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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
J Neurosci. Author manuscript; available in PMC 2010 June 15.
Published in final edited form as:
PMCID: PMC2885697

Chronic Sustained Hypoxia Enhances Both Evoked EPSCs and Norepinephrine Inhibition of Glutamatergic Afferent Inputs in the Nucleus of the Solitary Tract


The nucleus of the solitary tract (NTS) receives inputs from both arterial chemoreceptors and central noradrenergic neural structures activated during hypoxia. We investigated NE modulation of chemoreceptor afferent integration following a chronic exposure to sustained hypoxia (CSH; 7-8 days at 10% FIO2). Whole-cell recordings of NTS second-order neurons identified by DiA labeling of carotid bodies were obtained in a brain slice. Electrical stimulation of the solitary tract was used to evoke EPSCs (eEPSCs). CSH exposure increased eEPSC amplitude via both pre- and post-synaptic mechanisms. NE dose-dependently decreased the amplitude of eEPSCs. NE increased the paired-pulse ratio of eEPSCs and reduced the frequency of miniature EPSCs, suggesting a pre-synaptic mechanism. EC50 of NE inhibition of eEPSCs was lower in CSH cells (3.0±0.9 μM, n=5) than in normoxic (NORM) cells (7.6±1.0 μM, n=7, p<0.01). NE (10μM) elicited greater inhibition of eEPSCs in CSH cells (63±2%, n=16) than NORM cells (45±3%, n=21, p<0.01). The α-adrenoreceptor antagonist phentolamine abolished NE inhibition of eEPSCs. CSH enhanced α2-adrenoreceptor agonist clonidine-mediated inhibition (3 μM, NORM 23±2%, n=5 vs. CSH 44±5%, n=4, p<0.05), but attenuated α1-adrenoreceptor agonist phenylephrine-mediated inhibition (40 μM, NORM 36±2%, n=11 vs. CSH 26±4%, n=6, p<0.05). The α2-adrenoreceptor antagonist yohimbine abolished CSH-induced enhancement of NE inhibition of eEPSCs. These results demonstrate that CSH increases evoked excitatory inputs to NTS neurons receiving arterial chemoreceptor inputs. CSH also enhances NE inhibition of glutamate release from inputs to these neurons via pre-synaptic α2-adrenoreceptors. These changes represent central neuronal adaptations to CSH.

Keywords: chemoreflex, chemoreceptor, electrophysiology, respiration


During many pathophysiological conditions, periods of general or focal brain ischemia/hypoxia occur which can result in neuronal injury and interruption of synaptic transmission as well as a dramatic change in the extracellular concentration of various neurochemicals (Lipton, 1999). In particular, excessive extracellular glutamate may have an excitotoxic effect depending on the severity of the tissue hypoxia. At the same time, other neurochemicals are released to counteract the toxic glutamate effect by depressing excitatory synaptic transmission and reducing glutamate release. One such neurochemical is norepinephrine (NE) and NE levels increase dramatically in mammalian brain and spinal cord during hypoxia (Globus et al., 1989; Nakai et al., 1999; Perego et al., 1992).

A unique scenario occurs in the central pathways that mediate arterial chemoreflexes. These neural structures experience tissue hypoxia during acute or chronic hypoxia. However, synaptic transmission of chemoreceptor afferent inputs has to be sustained to maintain the reflex response to hypoxia, i.e. increased sympathetic nerve activity and respiratory activity (Guyenet, 2000). The nucleus of the solitary tract (NTS) receives the first central projections of the arterial chemoreceptors and is the first central integration site in arterial chemoreflex pathways (Loewy, 1990; Mifflin, 1992). The NTS contains its own noradrenergic neurons the A2 cell group, and also receives inputs from multiple central noradrenergic structures (Loewy, 1990).

The α1-, α2- and β-adrenoreceptor subtypes have been detected in the NTS (Aoki et al., 1989; Dashwood et al., 1985; Day et al., 1997; Jones et al., 1985; Young and Kuhar, 1980). Microinjection of NE into the NTS attenuated cardiovascular responses in both arterial chemoreflex and arterial baroreflex (De Jong, 1974; Silva de Oliveira et al., 2007). Further evidence suggests that activation of both α1- and α2-adrenoreceptors has inhibitory effect in the NTS (Feldman and Moises, 1987; Feldman and Moises, 1988; Feldman and Felder, 1989a, 1989b; Moore and Guyenet, 1983; Zhang and Mifflin, 2007), although the contribution of different adrenoreceptor subtypes is still unknown.

Chronic sustained hypoxia (CSH) is a pathophysiological situation associated with chronic pulmonary and heart disease and travel to high altitude. CSH alters the neural control of cardio-respiratory activity (Powell et al., 2000; Prabhakar and Jacono, 2005). Whether CSH induces adaptive changes in synaptic integration of arterial chemoreceptor inputs in the NTS has not been investigated. We report that CSH enhances glutamatergic synaptic transmission of arterial chemoreceptor inputs in the NTS via pre- and post-synaptic mechanisms. In addition, CSH has been shown to induce significant changes in NE metabolism in the NTS and in other noradrenergic structures that project to the NTS (Schmitt et al., 1994; Soulier et al., 1992). Whether CSH alters NE modulation of synaptic transmission within the NTS is currently unknown. Our results show CSH enhances NE, α2-adrenoreceptor inhibition of glutamate release within the NTS.


All experimental protocols were approved by the Institutional Animal Care and Use Committee at the University of Texas Health Science Center at San Antonio.

Surgical preparation for labeling carotid body

All experiments were performed on male Sprague-Dawley rats (125–145 g, Charles River Laboratories, Wilmington, MA). Rats were anaesthetized with a combination of ketamine (75 mg/kg, I.P., Fort Dodge) and medetomidine (0.5 mg/kg, I.P., Pfizer). Crystals of anterograde fluorescent dye 1,1′-dilinoleyl-3,3,3′,3′-tetra-methylindocarbocyanine, 4-chlorobenzenesulphonate (DiA) were applied unilaterally to the carotid body region to visualize the chemoreceptor synaptic terminals in the NTS and neurons receiving these synaptic contacts (de Paula et al., 2007; Tolstykh et al., 2004; Zhang and Mifflin, 2007; Zhang et al., 2008). The area was then embedded with silicone adhesive (Kwik-Sil, WPI, Sarasota, FL). Anesthesia was terminated by atipamezole (1 mg/kg I.P., Pfizer) and post-operative analgesics (nubaine, I.M.) were available.

Chronic hypoxia exposure

Rats were divided into two groups: chronic hypoxia (CSH) group and normoxic control (NORM) group. CSH rats were housed in their home cages within a normobaric hypoxia chamber at oxygen levels of 10 ± 0.5% for 7-8 days as described previously (Ilyinsky and Mifflin, 2005; Ilyinsky et al., 2003; Tolstykh et al., 2004; Zhang et al., 2008). The air within the chamber was recycled and the oxygen level was controlled by a computer-driven set of valves and pumps. A hypoxic environment was achieved by the addition of nitrogen gas to room air. Oxygen levels within the chamber were monitored with an electrochemical sensor, and this information fed into the computer-driven feedback circuit so that deviations in the oxygen level from preset value were rapidly corrected by adding appropriate gas. Temperature and humidity were monitored, and the recycled air was passed through a desiccant and CO2 scrubber. NORM rats were maintained in a similar environment while breathing room air.

Brain slice preparation

Anesthesia was induced with isoflurane and the brainstem rapidly removed and placed in ice-cold, high-sucrose, artificial cerebrospinal fluid (aCSF) that contained (in mM): 3 KCl, 1 MgCl2, 1 CaCl2, 2 MgSO4, 1.25 NaH2PO4, 26 NaHCO3, 10 glucose, and 206 sucrose, pH 7.4 when continuously bubbled with 95% O2/5% CO2. The brainstem was mounted in a vibrating microtome (VT1000E, Leica Microsystems, Bannockburn, IL) and horizontal slices (250 μm thickness) were cut with a sapphire knife (Delaware Diamond Knives, Wilmington, DE). The slices were incubated for at least one hour in normal aCSF that contained (in mM): 124 NaCl, 3 KCl, 2 MgSO4, 1.25 NaH2PO4, 26 NaHCO3, 10 glucose and 2 CaCl2, pH 7.4 when continuously bubbled with 95% O2/5% CO2.

Electrophysiological Recording

Whole-cell patch-clamp recordings were performed in the recording chamber on an upright epifluorescent microscope (Olympus BX50WI, Tokyo) equipped with infrared differential interference contrast (IR-DIC) and an optical filter set for visualization of DiA. The slice was held in place with a nylon mesh, submerged in normal aCSF equilibrated with 95% O2/5% CO2 and perfused at a rate of 2-3 ml/min. All images were captured with a charge-coupled device (CCD) camera (IR-1000, CCD-100; Dage-MTI, Michigan City, IN) displayed on a TV monitor and stored in a PC computer. Patch pipettes were pulled from borosilicate glass capillaries with an inner filament (0.90 mm ID, 1.2 mm OD, WPI, Sarasota, FL) on a pipette puller (Model P-2000, Sutter Instrument Company, Novato, CA) and were filled with a solution that contained (in mM): 145 K-gluconate, l MgCl2, 10 HEPEs, 1.1 EGTA, 2 Mg2ATP, and 0.3 Na3GTP. The pH was adjusted to 7.3 with KOH. The pipette resistance ranged from 3 to 6 MΩ. A seal resistance of at least 1 GΩ or above, and an access resistance < 20 MΩ which changed <15% during recording were considered acceptable. Series resistance was optimally compensated. Recordings of post-synaptic currents began 5 min after the whole cell access was established and the holding current reached a steady state. Recordings were made with an AxoPatch 200B patch-clamp amplifier and pClamp software version 8 (Axon Instruments, Union City, CA). Whole-cell currents were filtered at 2 kHz, digitized at 10 kHz with the DigiData 1200 Interface (Axon Instruments) and stored in a PC computer for offline analysis. All experiments were performed at room temperature.

Whole-cell voltage-clamp recordings were performed on second-order NTS arterial chemoreceptor neurons labeled with fluorescent DiA (Fig. 1A). Cells were clamped at a membrane potential of −60 mV. Input resistance was monitored by frequently applying a 10 mV hyperpolarizing voltage step (100 ms duration) from a holding potential of −60 mV. Evoked EPSCs (eEPSCs) were elicited by electrical stimulation of the ipsilateral solitary tract (ST) using a concentric bipolar electrodes (FHC, Bowdoinham, ME) with a tip diameter of 200 μm. Square electric pulses of 0.1 ms duration with a frequency of 0.2 Hz were delivered through a stimulus isolator A360 (WPI, Sarasota, FL), in series with a programmable stimulator (Master8, AMPI, Jerusalem, Israel). All data were collected while the ST was stimulated at 2.5× threshold which was defined as the lowest stimulation intensity to reliably evoke monosynaptic EPSCs. Recordings of glutamatergic EPSCs were performed in the presence of the GABAA receptor antagonist 25 μM gabazine. Bath application of drugs typically lasted about 3-5 min to achieve steady state and begin drug effect tests.

Figure 1
Second-order neurons of arterial chemoreceptor reflex in horizontal NTS slice

For frequency-dependent depression of eEPSCs, a train of 30 stimuli was delivered to ST at frequencies of 0.2, 1, 3, 10, and 20 Hz. The last 20 responses in each train were averaged. A paired-pulse stimulation protocol was used as one means of identifying potential pre-synaptic mechanism. Two synaptic responses (A1 and A2) were evoked by a pair of stimuli given at an interval ranging from 20 to 200 ms. Paired-pulse ratio (PPR) was calculated as the amplitude ratio of the second synaptic response to the first synaptic response (A2/A1). A second analysis of pre-synaptic effects involved examination of miniature EPSCs (mEPSCs). The mEPSCs were recorded in the presence of the sodium channel blocker tetrodotoxin (TTX, 1 μM), and the GABAA receptor antagonist gabazine (25 μM). At least 200 miniature events were collected before, during drug application and 15 min after washout.

Puff drug application

Puff application of glutamate (1 mM) was performed using a syringe pump delivery system (Model 310, Stoelting, Wood Dale, IL). Glutamate was delivered from a patch pipette (~10 μm tip diameter, volume 10 nl, injection duration 0.5 sec) positioned about 25 μm from the recorded neuron to elicit a reproducible post-synaptic response. Post-synaptic responses were recorded at a holding potential of −60 mV in the presence of 1 μM TTX, 300 μM picrotoxin and 100 μM AP-5. In control studies, injection of the same volume of aCSF did not elicit significant changes in holding current (n=4, data not shown), indicating minimal mechanical disturbance by our puff application method.

Western blot analysis

Each rat was anesthetized with isoflurane and decapitated. The brain was removed from the skull and the brainstem was placed in a commercially available brain matrix (Stoelting). The brain stem was then cut into 1 mm thick coronal sections with razor blades. Under a dissecting microscope the caudal NTS was dissected from the sections, placed in microcentrifuge tubes and flash frozen with liquid nitrogen. The caudal NTS was then sonicated in 50 μl of modified RIPA-buffer supplemented with protease and phosphatase inhibitors (Sigma) followed by 30 min incubation on ice. The total homogenate was then centrifuged at 4°C for 30 min at 14,000rpm and the total protein for each sample was determined by the Bradford method. Six μg (GluR1) or 12 μg (GluR2) of the cleared total lysate was loaded onto 10% acrylamide SDS gel, eletrophoresed in Tris-glycine buffer under denaturing conditions and transferred to nitrocellulose membrane (BioRad) in Tris-glycine buffer with 20% methanol. Membranes were blocked for 1h at room temperature with 5% non fat milk in Tris-buffered saline-Tween 20 (TBS-Tween; 50mM Tris base, 200 mM NaCl, 0.1% Tween 20), and then incubated overnight at 4°C with primary antibodies against AMPA receptor subunit GluR1 and GluR2 (both 1: 1000, Calbiochem, Gibbstown, NJ) or b-actin (1:600, Sigma, St Louis, MO). Blots were rinsed 3 times, 10 min each with TBS-Tween and then incubated at room temperature for 1 hour in a horseradish peroxidase conjugated secondary antibody against respective primary antibody host species (1:5000, Sigma, St Louis, MO). The respective immunoreactive bands were detected by enhanced chemiluminescence (ECL reagents, Amersham) and exposed to radiographic film (Hyperfilm ECL, Amersham).

Data analysis

Data are presented as mean ± SEM. Peak amplitudes of averaged evoked post-synaptic currents (≥ 10 sweeps) were calculated as the difference from the baseline measured several milliseconds before the stimulation artifact. Differences in drug effects were tested by ANOVA or t-test. All miniature events were detected with MiniAnalysis software (v6.0, Synaptosoft Inc., Fort Lee, NJ). The threshold value for detecting mEPSCs was set as four times the root-mean-square baseline noise and all miniature events detected by the software were visually checked to minimize errors. Cumulative distributions of miniature synaptic current amplitudes and frequencies were compared using Kolmogorov-Smirnov (K-S test) nonparametric analysis. For western blot analysis, the immunoreactive bands of interest were quantified by densitometry using the Scion program, and the respective values were normalized against β-actin densitometry. Data were expressed as arbitrary densitometric units (ADU). Statistical tests were performed with SigmaStat (v2.03, SPSS software, Chicago, IL), and graphs were made in SigmaPlot (v8.0, SPSS software, Chicago, IL). Values of p<0.05 were considered significant.


DiA was obtained from Molecular Probes (Eugene, OR). Gabazine (SR 95531 hydrobromide) was obtained from Tocris (Ballwin, MO). (R)-(−)-phenylephrine HCl, yohimbine HCl, 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX), (±)-Propranolol HCl, L(−)-norepineprine bitartrate, clonidine HCl and other chemicals were obtained from Sigma (St Louis, MO). To protect NE from degradation, all buffers included the antioxidant sodium metabisulfite (10 μM), and solutions were prepared freshly before the experiment and protected from bright light.


All data were obtained from second-order neurons in the NTS, identified by presence of DiA-labeled somatic appositions as shown in Fig. 1A. Whole-cell patch-clamp recordings were performed on 201 DiA-labeled NTS cells from 69 NORM rats and 49 CSH rats.

CSH alters glutamatergic synaptic transmission in the NTS

There was no statistical difference between NORM (n=113) and CSH neurons (n=88) in resting membrane potential (NORM: 55.5 ± 0.4 mV, CSH: 56.0 ± 0.5 mV) and input resistance (NORM: 1.00 ± 0.05 GΩ, CSH: 1.06 ± 0.07 GΩ).

Electrical stimulation of the solitary tract evoked EPSCs (eEPSCs) in second-order neurons of arterial chemoreceptors in the NTS when GABAergic inputs were blocked with gabazine (Fig. 1B). The variability of onset latencies, as calculated from the standard deviation of the response onset latency from 10 sweeps, was less than 200 μs (median value 88 μs), further suggesting a mono-synaptic input from afferent terminals (Doyle et al., 2001; Zhang and Mifflin 2007). The eEPSCs were mediated by activation of non-NMDA receptors since the non-NMDA receptor antagonist CNQX (10 μM) abolished eEPSCs (Fig. 1B, also see Fig. 1 in Zhang and Mifflin 2007). Similar to findings from others (Bailey et al., 2006), eEPSCs elicited from the tractus were all-or-none responses. Further increases in stimulation intensity above threshold did not change the onset latency or amplitude of the evoked responses. There was no difference in the threshold stimulation intensity (NORM: 104±5 μA vs CSH: 101±9 μA, p>0.05) or the all-or-none nature of the eEPSC comparing NORM and CSH rats (Fig. 1C).

The maximal amplitude of eEPSCs was significantly greater in CSH than in NORM neurons (296.5±20.4 pA, n=76 vs 220.7±13.2 pA, n=92, p<0.01) (Fig. 2A&B). A separate group of rats were allowed to recover from exposure to CSH by remaining in room air for 7 days after CSH treatment. The amplitude of eEPSCs in these recovered rats was not significantly different from that observed in NORM rats (Recovery: 185.5±15.3 pA, n=10). Onset latency of the eEPSC was not affected by CSH (NORM: 4.9±0.2 ms vs CSH: 4.7±0.2 ms vs Recovery: 4.9±0.5 ms). We next investigated potential pre- and/or post-synaptic mechanisms underlying the CSH enhancement of the eEPSC.

Figure 2
Chronic hypoxia enhances eEPSCs elicited by tractus stimulation

We first examined the post-synaptic currents elicited by direct puff application of 1mM glutamate in the presence of GABAA and NMDA receptor blockade in NTS neurons from both NORM and CSH rats (Fig. 2C). The glutamate-evoked inward current in DiA-labeled NTS neurons was significantly greater in CSH than in NORM neurons (127.8±23.5 pA, n=9 vs 55.3±16.8 pA, n=8, p<0.05). Western blot analysis of the caudal NTS indicated an increased level of GluR2 subunit in the NTS of CSH rats (266% increase compared with NORM rats, both n=5) (Fig. 2D). There was no significant difference in the level of GluR1 subunit comparing CSH and NORM rats (Fig. 2D).

Frequency-dependent depression of visceral afferent inputs in the NTS has been proposed to optimize information transfer within central networks (Chen et al., 1999; Doyle et al., 2001; Glaum et al., 1993; Miles 1986; Scheuer et al., 1996; Schild et al., 1998). This phenomenon is considered to be primarily mediated by pre-synaptic mechanisms and related to neurotransmitter release probability (Zucker and Regehr, 2002).A solitary tract stimulation frequency of 0.2 Hz was chosen as the control since there was no significant change in eEPSC amplitude during stimulation at this frequency (Fig. 1B and and2E).2E). The amplitudes of eEPSCs decreased as the frequency of ST stimulation increased. CSH attenuated frequency-dependent depression of eEPSCs (Fig. 2E).

We further examined alterations in pre-synaptic glutamate release in the NTS by analyzing action potential-independent spontaneous glutamate release from pre-synaptic terminals, miniature EPSCs (mEPSCs). The bath solution included gabazine (25 μM) to block GABAA receptors and the sodium channel blocker TTX (1 μM). The mEPSCs were glutamatergic in origin as they were completely abolished by the non-NMDA receptor antagonist CNQX (10 μM) (data not shown). The frequency of mEPSCs in CSH neurons (1.83±0.26 Hz) was significantly less than that observed in NORM neurons (3.62±0.59 Hz, p<0.05) (Fig. 2G,H). There was no difference in the amplitude of mEPSCs between CSH neurons and NORM neurons (CSH: 28.0±3.0 pA, n=11 vs NORM: 26.7±2.3 pA, n=12, p>0.05) (Fig. 2H).

These data indicate that CSH exposure increases both glutamate release probability and the post-synaptic response to glutamate, resulting in enhanced glutamatergic synaptic transmission of arterial chemoreceptor inputs in the NTS.

CSH enhances NE-mediated inhibition of eEPSCs

Bath application of 10 μM NE reduced the amplitude of eEPSCs without a significant change in onset latency in both NORM and CSH neurons (Fig. 3A). In CSH neurons, NE-mediated inhibition of eEPSCs was enhanced compared to NORM neurons. In cells in which all doses of NE were tested, the EC50 of the NE inhibition of eEPSCs was less in CSH neurons (3.0±0.9 μM, n=5) than in NORM neurons (7.6±1.0 μM, n=7, p<0.01) (Fig. 3B). At a concentration of 10 μM, NE caused greater inhibition of eEPSCs in CSH neurons than in NORM neurons (63±2%, n=9 vs 45±3%, n=8, p<0.01).

Figure 3
NE inhibits tractus evoked EPSCs

In most second-order neurons, there was no discernable change in holding current and input resistance during application of 10 μM NE, suggesting the primary site of NE inhibition is pre-synaptic. In cells with a post-synaptic response, 10 μM NE induced outward currents of 18.3±3.6 pA in NORM neurons (9/53) and 15.3±3.2 pA in CSH neurons (5/29), and a decrease in input resistance of 51±4% in NORM neurons and 51±7% in CSH neurons. No significant difference in the post-synaptic response to NE was observed between NORM and CSH neurons that exhibited a post-synaptic response to NE.

CSH enhances NE increase of paired-pulse ratio

To further investigate the pre-synaptic effect of NE, we examined paired-pulse stimulation responses. A change in paired-pulse ratio (PPR, the ratio of second evoked response amplitude to the amplitude of the first evoked response) indicates a pre-synaptic site of action (Zucker and Regehr, 2002). NE (10 μM) increased PPR (Fig. 4A), supporting a pre-synaptic site for NE inhibition of eEPSCs. The PPR was significantly increased in CSH neurons compared with NORM neurons (p<0.01, Fig. 4B). NE (10 μM) significantly increased PPR at a pulse interval of 40 ms (p<0.001) in both NORM and CSH neurons (Fig. 4C). However, the increase was greater in CSH neurons than in NORM neurons (180±11%, n=10 vs 150±8%, n=13, p<0.05).

Figure 4
NE modulates paired-pulse stimulation response

NE inhibition of mEPSCs

We further examined alterations in NE inhibition of pre-synaptic glutamate release in the NTS by recording action potential-independent spontaneous glutamate release from pre-synaptic terminals (mEPSCs). The bath solution included 25 μM gabazine to block GABAA receptors and 1 μM sodium channel blocker TTX to record mEPSCs. NE (10 μM) significantly reduced the frequency of mEPSCs in both CSH neurons and NORM neurons (p<0.001), but not the amplitude of mEPSCs (Fig. 5). As previously discussed, the frequency of mEPSCs was reduced after exposure to CSH. However, there was no difference in NE inhibition of mEPSC frequency comparing NORM (44.0±5.5%, n=7) to CSH (44.9±5.0%, n=8, p>0.05) neurons (Fig. 5B).

Figure 5
NE modulates mEPSCs

NE effect in the NTS is mediated primarily by α-adrenoreceptors

To determine which adrenoreceptors mediate NE inhibition of glutamate release in the NTS, we applied NE (10 μM) alone and in combination with an adrenoreceptor antagonist (Fig. 6A). There was no significant difference in the inhibition of eEPSCs between two repeated applications separated by 15 min of 10 μM NE alone (57±5% vs 58±9%, p>0.05, combined data from 3 NORM and 2 CSH neurons), suggesting no discernable desensitization of adrenoreceptors in this protocol. Bath application the αadrenoreceptor antagonist phentolamine (10 μM) did not significantly alter the amplitudes of eEPSCs in neurons from both groups (97±5% of control, from 4 NORM neurons and 3 CSH neurons), suggesting no tonic activation NTS α-adrenoreceptors in our preparation. Phentolamine nearly abolished NE-mediated inhibition of eEPSCs in both NORM (41±8% vs 6±5%, n=3, p<0.05) and CSH neurons (73±18% vs 8±8%, n=3, p<0.05) (Fig. 6B). These data suggest that NE inhibition in the NTS is primarily mediated by α-adrenoreceptors.

Figure 6
NE effect in the NTS is mediated by α-adrenoreceptors

We further tested the effect of the β-adrenoreceptor antagonist propranolol using the same protocol. Propranolol (10 μM) had no significant effect on eEPSCs (99±3% of control, combined data from 4 NORM neurons and 3 CSH neurons), and NE-mediated inhibition of eEPSCs in both NORM and CSH neurons (99±10 % of NE alone, p>0.05, combined data from 3 NORM neurons and 3 CSH neurons). These data suggest that there was no tonic activation of NTS β-adrenoreceptors in our preparation, and βadrenoreceptors play only a minor role in modulating NE inhibition of eEPSCs in the NTS in our preparation.

CSH has different effect on the function of NTS α1- and α2-adrenoreceptors

The relative role of α1- and α2-adrenoreceptors in NE-mediated inhibition of eEPSCs in the NTS was examined. The α1-adrenoreceptor agonist phenylephrine (40 μM) and the α2-adrenoreceptor agonist clonidine (3 μM) both inhibited eEPSCs (Fig. 7A). However, the effect of CSH on the inhibition mediated by these two α-adrenoreceptors was different (Fig. 7B). CSH attenuated phenylephrine inhibition of eEPSCs (p<0.05), but enhanced clonidine inhibition of eEPSCs (p<0.05).

Figure 7
CSH effect on NTS α1- and α2-adrenoreceptors

In second-order NTS neurons, co-application of 10 μM NE and the α2-adrenoreceptor antagonist yohimbine significantly attenuated NE-mediated inhibition of eEPSCs (Fig. 8A). Yohimbine itself did not significantly change the amplitudes of eEPSCs (NORM: 100±2% of control, n=6 and CSH 99±2% of control, n=5), again suggesting no tonic activation of NTS α2-adrenoreceptor in our preparation. In NORM neurons (n=8), yohimbine reduced NE-mediated inhibition of eEPSCs from 42±7 % to 14±2 %, a 61±8 % attenuation. In CSH neurons (n=5), yohimbine reduced NE-mediated inhibition of eEPSCs from 63±9 % to 16±8 %, an attenuation of 76±11 % (Fig. 8B, repeated). There was a significant difference in baseline NE-mediated inhibition between the two groups (p<0.01). There was no significant difference between NORM and CSH neurons in the NE-mediated inhibition that remained after blocking α2-adrenoreceptors with yohimbine.

Figure 8
The role of α2-adrenoreceptors in NE inhibition in the NTS

In a separate group of neurons, NE (10 μM) and yohimbine were applied simultaneously without prior application of NE alone to ensure that the antagonist effect was not altered by prior exposure to NE. In this protocol the NE-mediated inhibition remaining after blocking α2 adrenoreceptors with yohimbine was not different between NORM neurons (20±4 %, n=4) and CSH neurons (22±5 %, n=7) (Fig. 8B, single). No difference was observed in NE inhibition comparing protocols where antagonist effects were measured in the absence of prior exposure to NE or after prior exposure to NE. The data from these 2 protocols strongly suggests that although both α1- and α2-adrenoreceptors mediate NE inhibition in the NTS, the enhanced NE inhibition of eEPSCs observed following CSH is primarily mediated by α2-adrenoreceptors.


We report two significant new findings. First, exposure to CSH enhances the amplitude of the glutamatergic EPSC evoked by peripheral afferent stimulation in NTS neurons receiving monosynaptic arterial chemoreceptor inputs. Both pre-synaptic and post-synaptic factors appear to contribute to the enhanced eEPSC. Second, presynaptic inhibition of glutamate release from afferents to these same neurons by α2-adrenoreceptors is enhanced following chronic exposure to hypoxia.

We found that the amplitude of eEPSCs was greater in NTS neurons from CSH rats compared to NORM rats. This is the result of an increased post-synaptic sensitivity to excitatory amino acids as demonstrated by our puff applications of glutamate. Since these applications were carried out in the presence of NMDA receptor blockade, it is likely that this increased sensitivity reflects alterations in the AMPA receptor. In fact, we observed increased levels of AMPA receptor GluR2 subunit, but not GluR1 subunit, in the NTS after CSH exposure. Previous study of our lab provided electrophysiological evidence suggesting that the AMPA receptors on NTS neurons receiving monosynaptic arterial chemoreceptor inputs contain GluR2 subunits (de Paula et al., 2007). Possible mechanisms underlying increased expression and/or sensitivity of AMPA receptors include hypoxia-induced activation of multiple kinases leading to phosphorylation of AMPA receptor subunits, activity-dependent changes in AMPA receptor subunit composition and increased expression and/or trafficking of AMPA receptors (Derkach et al., 2007; Esteban et al., 2003; Gozal et al., 2000; Gozal et al., 1998; Schmid et al., 2008). Along with enhanced excitation of carotid bodies following exposure to CSH (Prabhakar and Jacono, 2005; Powell, 2007), these data suggest both peripheral and central neural mechanisms contribute to enhanced response to acute hypoxia after exposure to CSH (Powell et al., 2000).

We also found alterations in glutamate release following exposure to CSH. The frequency of mEPSCs was reduced, suggesting a reduction in release probability. This may not seem consistent with our finding of an increased eEPSC after exposure to CSH. However, it is important to keep in mind that the eEPSC reflects synchronous activation of peripheral afferent inputs to the cell, while mEPSCs reflect spontaneous release from any glutamatergic terminals in close proximity to the cell. As such, mEPSCs reflect potential glutamate release from not only peripheral afferents but also inputs from other central sites and/or local interneurons. The modulation of vesicular release mechanisms can differ depending upon whether the release sites are synaptic or ectopic (Matsui and Jahr, 2004). The eEPSC may be representative of release from synaptic sites while mEPSCs may reflect, at least in part, ectopic release.

A recent report found that exposure to chronic intermittent hypoxia (CIH) depressed eEPSCs in NTS neurons studied in a brain slice (Kline et al., 2007). The authors suggested that CIH-induced depression of EPSCs is mediated by a pre-synaptic mechanism without the apparent involvement of any post-synaptic changes. In this report the absence of post-synaptic changes was inferred from the lack of change in the amplitude of mEPSCs. In contrast, another report shows that CIH enhanced neuronal response to exogenous application of AMPA in enzymatically dissociated NTS second-order neurons without a significant change in EC50 (de Paula et al., 2007). In our present study, we also found no changes in the amplitude of mEPSCs after CSH, however direct testing of post-synaptic sensitivity to glutamate revealed a different story. Responses to exogenous applications of agonist could include receptors not normally activated by synaptically released glutamate or, as discussed above, analysis of spontaneous transmitter release may include inputs not derived from the primary afferent. Nonetheless, more work needs to be done to differentiate changes in pre-synaptic and post-synaptic function following chronic exposures to both sustained and intermittent hypoxia.

Our current results demonstrated that NE inhibition of eEPSCs is mediated by both pre-synaptic α1- and α2-adrenoreceptors, and this inhibitory effect is enhanced after CSH. Furthermore, pre-synaptic α2-adrenoreceptors mediate the CSH-enhanced NE inhibition. These results extend our previous investigation of pre-synaptic α1-adrenoreceptors in modulating synaptic transmission of arterial chemoreceptor inputs in the NTS (Zhang and Mifflin, 2007) and reveal a previously unrecognized central neural adaptive response to CSH. Our results are consistent with previous in vivo studies demonstrating inhibitory effects of NE mediated by both α1- and α2-adrenoreceptors in the NTS (De Jong, 1974; Feldman and Moises, 1987; Feldman and Moises, 1988; Feldman and Felder, 1989a, 1989b; Moore and Guyenet, 1983; Silva de Oliveira et al., 2007). However, one study reported that microinjection of an α2-adrenoreceptor antagonist into the NTS attenuated the arterial chemoreflex (Hayward, 2001). Such a result seems contradictory to our findings. We have observed that activation of α2-adrenoreceptors can also inhibit IPSCs in the NTS via a pre-synaptic mechanism (Zhang and Mifflin, unpublished observation). NE inhibits GABAergic synaptic transmission via pre-synaptic α2-adrenoreceptors in other central neural sites (Han et al., 2002; Hirono and Obata, 2006; Li et al., 2005). Thus the neuronal responses in the NTS will depend on the balance between excitatory glutamatergic inputs and inhibitory GABAergic inputs. This could be one possible mechanism underlying the inhibitory effect of α2-adrenoreceptor antagonist in arterial chemoreflexes (Hayward, 2001). Future studies will be needed to investigate NE-mediated modulation of IPSCs in the NTS and the impact of CSH.

This current study established a major role for α2-adrenoreceptors in mediating CSH-induced alterations of NE inhibition in the NTS. This could represent a neural adaptive response to CSH with potential physiological significance. Increased chemoreceptor discharge following CSH (Prabhakar and Jacono, 2005; Powell, 2007) should result in increased arterial chemoreceptor inputs to the NTS. Supporting this is the observation that the respiratory response to acute hypoxia is enhanced after CSH (Powell, 2000). Our study also found attenuated frequency-dependent depression of eEPSCs in brainstem slices collected from CSH rats, resulting in larger amplitude eEPSCs at any given tractus stimulation frequency, which suggests increased glutamate release from pre-synaptic afferent terminals. Enhanced pre-synaptic α2-adrenoreceptor-mediated inhibition of glutamate release from afferent terminals might be an adaptive mechanism to limit excitotoxic damage and maintain neuronal function. CSH enhanced α2-adrenoreceptor-mediated inhibition of neuronal excitability has been reported in the locus coeruleus (Chang et al., 2006). This post-synaptic change is correlated with an increased number of neuronal α2-receptor binding sites in locus coeruleus. There might be a similar up-regulation of pre-synaptic α2-adrenoreceptors in the NTS after CSH.

Under what conditions do NTS adrenoceptors modulate chemoreceptor afferent integration? CSH increased the activity of tyrosine hydroxylase, the rate-limiting enzyme in catecholamine biosynthesis, in the NTS including A2 cell group (Pepin et al., 1996; Soulier et al., 1995), the ventrolateral medulla (Schmitt et al., 1993) and the locus coeruleus (Schmitt et al., 1993). Furthermore, CSH increased NE turnover in A2 cell group, but decreased that in A5 cell group and locus coeruleus with no significant effect in A1 cell group (Soulier et al., 1992). These noradrenergic structures appear to be involved in arterial chemoreflex pathways since systemic hypoxia or carotid sinus nerve stimulation increases c-fos expression in these structures (Buller et al., 1999; Erickson and Millhorn, 1994; Smith et al., 1995; Teppema et al., 1997). Arterial chemoreceptor stimulation evoked neuronal discharge in these noradrenergic structures (Guyenet et al., 1993; Li et al., 1992), suggesting their active roles in modulating arterial chemoreflexes. In vivo studies have demonstrated that excitation or inhibition of these noradrenergic neural structures can modulate cardiorespiratory responses to arterial chemoreceptor stimulation (Hayward, 2001; Koshiya and Guyenet, 1994a; Koshiya and Guyenet, 1994b; Perez et al., 1998). These data suggest that activation of central noradrenergic neural structures during arterial chemoreceptor stimulation could release NE and modulate synaptic transmission in the NTS. Our current data cannot evaluate the contribution of different noradrenergic neural structures. The A2 cell group might be an important source of NE since it is located at the first central site of arterial chemoreflex pathways and receives increased afferent inputs after CSH. At least some A2 neurons are second-order neurons of arterial chemoreceptors (Kawai and Senba, 1999; Zhang and Mifflin, unpublished observation). These neurons may play a crucial role in neural adaptations to CSH.

In summary, our current project investigated one inhibitory mechanism in the NTS after CSH. Although the role of NE in arterial chemoreflexes requires further investigation (Joseph et al., 1998; McCrimmon et al., 1983; Schreihofer and Guyenet, 2000), we suggest that enhanced NE inhibition of glutamate release is a neural adaptive mechanism in response to increased afferent inputs during CSH. CSH results in enhanced excitation of carotid bodies (Prabhakar and Jacono, 2005; Powell, 2007), which may initiate changes in central synaptic transmission such as increased amplitude of solitary tract-evoked EPSCs and attenuated frequency-dependent depression of eEPSCs reported here. Tissue hypoxia may also contribute to the initiation and/or maintenance of these changes in synaptic transmission (Zhang et al., 2008). The synaptic integrative functions of the NTS will influence the flow of information downstream in the reflex pathway (Loewy, 1990; Mifflin, 1992). Alterations in NE inhibition could play a crucial role in neural plasticity of synaptic integration of arterial chemoreceptor inputs in the NTS.


Authors acknowledge the assistance from Myrna Herrera-Rosales and Melissa Vitela. This study was supported by National Heart, Lung, and Blood Institute grant HL-41894 (to S. W. Mifflin) and HL-62579 (to J. T. Cunningham)


  • Aoki C, Zemcik BA, Strader CD, Pickel VM. Cytoplasmic loop of beta-adrenergic receptors: synaptic and intracellular localization and relation to catecholaminergic neurons in the nuclei of the solitary tracts. Brain Res. 1989;493:331–347. [PubMed]
  • Bailey TW, Jin YH, Doyle MW, Smith SM, Andresen MC. Vasopressin inhibits glutamate release via two distinct modes in the brainstem. J Neurosci. 2006;26:6131–6142. [PMC free article] [PubMed]
  • Buller KM, Smith DW, Day TA. NTS catecholamine cell recruitment by hemorrhage and hypoxia. Neuroreport. 1999;10:3853–3856. [PubMed]
  • Chang KC, Yang JJ, Liao JF, Wang CSH, Chiu TH, Hsu FC. Chronic hypobaric hypoxia induces tolerance to acute hypoxia and up-regulation in alpha-2 adrenoceptor in rat locus coeruleus. Brain Res. 2006;1106:82–90. [PubMed]
  • Chen CY, Horowitz JM, Bonham AC. A pre-synaptic mechanism contributes to depression of autonomic signal transmission in NTS. Am J Physiol Heart Circ Physiol. 1999;277:H1350–H1360. [PubMed]
  • Dashwood MR, Gilbey MP, Spyer KM. The localization of adrenoceptors and opiate receptors in regions of the cat central nervous system involved in cardiovascular control. Neuroscience. 1985;15:537–551. [PubMed]
  • Day HE, Campeau S, Watson SJ, Jr, Akil H. Distribution of α1a-, α1b- and α1d-adrenergic receptor mRNA in the rat brain and spinal cord. J Chem Neuroanat. 1997;13:115–139. [PubMed]
  • De Jong W. Noradrenaline: Central inhibitory control of blood pressure and heart rate. Eur J Pharmacol. 1974;29:179–181. [PubMed]
  • de Paula PM, Tolstykh G, Mifflin S. Chronic intermittent hypoxia alters NMDA and AMPA-evoked currents in NTS neurons receiving carotid body chemoreceptor inputs. Am J Physiol Regul Integr Comp Physiol. 2007;292:R2259–R2265. [PubMed]
  • Derkach VA, Oh MC, Guire ES, Soderling TR. Regulatory mechanisms of AMPA receptors in synaptic plasticity. Nat Rev Neurosci. 2007;8:101–113. [PubMed]
  • Doyle MW, Andresen MC. Reliability of monosynaptic sensory transmission in brain stem neurons in vitro. J Neurophysiol. 2001;85:2213–2223. [PubMed]
  • Erickson JT, Millhorn DE. Hypoxia and electrical stimulation of the carotid sinus nerve induce Fos-like immunoreactivity within catecholaminergic and serotoninergic neurons of the rat brainstem. J Comp Neurol. 1994;348:161–182. [PubMed]
  • Esteban JA, Shi SH, Wilson C, Nuriya M, Huganir RL, Malinow R. PKA phosphorylation of AMPA receptor subunits controls synaptic trafficking underlying plasticity. Nat Neurosci. 2003;6:136–143. [PubMed]
  • Featherby T, Lawrence AJ. Chronic cold stress regulates ascending noradrenergic pathways. Neuroscience. 2004;127:949–960. [PubMed]
  • Feldman PD, Felder RB. Alpha-adrenergic influences on neuronal responses to visceral afferent input in the nucleus tractus solitarius. Neuropharmacology. 1989a;28:1081–1087. [PubMed]
  • Feldman PD, Felder RB. α2-Adrenergic modulation of synaptic excitability in the rat nucleus tractus solitarius. Brain Res. 1989b;480:190–197. [PubMed]
  • Feldman PD, Moises HC. Electrophysiological evidence for α1- and α2-adrenoceptors in solitary tract nucleus. Am J Physiol Heart Circ Physiol. 1988;254:H756–H762. [PubMed]
  • Feldman PD, Moises HC. Adrenergic responses of baroreceptive cells in the nucleus tractus solitarii of the rat: a microiontophoretic study. Brain Res. 1987;420:351–361. [PubMed]
  • Glaum SR, Miller RJ. Pre-synaptic metabotropic glutamate receptors modulate ω-Conotoxin-GVIA-insensitive calcium channels in the rat medulla. Neuropharmacology. 1995;34:953–964. [PubMed]
  • Globus MY, Busto R, Dietrich WD, Martinez E, Valdes I, Ginsberg MD. Direct evidence for acute and massive norepinephrine release in the hippocampus during transient ischemia. J Cereb Blood Flow Metab. 1989;9:892–896. [PubMed]
  • Gozal D, Gozal E, Simakajornboon N. Signaling pathways of the acute hypoxic ventilatory response in the nucleus of the solitary tract. Respir Physiol. 2000;121:209–221. [PubMed]
  • Gozal E, Roussel AL, Holt GA, Gozal L, Gozal YM, Torres JE, Gozal D. Protein kinase C modulation of ventilatory responses to hypoxia in nucleus tractus solitarii of conscious rats. J Appl Physiol. 1998;84:1982–1990. [PubMed]
  • Guyenet PG, Koshiya N, Huangfu D, Verberne AJ, Riley TA. Central respiratory control of A5 and A6 pontine noradrenergic neurons. Am J Physiol Regul Integr Comp Physiol. 1993;264:R1035–R1044. [PubMed]
  • Han SK, Chong W, Li LH, Lee IS, Murase K, Ryu PD. Noradrenaline excites and inhibits GABAergic transmission in parvocellular neurons of rat hypothalamic paraventricular nucleus. J Neurophysiol. 2002;87:2287–2296. [PubMed]
  • Hayward LF. Evidence for α-2 adrenoreceptor modulation of arterial chemoreflexes in the caudal solitary nucleus of the rat. Am J Physiol Regul Integr Comp Physiol. 2001;281:R1464–R1273. [PubMed]
  • Hirono M, Obata K. α-Adrenoceptive dual modulation of inhibitory GABAergic inputs to Purkinje cells in the mouse cerebellum. J Neurophysiol. 2006;95:700–708. [PubMed]
  • Ilyinsky O, Mifflin S. Chronic hypoxia abolishes expiratory prolongation following carotid sinus nerve stimulation in the anesthetized rat. Respir Physiol Neurobiol. 2005;146:269–277. [PubMed]
  • Ilyinsky O, Tolstykh G, Mifflin S. Chronic hypoxia abolishes posthypoxia frequency decline in the anesthetized rat. Am J Physiol Regul Integr Comp Physiol. 2003;285:R1322–R1330. [PubMed]
  • Jones LS, Gauger LL, Davis JN. Anatomy of brain alpha 1-adrenergic receptors: in vitro autoradiography with [125I]-heat. J Comp Neurol. 1985;231:190–208. [PubMed]
  • Joseph V, Dalmaz Y, Cottet-Emard JM, Pequignot JM. Dexamethasone's influence on tyrosine hydroxylase activity in the chemoreflex pathway and on the hypoxic ventilatory response. Pflugers Arch. 1998;435:834–839. [PubMed]
  • Kawai Y, Senba E. Electrophysiological and morphological characterization of cytochemically-defined neurons in the caudal nucleus of tractus solitarius of the rat. Neuroscience. 1999;89:1347–1355. [PubMed]
  • Kline DD, Ramirez-Navarro A, Kunze DL. Adaptive depression in synaptic transmission in the nucleus of the solitary tract after in vivo chronic intermittent hypoxia: evidence for homeostatic plasticity. J Neurosci. 2007;27:4663–4673. [PubMed]
  • Koshiya N, Guyenet PG. Role of the pons in the carotid sympathetic chemoreflex. Am J Physiol Regul Integr Comp Physiol. 1994a;267:R508–R518. [PubMed]
  • Koshiya N, Guyenet PG. A5 noradrenergic neurons and the carotid sympathetic chemoreflex. Am J Physiol Regul Integr Comp Physiol. 1994b;267:R519–R526. [PubMed]
  • Li DP, Atnip LM, Chen SR, Pan HL. Regulation of synaptic inputs to paraventricular-spinal output neurons by α2 adrenergic receptors. J Neurophysiol. 2005;93:393–402. [PubMed]
  • Li YW, Gieroba ZJ, Blessing WW. Chemoreceptor and baroreceptor responses of A1 area neurons projecting to supraoptic nucleus. Am J Physiol Regul Integr Comp Physiol. 1992;263:R310–R317. [PubMed]
  • Lipton P. Ischemic cell death in brain neurons. Physiol Rev. 1999;79:1431–1568. [PubMed]
  • Loewy AD. In: Central autonomic pathways, In: Central regulation of autonomic functions. Loewy AD, Spyer KM, editors. Oxford; New York: 1990. pp. 88–103.
  • Matsui K, Jahr CE. Differential control of synaptic and extopic vesicular releae of glutamate. J Neurosci. 2004;24:8932–8939. [PubMed]
  • McCrimmon DR, Dempsey JA, Olson EB., Jr Effect of catecholamine depletion on ventilatory control in unanesthetized normoxic and hypoxic rats. J Appl Physiol. 1983;55:522–528. [PubMed]
  • Mifflin SW. Arterial chemoreceptor input to nucleus tractus solitarius. Am J Physiol Regul Integr Comp Physiol. 1992;263:R368–R375. [PubMed]
  • Miles R. Frequency dependence of synaptic transmission in nucleus of the solitary tract in vitro. J Neurophysiol. 1986;55:1076–1090. [PubMed]
  • Moore SD, Guyenet PG. Alpha-receptor mediated inhibition of A2 noradrenergic neurons. Brain Res. 1983;276:188–191. [PubMed]
  • Nakai T, Milusheva E, Baranyi M, Uchihashi Y, Satoh T, Vizi ES. Excessive release of [3H]noradrenaline and glutamate in response to simulation of ischemic conditions in rat spinal cord slice preparation: effect of NMDA and AMPA receptor antagonists. Eur J Pharmacol. 1999;366:143–150. [PubMed]
  • Pepin JL, Levy P, Garcin A, Feuerstein G, Savasta M. Effects of long-term hypoxia on tyrosine hydroxylase protein content in catacholaminergic rat brainstem areas: a quantitative autoradiographic study. Brain Res. 1996;733:1–8. [PubMed]
  • Perego C, Gatti S, Vetrugno GC, Marzatico F, Algeri S. Correlation between electroencephalogram isoelectric time and hippocampal norepinephrine levels, measured by microdialysis, during ischemia in rats. J Neurochem. 1992;59:1257–1262. [PubMed]
  • Perez H, Ruiz S, Laurido C, Hernandez A. Locus coeruleus-mediated inhibition of chemosensory responses in the rat nucleus tractus solitarius is mediated by α2-adrenoreceptors. Neurosci Lett. 1998;249:37–40. [PubMed]
  • Powell FL, Huey KA, Dwinell MR. Central nervous system mechanisms of ventilatory acclimatization to hypoxia. Respir Physiol. 2000;121:223–236. [PubMed]
  • Powell FL. The influence of chronic hypoxia upon chemoreception. Resp Physiol Neurobiol. 2007;157:154–161. [PMC free article] [PubMed]
  • Prabhakar NR, Jacono FJ. Cellular and molecular mechanisms associated with carotid body adaptations to chronic hypoxia. High Alt Med Biol. 2005;6:112–120. [PubMed]
  • Scheuer DA, Zhang J, Toney GM, Mifflin SW. Temporal processing of aortic nerve evoked activity in the nucleus of the solitary tract. J Neurophysiol. 1996;76:3750–3757. [PubMed]
  • Schild JH, Clark JW, Canavier CC, Kunze DL, Andresen MC. Afferent synaptic drive of rat medial nucleus tractus solitarius neurons: dynamic simulation of graded vesicular mobilization, release, and non-NMDA receptor kinetics. J Neurophysiol. 1998;74:1529–1548. [PubMed]
  • Schmid A, Hallermann S, Kittel RJ, Khorramshahi O, Frölich AM, Quentin C, Rasse TM, Mertel S, Heckmann M, Sigrist SJ. Activity-dependent site-specific changes of glutamate receptor composition in vivo. Nat Neurosci. 2008;11:659–666. [PubMed]
  • Schmitt P, Pequiqnot J, Garcia C, Pujol JF, Pequiqnot JM. Regional specificity of the long-term regulation of tyrosine hydroxylase in some catecholaminergic rat brainstem areas. I. influence of long-term hypoxia. Brain Res. 1993;611:53–60. [PubMed]
  • Schmitt P, Soulier V, Pequignot JM, Pujol JF, Denavit-Saubie M. Ventilatory acclimatization to chronic hypoxia: relationship to noradrenaline metabolism in the rat solitary complex. J Physiol (London) 1994;477:331–337. [PubMed]
  • Schreihofer AM, Guyenet PG. Sympathetic reflexes after depletion of bulbospinal catecholaminergic neurons with anti-DβH-saporin. Am J Physiol Regul Integr Comp Physiol. 2000;279:R729–R742. [PubMed]
  • Silva de Oliveira LC, Bonagamba LG, Machado BH. Noradrenergic inhibitory modulation in the caudal commissural NTS of the pressor response to chemoreflex activation in awake rats. Auton Neurosci. 2007;136:63–68. [PubMed]
  • Smith DW, Buller KM, Day TA. Role of ventrolateral medulla catecholamine cells in hypothalamic neuroendocrine cell responses to systemic hypoxia. J Neurosci. 1995;15:7979–7988. [PubMed]
  • Soulier V, Cottet-Emard JM, Pequignot J, Hanchin F, Peyrin L, Pequignot JM. Differential effects of long-term hypoxia on norepinephrine turnover in brain stem cell groups. J Appl Physiol. 1992;73:1810–1814. [PubMed]
  • Soulier V, Dalmaz Y, Cottet-Emard JM, Kitahama K, Pequignot JM. Delayed increase of tyrosine hydroxylation in the rat A2 medullary neurons upon long-term hypoxia. Brain Res. 1995;674:188–195. [PubMed]
  • Teppema LJ, Veening JG, Kranenburg A, Dahan A, Berkenbosch A, Olievier C. Expression of c-fos in the rat brainstem after exposure to hypoxia and to normoxic and hyperoxic hypercapnia. J Comp Neurol. 1997;388:169–190. [PubMed]
  • Tolstykh G, Belugin S, Mifflin S. Responses to GABAA receptor activation are altered in NTS neurons isolated from chronic hypoxic rats. Brain Res. 2004;1006:107–113. [PubMed]
  • Young WS, III, Kuhar MJ. Noradrenergic α1 and α2 receptors: light microscopic autoradiographic localization. Proc Natl Acad Sci USA. 1980;77:1696–1700. [PubMed]
  • Zhang W, Mifflin SW. Modulation of synaptic transmission to second-order peripheral chemoreceptor neurons in caudal nucleus tractus solitarius by α1-adrenoreceptors. J Pharmacol Exp Ther. 2007;320:670–677. [PubMed]
  • Zhang W, Carreño FR, Cunningham JT, Mifflin SW. Chronic sustained and intermittent hypoxia reduce function of ATP-sensitive potassium channels in nucleus of the solitary tract. Am J Physiol Regul Integr Comp Physiol. 2008;295:R1555–R1562. [PubMed]
  • Zucker RS, Regehr WG. Short-term synaptic plasticity. Annu Rev Physiol. 2002;64:355–405. [PubMed]