N=4, n=9, not shown) did alter the number of events within a current step (Veh in Fig 3C); OT at 600 (N=4, n=11) and 1000 nM (N=2, n=5) increased discharge at the highest current steps. By contrast, after CIH OT decreased discharge which was significant at 300 (N=4, n=9, not shown) and 600 (N=3, n=5) nM whereas Veh (N=3, n=8) was without effect (Fig 3D). The properties of the initial evoked AP (i.e., amplitude, afterhyperpolarization, half-width, threshold, rise and decay) were not altered by Veh after Norm or CIH. These properties were also not altered by any concentration of OT in either group, one exception was a hyperpolarization of threshold by 600 nM OT after CIH (example in Fig 3D, inset; Bsl, −19.6 ± 5.3 vs. OT, −24.0 ± 5.8 mV, paired t-test, p = 0.04, N=3, n=5). Together, these results show that after CIH OT depolarized Vm and modified AP discharge.

3.4. OTR activation after CIH reduces total transmembrane outward potassium currents (IK).

Potassium currents influence membrane potential, neuronal excitability and AP properties (Glazebrook et al., 2002; Pongs, 1999). As such, nodose neuron IK was examined after Norm and CIH, and the influence of OT on these currents was examined. In Norm and CIH-exposed neurons, depolarizing voltage steps (140 ms, 10 pA steps) induced progressively greater currents, as shown in Fig 4A&C. OT at 600 nM appeared to decrease IK in CIH but not Norm-exposed neurons. We quantified the current-voltage (IV) activation profile of the early and steady state currents. Early and steady state IK was not altered by Veh application in either group (Norm, N=3, n=7; CIH, N=4, n=7; not shown). In Norm-exposed neurons, 300 and 600 nM OT (N=4 & 4, n=8 & 9) had minimal effect on IK (600 nM shown in Fig 4B). By contrast, after CIH 300 and 600 nM OT (N= 4 & 3, n=7 & 6) produced a progressively greater decrease in both the early and steady state currents. Quantitatively, 600 nM OT decreased the early and steady state IK in CIH-exposed neurons (Fig 4D, two-way RM ANOVA). The OT-sensitive current, as shown in the representative example in Fig 4E was greater after CIH compared to Norm (Fig 4F). Taken together, these results suggest CIH enhances the responsiveness of OT to influence potassium currents.

Figure 4. Following CIH OT decreases transmembrane voltage-gated potassium currents (IK).

Figure 4.

Open in a new tab

A, C. Representative Ik from a nodose neuron during baseline (Bsl) and 600 nM OT (OT) in a Norm- (A) and CIH- (C) exposed neuron. Capacitive currents reduced for clarity. Scale, 1000 pA and 100 ms. Shaded areas denote time in which early and steady state currents were measured. Note the reduction in IK by OT after CIH. B,D. Early current (e in panel A) and steady state (ss in panel A) were not altered by 600 nM OT in Norm (B) neurons, but were reduced significantly after CIH (D). E. Representative OT-sensitive IK of Norm and CIH neurons after 600 nM OT exposure. Scale, 250 pA and 100 ms. Note the increase in CIH, which is quantified in F. *, p < 0.05 vs Bsl (B,D) or Norm vs CIH (E), Two-way RM ANOVA with Holm-Sidak (B,D) or Fisher’s LSD (E). Bsl was physiological solution.

OT may bind to arginine vasopressin (AVP) receptors (Chini et al., 2008). To confirm the enhanced CIH responsiveness of IK was due to activation of the OT receptor rather than AVP receptor, we utilized two approaches. Application of the OTR antagonist L-368,899 (OTR-x, 1 μM, N=3, n=7) prior to 600 nM OT prevented the decrease in IK (Fig 5A example, B). In a separate set of neurons, application of the OTR agonist (TGOT, 600 nM, N=3, n=4) which has a higher affinity for OT than AVP receptor (Chini et al., 2008), decreased IK in CIH neurons (Fig 5C) mimicking the response to OT. In Norm-exposed neurons, neither OTR-x + OT nor TGOT effected IK (not shown). Thus, in the nodose ganglia OT acts via the oxytocin receptor.

Figure 5. OTR antagonist (OTR-x) prevents the decrease in IK after CIH, and OTR agonist decreased IK.

Figure 5.

Open in a new tab

A. Representative IK of a nodose neuron after CIH exposed to 1 μM OTR-x and to OTR-x combined with 600 nM OT. Note that the OT induced decrease in IK seen in Fig 3 is absent during OTR block. Scale, 1000 pA and 100 ms. B. Quantification demonstrating OTR-x blocked the steady state decrease of steady state IK in response to 600 nM OT. C. An OT agonist (600 nM TGOT) decreased steady state IK in CIH neurons. *, p < 0.05 vs Bsl, Two-way RM ANOVA with Holm-Sidak. Bsl was physiological solution.

3.5. OT alters decreases IK independent of other currents via TEA-sensitive channels.

OT alters calcium channels in the mouse nodose ganglia, as well as a multitude of channels in the central nervous system (Breton et al., 2009; Raggenbass et al., 1992; van den Burg et al., 2015). To provide greater focus on the IK channels without the potential influence of other channels we utilized an extracellular solution utilizing the general sodium channel blocker NMDG and lowered calcium to remove the influence of calcium channels and calcium-activated IK. The latter have been shown to be voltage dependent (Schild et al., 2012). As shown in the CIH example in Fig 6A, OT decreased IK in NMDG solution as in physiological-like solutions shown in Fig 3. Across all neurons studied, Veh application did not alter steady state IK in Norm (N=7, n=9)- or CIH (N=4, n=5)-exposed cells. OT applied at 300 nM decreased IK in CIH neurons at voltages between 40 and 60 mV (N=6, n=8, not shown), whereas there was no effect in Norm neurons (N=5, n=7). Increasing OT to 600 nM in Norm neurons reduced IK only at the highest voltage (Fig 6B) whereas there was a continued pronounced effect in CIH neurons (Fig 6C, N=8, n=12). Examination of the OT-sensitive currents confirm a greater sensitivity in CIH neurons (Fig 6D).

Figure 6. CIH neurons in the presence of N-Methyl-D-glucamine (NMDG) are more responsive to OT.

Figure 6.

Open in a new tab

A. Representative IK of a CIH NPG neuron after exposure to 600 nM OT in the presence of NMDG solution to reduce sodium and calcium currents. Scale, 1000 pA and 100 ms. B, C. 600 nM OT reduced early and steady state IK only at high voltage steps in Norm neurons (B) but had a more prominent response after CIH (C). D. OT-sensitive currents are greater in CIH neurons exposed to 600 nM OT. *, p < 0.05 vs Bsl, Two-way RM ANOVA with Holm-Sidak (B–C) or Fisher’s LSD (D). Bsl consisted of NMDG solution.

In CIH-exposed neurons, we examined the pharmacological profile of OT-sensitive currents. We utilized the general IK blockers 4-aminopuridine (4-AP, 5 mM) that inhibits early transient currents and tetraethylammonium (TEA, 10 mM) that inhibits the delayed rectifier current. 4-AP did not inhibit the reduction of the early or steady state IK by OT (600 nM, Fig 7A, N=4, n=4). On the other hand, in the presence of TEA OT failed to effect IK in CIH neurons (Fig 7B, N=4, n=7).

Figure 7. Characterization of CIH OT-sensitive currents.

Figure 7.

Open in a new tab

A. OT (600 nM) reduced the early and steady state IK in the presence of 4-aminopuridine (4-AP). B. Tetraethylammonium (TEA) prevented the 600 nM OT-reduction in early or steady state currents. *, p < 0.05 vs Bsl, Two-way RM ANOVA with Holm-Sidak. Bsl contained either 4-AP or TEA in NMDG solution.

Across all potassium currents studied, when examined in normal physiological solutions prior to any further manipulation, the current-voltage relationship was similar between Norm and CIH (n = 79 & 72, respectively, supplemental figure 1).

4. DISCUSSSION

In this study, we present several major findings. Specifically, we show in nodose neurons that OTR expression is not altered following CIH yet its activation depolarizes membrane potential, attenuates action potential discharge, and decreases potassium currents. These results provide mechanistic insight into the potential contribution of OT in autonomic (dys)function via sensory afferents in patients with obstructive sleep apnea.

Periods of low systemic hypoxia increases circulating OT as well as activation of OT projections to the nTS that originate from the hypothalamus (Goraca, 2004; Kelestimur et al., 1997; Ruyle et al., 2018; Stegner et al., 1984). Vagal afferent neurons are initially integrated and modulated in the brainstem nTS (Andresen et al., 1994). Thus, increased hypoxia-induced OT may alter vagal afferent activity at the level of the soma in the nodose ganglia or their central terminals. Examining OT plasma immediately following the last hypoxic episode on the 10th day of CIH exposure illustrated an increase in OT compared to normoxic controls, consistent with Wilson et al. (2018). By contrast, OT plasma in CIH and Norm rats 12–16 hours after the last hypoxia period (a period in which we took tissue and recorded) was comparable. This is not unexpected as the half-life of OT in the plasma is several minutes (Leng et al., 2016; Veening et al., 2010). Further studies will need to examine after CIH cerebrospinal OT, which is maintained longer than in plasma (Veening et al., 2010). We confirm the existence of OTR transcript and protein in the normoxic nodose ganglion (Welch et al., 2009) which was localized to somas and fibers that may terminate in the nTS. The later expression may be responsible for the potentiating effects of OT in afferent glutamate release in the nTS (Peters et al., 2008). We extend these findings to show that after CIH, OTR protein remains yet is functionally exaggerated. The mechanism for this functional augmentation is likely multifaceted but could include increased receptor sensitivity, affinity or second messenger pathways. The nodose is a heterogenous ganglia (Kupari et al., 2019), and thus one limitation of this study is we do not know the modality (i.e., respiratory, gastrointestinal or cardiovascular), cell type (A- or C-fibers) or nTS projection site of these OTR-positive neurons. In addition, our cultures may also contain neurons from the adjacent petrosal ganglion. Physiologically, our study is in agreement with others that 7–10 days CIH increases MAP and HR (Yamamoto et al., 2013; Zoccal et al., 2007; Zoccal et al., 2009) but does not alter baroreflex sensitivity (BRS) in conscious rats, as determined by the sequence method (Yamamoto et al., 2013; Zoccal et al., 2007). Longer durations of CIH may decrease BRS (Gu et al., 2007; Lai et al., 2006) and the lack of altered sBRS in the present work may be due, in part, to the central synaptic plasticity demonstrated in the nTS (Kline et al., 2007). Alternatively, while CIH may decrease BRS, microinjection of OT into the nTS enhances BRS (Higa et al., 2002); one may speculate that increased OT may help offset the decrease in BRS during CIH although this requires additional investigation. While our studied neurons may contribute to these cardiovascular alterations and pathways, the expression, target sites and physiological implications of specific OTR positive neurons requires further study.

In isolated neurons following CIH, nanomolar OT has a concentration-related effect of membrane potential and potassium currents. These responses are likely due to OT acting on OTRs rather than vasopressin receptors, as suggested by the block of OT current responses by an OTR antagonist and the mimicked decrease in IK by an OT agonist. The OT-induced depolarization may be due to OT acting on one or more ion channels expressed on vagal neurons (Schild et al., 2005). For instance, in other neurons OT increases Ca2+ in part via TRPV2 channels (van den Burg et al., 2015), activates a persistent sodium current in dorsal vagal neurons (Raggenbass et al., 1992) and decreases potassium currents in the spinal cord (Breton et al., 2009). The depolarization of membrane potential by OT at negative voltages (~ −50 mV) may also suggest inhibition of potassium leak channels. Thus, OT via its G protein coupled receptor may modify a number of ion channels resulting in depolarization. Our current study shows that OT also attenuates voltage-gated IK in sensory neurons, and this occurs primarily after CIH. The hyperpolarization of AP threshold by OT, which is within the range of IK activation, is consistent with OT subtly altering IK to allow easier Vm depolarization and discharge (Bean, 2007; Schild et al., 2005; Schild et al., 2012). However, the reduction in discharge by OT is not entirely consistent with OT working solely through IK as its reduction should allow for greater discharge rates (Pongs, 1999). Rather, it is likely due to an OT effect on the available compliment of the ion channels.

Nodose neurons have a multitude of voltage-gated K+ currents (Schild et al., 2005; Schild et al., 2012). We subsequently utilized a pharmacological approach to distinguish some of the currents that may be affected. Our data shows that 4-AP did not prevent the OT-attenuation of IK. 4-AP is a prototypical blocker of currents with fast transient (IA) currents. At the utilized concentration of 4-AP, the continued influence of OT suggests it does not influence members of the KV1.4 and KV4 families that often mediate IA (Rudy, 2010). On the other hand, inclusion of TEA, a ubiquitous voltage-gated potassium channel blocker, eliminated the influence of OT on IK. Seeing as OT continued to decrease both the early and steady state currents in the presence of 4-AP, and are TEA-sensitive, OT may alter a number of channel families including members of the KV1 and KV2 families (Glazebrook et al., 2002; Rudy, 2010). Regardless of these results, additional studies are required using more family specific antagonists or toxins as well as examination of their current properties (i.e., inactivation, recovery) to fully identify the channel altered by OT. Nevertheless, the influence of OT on potassium currents is consistent with its effects on KV channel in the spinal cord (Breton et al., 2009) and inward rectifiers in the olfactory bulb (Gravati et al., 2010).

5. CONCLUSION.

In conclusion, we show that CIH enhances the membrane voltage and potassium current responses to oxytocin. This occurs in the absence of changes in the OTR expression. Given the multi-modalities that vagal nodose neurons serve, it is likely that OT modulates sensory input to the brainstem from a variety of viscera to ultimately effect end-organ function.

Supplementary Material

1

Supplemental Figure 1. Potassium currents (IK) are not altered after CIH. Across all neurons studied, IK was similar between Norm (n=79) and CIH (n=72). Two-way ANOVA.

HIGHLIGHTS.

  • Oxytocin receptors are localized to nodose ganglia

  • After CIH, exogenous oxytocin depolarizes nodose neurons

  • Oxytocin decreases voltage-gated potassium currents following CIH

  • The decrease in potassium currents is prevented by the blocker TEA

ACKNOWLEDGEMENTS

We thank Drs. Diana Martinez and Gabrielle Hofmann for critical reading of drafts of the manuscript.

FUNDING

This study was supported by HL128454 & HL 098602 (DDK)

ABBREVIATIONS

4-AP

4-Aminopyridine

AP

Action potential

CIH

Chronic intermittent hypoxia

IK

Potassium Current

Norm

Normoxia

OT

Oxytocin

OTR

Oxytocin receptor

TEA

Tetraethylammonium

Vm

Membrane potential

Footnotes

Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Conflict of Interest: The authors declare no competing financial interests.

REFERENCES

  1. Almado CE, Machado BH, Leao RM 2012. Chronic intermittent hypoxia depresses afferent neurotransmission in NTS neurons by a reduction in the number of active synapses. J Neurosci 32, 16736–16746. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Andresen MC, Kunze DL 1994. Nucleus tractus solitarius--gateway to neural circulatory control. Annu. Rev. Physiol 56, 93–116. [DOI] [PubMed] [Google Scholar]
  3. Bean BP 2007. The action potential in mammalian central neurons. Nat Rev Neurosci 8, 451–465. [DOI] [PubMed] [Google Scholar]
  4. Blevins JE, Eakin TJ, Murphy JA, Schwartz MW, Baskin DG 2003. Oxytocin innervation of caudal brainstem nuclei activated by cholecystokinin. Brain Res 993, 30–41. [DOI] [PubMed] [Google Scholar]
  5. Breton JD, Poisbeau P, Darbon P 2009. Antinociceptive action of oxytocin involves inhibition of potassium channel currents in lamina II neurons of the rat spinal cord. Mol Pain 5, 63. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Chini B, Manning M, Guillon G 2008. Affinity and efficacy of selective agonists and antagonists for vasopressin and oxytocin receptors: an “easy guide” to receptor pharmacology. Prog Brain Res 170, 513–517. [DOI] [PubMed] [Google Scholar]
  7. Gimpl G, Fahrenholz F 2001. The oxytocin receptor system: structure, function, and regulation. Physiol Rev 81, 629–683. [DOI] [PubMed] [Google Scholar]
  8. Glazebrook PA, Ramirez AN, Schild JH, Shieh C-C, Doan T, Wible BA, Kunze DL 2002. Potassium channels Kv1.1, Kv1.2 and Kv1.6 influence excitability of rat visceral sensory neurons. The Journal of Physiology 541, 467–482. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Gong L, Gao F, Li J, Li J, Yu X, Ma X, Zheng W, Cui S, Liu K, Zhang M, Kunze W, Liu CY 2015. Oxytocin-induced membrane hyperpolarization in pain-sensitive dorsal root ganglia neurons mediated by Ca(2+)/nNOS/NO/KATP pathway. Neuroscience 289, 417–428. [DOI] [PubMed] [Google Scholar]
  10. Gonzalez-Iglesias AE, Fletcher PA, Arias-Cristancho JA, Cristancho-Gordo R, Helena CV, Bertram R, Tabak J 2015. Direct stimulatory effects of oxytocin in female rat gonadotrophs and somatotrophs in vitro: comparison with lactotrophs. Endocrinology 156, 600–612. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Goraca A 2004. Oxytocin content in the venous blood outflowing from the vicinity of the cavernous sinus and from the femoral vein. Endocr Regul 38, 119–125. [PubMed] [Google Scholar]
  12. Gravati M, Busnelli M, Bulgheroni E, Reversi A, Spaiardi P, Parenti M, Toselli M, Chini B 2010. Dual modulation of inward rectifier potassium currents in olfactory neuronal cells by promiscuous G protein coupling of the oxytocin receptor. J Neurochem 114, 1424–1435. [DOI] [PubMed] [Google Scholar]
  13. Gross PM, Wall KM, Pang JJ, Shaver SW, Wainman DS 1990. Microvascular specializations promoting rapid interstitial solute dispersion in nucleus tractus solitarius. Am J Physiol 259, R1131–1138. [DOI] [PubMed] [Google Scholar]
  14. Gu H, Lin M, Liu J, Gozal D, Scrogin KE, Wurster R, Chapleau MW, Ma X, Cheng ZJ 2007. Selective impairment of central mediation of baroreflex in anesthetized young adult Fischer 344 rats after chronic intermittent hypoxia. Am J Physiol Heart Circ. Physiol 293, H2809–H2818. [DOI] [PubMed] [Google Scholar]
  15. Han RT, Kim HB, Kim YB, Choi K, Park GY, Lee PR, Lee J, Kim HY, Park CK, Kang Y, Oh SB, Na HS 2018. Oxytocin produces thermal analgesia via vasopressin-1a receptor by modulating TRPV1 and potassium conductance in the dorsal root ganglion neurons. Korean J Physiol Pharmacol 22, 173–182. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Henry JL, Sessle BJ 1989. Vasopressin and oxytocin express excitatory effects on respiratory and respiration-related neurones in the nuclei of the tractus solitarius in the cat. Brain Res 491, 150–155. [DOI] [PubMed] [Google Scholar]
  17. Higa KT, Mori E, Viana FF, Morris M, Michelini LC 2002. Baroreflex control of heart rate by oxytocin in the solitary-vagal complex. Am J Physiol Regul Integr Comp Physiol 282, R537–545. [DOI] [PubMed] [Google Scholar]
  18. Iwasaki Y, Maejima Y, Suyama S, Yoshida M, Arai T, Katsurada K, Kumari P, Nakabayashi H, Kakei M, Yada T 2015. Peripheral oxytocin activates vagal afferent neurons to suppress feeding in normal and leptin-resistant mice: a route for ameliorating hyperphagia and obesity. Am J Physiol Regul Integr Comp Physiol 308, R360–369. [DOI] [PubMed] [Google Scholar]
  19. Jameson H, Bateman R, Byrne P, Dyavanapalli J, Wang X, Jain V, Mendelowitz D 2016. Oxytocin neuron activation prevents hypertension that occurs with chronic intermittent hypoxia/hypercapnia in rats. Am J Physiol Heart Circ Physiol 310, H1549–1557. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Kelestimur H, Leach RM, Ward JP, Forsling ML 1997. Vasopressin and oxytocin release during prolonged environmental hypoxia in the rat. Thorax 52, 84–88. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Kline DD, Hendricks G, Hermann G, Rogers RC, Kunze DL 2009. Dopamine inhibits N-type channels in visceral afferents to reduce synaptic transmitter release under normoxic and chronic intermittent hypoxic conditions. J Neurophysiol 101, 2270–2278. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Kline DD, Ramirez-Navarro A, Kunze DL 2007. Adaptive depression in synaptic transmission in the nucleus of the solitary tract after in vivo chronic intermittent hypoxia: evidence for homeostatic plasticity. J. Neurosci 27, 4663–4673. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Kline DD, Wang S, Kunze DL 2019. TRPV1 channels contribute to spontaneous glutamate release in nucleus tractus solitarii following chronic intermittent hypoxia. J Neurophysiol 121, 881–892. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Kunze DL, Kline DD, Ramirez-Navarro A 2007. Hypertension in sleep apnea: The role of the sympathetic pathway. Cleveland Clinic Journal of Medicine 74, S34–S36. [DOI] [PubMed] [Google Scholar]
  25. Kupari J, Haring M, Agirre E, Castelo-Branco G, Ernfors P 2019. An Atlas of Vagal Sensory Neurons and Their Molecular Specialization. Cell Rep 27, 2508–2523 e2504. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Lacolley P, Owen JR, Sandock K, Lewis TH, Bates JN, Robertson TP, Lewis SJ 2006. Occipital artery injections of 5-HT may directly activate the cell bodies of vagal and glossopharyngeal afferent cell bodies in the rat. Neuroscience 143, 289–308. [DOI] [PubMed] [Google Scholar]
  27. Lai CJ, Yang CC, Hsu YY, Lin YN, Kuo TB 2006. Enhanced sympathetic outflow and decreased baroreflex sensitivity are associated with intermittent hypoxia-induced systemic hypertension in conscious rats. J Appl Physiol (1985) 100, 1974–1982. [DOI] [PubMed] [Google Scholar]
  28. Leng G, Sabatier N 2016. Measuring Oxytocin and Vasopressin: Bioassays, Immunoassays and Random Numbers. J Neuroendocrinol 28. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Lin M, Liu R, Gozal D, Wead WB, Chapleau MW, Wurster R, Cheng ZJ 2007. Chronic intermittent hypoxia impairs baroreflex control of heart rate but enhances heart rate responses to vagal efferent stimulation in anesthetized mice. Am. J. Physiol Heart Circ. Physiol 293, H997–1006. [DOI] [PubMed] [Google Scholar]
  30. Maley BE 1996. Immunohistochemical localization of neuropeptides and neurotransmitters in the nucleus solitarius. Chem Senses 21, 367–376. [DOI] [PubMed] [Google Scholar]
  31. Maolood N, Meister B 2009. Protein components of the blood-brain barrier (BBB) in the brainstem area postrema-nucleus tractus solitarius region. J Chem Neuroanat 37, 182–195. [DOI] [PubMed] [Google Scholar]
  32. Martinez D, Rogers RC, Hasser EM, Hermann GE, Kline DD 2020. Loss of excitatory amino acid transporter restraint following chronic intermittent hypoxia contributes to synaptic alterations in nucleus tractus solitarii. J Neurophysiol 123, 2122–2135. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Matott MP, Hasser EM, Kline DD 2020. Sustained Hypoxia Alters nTS Glutamatergic Signaling and Expression and Function of Excitatory Amino Acid Transporters. Neuroscience 430, 131–140. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Michelini LC 2001. Oxytocin in the NTS. A new modulator of cardiovascular control during exercise. Ann N Y Acad Sci 940, 206–220. [PubMed] [Google Scholar]
  35. Michelini LC 2007. Differential effects of vasopressinergic and oxytocinergic pre-autonomic neurons on circulatory control: reflex mechanisms and changes during exercise. Clin Exp Pharmacol Physiol 34, 369–376. [DOI] [PubMed] [Google Scholar]
  36. Moak JP, Kunze DL 1993. Potassium currents of neurons isolated from medical nucleus tractus solitarius. Am. J. Physiol 265, H1596–H1602. [DOI] [PubMed] [Google Scholar]
  37. Peters JH, McDougall SJ, Kellett DO, Jordan D, Llewellyn-Smith IJ, Andresen MC 2008. Oxytocin enhances cranial visceral afferent synaptic transmission to the solitary tract nucleus. J Neurosci 28, 11731–11740. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Pongs O 1999. Voltage-gated potassium channels: from hyperexcitability to excitement. FEBS Lett 452, 31–35. [DOI] [PubMed] [Google Scholar]
  39. Prabhakar NR, Kline DD 2002. Ventilatory changes during intermittent hypoxia: importance of pattern and duration. High Alt. Med. Biol 3, 195–204. [DOI] [PubMed] [Google Scholar]
  40. Raggenbass M, Dreifuss JJ 1992. Mechanism of action of oxytocin in rat vagal neurones: induction of a sustained sodium-dependent current. J Physiol 457, 131–142. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Raggenbass M, Tribollet E, Dubois-Dauphin M, Dreifuss JJ 1989. Vasopressin receptors of the vasopressor (V1) type in the nucleus of the solitary tract of the rat mediate direct neuronal excitation. J Neurosci 9, 3929–3936. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Ramirez-Navarro A, Glazebrook PA, Kane-Sutton M, Padro C, Kline DD, Kunze DL 2011. Kv1.3 channels regulate synaptic transmission in the nucleus of solitary tract. J. Neurophysiol 105, 2772–2780. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Rudy B 2010. Ion Channels: From Structure to Function. In: Kew JNC, Davies CH, (Eds.). Oxford University Press. [Google Scholar]
  44. Ruyle BC, Klutho PJ, Baines CP, Heesch CM, Hasser EM 2018. Hypoxia activates a neuropeptidergic pathway from the paraventricular nucleus of the hypothalamus to the nucleus tractus solitarii. Am J Physiol Regul Integr Comp Physiol 315, R1167–R1182. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Schild JH, Alfrey KD, Li BY 2005. Voltage-gated ion channels in vagal afferent neurons In: Undem BJ, Weinreich D, (Eds.), Advances in Vagal Afferent Neurobiology. , 1 ed. CRC Press, Boca Raton: pp. 77–99. [Google Scholar]
  46. Schild JH, Kunze DL 2012. Differential distribution of voltage-gated channels in myelinated and unmyelinated baroreceptor afferents. Auton Neurosci 172, 4–12. [DOI] [PubMed] [Google Scholar]
  47. Sofroniew MV, Schrell U 1980. Hypothalamic neurons projecting to the rat caudal medulla oblongata, examined by immunoperoxidase staining of retrogradely transported horseradish peroxidase. Neurosci Lett 19, 257–263. [DOI] [PubMed] [Google Scholar]
  48. Son SJ, Filosa JA, Potapenko ES, Biancardi VC, Zheng H, Patel KP, Tobin VA, Ludwig M, Stern JE 2013. Dendritic peptide release mediates interpopulation crosstalk between neurosecretory and preautonomic networks. Neuron 78, 1036–1049. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Stegner H, Leake RD, Palmer SM, Oakes G, Fisher DA 1984. The effect of hypoxia on neurohypophyseal hormone release in fetal and maternal sheep. Pediatr Res 18, 188–191. [DOI] [PubMed] [Google Scholar]
  50. van den Burg EH, Stindl J, Grund T, Neumann ID, Strauss O 2015. Oxytocin Stimulates Extracellular Ca2+ Influx Through TRPV2 Channels in Hypothalamic Neurons to Exert Its Anxiolytic Effects. Neuropsychopharmacology 40, 2938–2947. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Veening JG, de Jong T, Barendregt HP 2010. Oxytocin-messages via the cerebrospinal fluid: behavioral effects; a review. Physiol Behav 101, 193–210. [DOI] [PubMed] [Google Scholar]
  52. Voorn P, Buijs RM 1983. An immuno-electronmicroscopical study comparing vasopressin, oxytocin, substance P and enkephalin containing nerve terminals in the nucleus of the solitary tract of the rat. Brain Res 270, 169–173. [DOI] [PubMed] [Google Scholar]
  53. Weiss JW, Liu MD, Huang J 2007. Physiological basis for a causal relationship of obstructive sleep apnoea to hypertension. Exp Physiol 92, 21–26. [DOI] [PubMed] [Google Scholar]
  54. Welch MG, Tamir H, Gross KJ, Chen J, Anwar M, Gershon MD 2009. Expression and developmental regulation of oxytocin (OT) and oxytocin receptors (OTR) in the enteric nervous system (ENS) and intestinal epithelium. J Comp Neurol 512, 256–270. [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. Wilson EN, Anderson M, Snyder B, Duong P, Trieu J, Schreihofer DA, Cunningham RL 2018. Chronic intermittent hypoxia induces hormonal and male sexual behavioral changes: Hypoxia as an advancer of aging. Physiol Behav 189, 64–73. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Yamamoto K, Eubank W, Franzke M, Mifflin S 2013. Resetting of the sympathetic baroreflex is associated with the onset of hypertension during chronic intermittent hypoxia. Auton Neurosci 173, 22–27. [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Zoccal DB, Bonagamba LG, Oliveira FR, Antunes-Rodrigues J, Machado BH 2007. Increased sympathetic activity in rats submitted to chronic intermittent hypoxia. Exp Physiol 92, 79–85. [DOI] [PubMed] [Google Scholar]
  58. Zoccal DB, Bonagamba LG, Paton JF, Machado BH 2009. Sympathetic-mediated hypertension of awake juvenile rats submitted to chronic intermittent hypoxia is not linked to baroreflex dysfunction. Exp Physiol 94, 972–983. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

1

Supplemental Figure 1. Potassium currents (IK) are not altered after CIH. Across all neurons studied, IK was similar between Norm (n=79) and CIH (n=72). Two-way ANOVA.

NIHMS1635282-supplement-1.pdf (382.1KB, pdf)

RESOURCES