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Department of Pharmacology and Physiology, George Washington University, Washington, District of Columbia
Submitted 12 September 2006; accepted in final form 1 November 2006
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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). The activity of CVNs is intricately linked to respiratory function and respiratory-related inputs to CVNs are critical for coordinated homeostatic interactions. For example, respiratory sinus arrhythmia, in which heart rate increases during each inspiration to optimize blood flow to the lungs, is mediated by increases in both inhibitory GABAergic and glycinergic neurotransmission to CVNs during inspiration (Neff et al. 2003).
The cardiorespiratory system in the brain stem exhibits a broad range of plasticity in response to physiological challenges. For example, hypoxia transforms eupnea to gasping and simultaneously converts an inspiratory evoked increase in heart rate to a pronounced parasympathetically mediated bradycardia, which decreases the metabolic demands of the heart and increases the likelihood of survival (Guntheroth and Kawabori 1975). The biphasic respiratory and heart rate response to hypoxia is paralleled by a biphasic change in inhibitory neurotransmission to CVNs (Neff et al. 2004). In addition to transient responses to a single episode of hypoxia, intermittent hypoxia can have distinct and long-lasting effects on cardiorespiratory network plasticity and alter responses to subsequent hypoxias, such as the long-term facilitation of respiratory activity (Baker and Mitchell 2000).
In most other brain regions, reactive oxygen species (ROS) generation is associated with and contributes to neuronal death resulting from excitotoxicity, hypoxia, and/or hypercapnia and various neurodegenerative disorders (Butterfield and Lauderback 2002; Rego and Oliveira 2003; Xu et al. 2004). However, ROS are increasingly recognized as a key component of homeostasis, participating in neuronal signaling and plasticity. For example, high concentrations of extracellular antioxidants attenuate long-term potentiation and synaptic transmission in the hippocampus, and superoxide is required for long-term potentiation (Kamsler and Segal 2003; Knapp and Klann 2002). Further, ROS are implicated in central autonomic network function; for example ROS are required for Angiotensin II signaling in the NTS, hypothalamus, and cortex (Kim-Mitsuyama et al. 2005; Sun et al. 2005; Wang et al. 2004).
Whereas the cardiorespiratory responses to a single period of hypoxia can be mediated by changes in inhibitory neurotransmission without changes in excitatory neurotransmission to CVNs, intermittent hypoxia elicits a different pattern of network responses. This study examines the role of ROS in the cardiorespiratory responses to intermittent hypoxia and has three main objectives: to test the hypothesis that acute intermittent hypoxia incrementally recruits a respiratory-related excitatory neurotransmission to CVNs not present during an equivalent duration of continuous hypoxia; to test the hypothesis that ROS generation is a crucial signal that is required for the respiratory-related increases in excitatory neurotransmission during intermittent hypoxia; and to identify candidate neurons that incrementally generate ROS and may constitute the source of excitatory neurotransmission to CVNs evoked during intermittent hypoxia.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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CVNs were retrogradely labeled from the heart as described previously (Mendelowitz and Kunze 1991) in Sprague Dawley rat pups (P1–P3; Hilltop). After 
24 h recovery, pups (P2–P7) were anesthetized and sacrificed, and the hindbrain was removed and placed in cold physiological saline solution [which contained (in mM) 140 NaCl, 5 KCl, 2 CaCl2, 5 glucose, and 10 HEPES, bubbled with 100% O2, pH 7.4]. A slice of the medulla (800 µm) was obtained that included parasympathetic cardiac neurons, the rostral hypoglossal nucleus and rootlets, and the pre-Botzinger complex (Neff et al. 2003). This slice was perfused in a recording chamber with artificial cerebrospinal fluid (ACSF) [containing (in mM) 125 NaCl, 3 KCl, 2 CaCl2, 26 NaHCO3, 5 glucose, and 5 HEPES, equilibrated with 95% O2-5% CO2, pH 7.35–7.4]. All animal procedures were performed with the approval of the Animal Care and Use Committee of The George Washington University in accordance with the recommendations of the panel on euthanasia of the American Veterinary Medical Association and the National Institutes of Health Guide for the Care and Use of Laboratory Animals.
Electrophysiology
Simultaneous recordings of glutamatergic synaptic events in fluorescently identified parasympathetic cardiac neurons in the external formation of the nucleus ambiguus and extracellular recordings of respiratory-related activity from the hypoglossal rootlet were obtained. Spontaneous respiratory-related activity was recorded by monitoring motor-neuron population activity from hypoglossal nerve rootlets using a suction electrode. Hypoglossal rootlet activity was amplified (50,000 times) and filtered (10- to 300-Hz band-pass; CWE, Ardmore, PA) and electronically integrated (
= 50 ms; CWE). Parasympathetic cardiac neurons were identified by the presence of the fluorescent tracer as described previously (Mendelowitz 1996). Recent work has shown this method does not label neurons in the compacta formation but identifies cardiac vagal neurons localized in the external formation of the NA (Bouairi et al. 2006). Patch pipettes (2.5–3.5 M
) were visually guided to the surface of individual CVNs using differential interference optics and infrared illumination (Zeiss, Oberkochen, Germany). CVNs were voltage-clamped at a holding potential of –80 mV. The patch pipettes were filled with a solution consisting of (in mM) 135 K-gluconic acid, 10 HEPES, 10 EGTA, 1 CaCl2, and 1 MgCl2, at a pH of 7.35–7.4. Focal drug application was performed using a pneumatic picopump pressure delivery system (WPI, Sarasota, FL). Drugs were ejected from a patch pipette positioned within 30 µm from the patched CVN. The maximum range of drug application has been determined previously to be 100–120 µm downstream from the drug pipette and considerably less behind the drug pipette (Wang et al. 2002). Glutamatergic postsynaptic currents were isolated by the focal application of gabazine (25 µM) and strychnine (1 µM) to block GABAergic and glycinergic postsynaptic currents, respectively. One experiment was performed per preparation. All synaptic events were blocked at the end of each experiment with the addition of gabazine (25 µM), strychnine (1 µM), 2-amino-5-phosphonopentanoic acid (AP5; 50 µM), and 6-cyano-7-nitroquinoxalene-2,3-dione (CNQX; 50 µM) to the perfusate to block GABAergic, glycinergic, N-methyl-D-aspartate (NMDA) and non-NMDA currents, respectively.
Exogenous glutamate application
Glutamate (50 µM) was focally applied to parasympathetic cardiac neurons using a picospritzer and ejected from a patch pipette positioned within 30 µ of the patched neuron. Glutamate was applied once every 30 s for the extent of the intermittent hypoxia protocol, or in the presence of 4-hydroxytetramethyl-piperidine-1-oxyl (Tempol; EMD Biosciences; 1 mM).
Intermittent hypoxia
Rhythmic inspiratory-related activity and glutamatergic synaptic events in a single parasympathetic cardiac neuron were recorded simultaneously for 6 min in ACSF bubbled with 95% O2-5% CO2. Slices were exposed to three periods of hypoxia of 5-min duration by perfusion of isocapnic ACSF equilibrated with 75% N2-20% O2-5% CO2. After each hypoxic exposure, slices were returned to control perfusate for 15 min. In some experiments, Tempol (1 mM) was included in the perfusate.
ROS imaging and immunohistochemistry
Slices were incubated in 5-(and-6)-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate (CM-H2DCFDA; Molecular probes; 10 µM) for 1 h. CM-H2DCFDA is converted to 2'-7'-dichlorodihydrofluorescein within the cell and subsequently fluoresces when converted to 2'-7'-dichlorofluorescein (DCF) by reactive oxygen species. Time-lapse recordings of DCF fluorescence once every 10 s were obtained using a cooled CCD camera (RTE/CCD-1300-Y/HS; Princeton Instruments; Trenton, NJ) and MetaMorph imaging systems (Molecular Devices), and DCF fluorescence intensity throughout the intermittent hypoxia protocol was calculated. Tempol (1 mM) was added to the perfusate in some experiments. Time-dependent controls were obtained in separate slices by identical time-lapse experiments with only control ACSF exposure. Due to the photosensitivity of DCF (Afzal et al. 2003), ROS imaging was not performed simultaneously with patch-clamp experiments.
For identification of glutamatergic neurons, an antibody to phosphate-activated glutaminase (PAG; kindly provided by Dr. Takeshi Kaneko, Kyoto University), the slice was subsequently fixed overnight at 4°C in 75.7% picric acid/0.81% paraformaldehyde in phosphate buffer, cryoprotected at 4°C in 30% sucrose solution for 2 h and embedded in O.C.T embedding medium at –20°C. Sections (60 µm) were obtained on a cryostat, washed three times in 0.1M PBS, and incubated for 1 h in 5% normal goat serum/0.01% Triton X in 0.1M PBS. Slices were transferred to primary antibody mixture containing the 40 µg/ml mouse anti-PAG/0.01% triton X in 0.1M PBS incubated overnight at 4°C. Slices were rinsed three times in 0.1M PBS at room temperature, incubated in Alexa Fluor 647 goat anti-mouse IgM secondary antibody (1:200; Invitrogen) in 0.1M PBS for 2 h at room temperature, and washed three times with 0.1M PBS. For control samples in adjacent slices, PAG antibody was substituted with normal mouse serum in a comparable dilution to the antibody. Published studies (Kaneko et al. 1987; Van der Gucht et al. 2003) have also confirmed the anti-PAG primary antibody specificity. Images were acquired with an Axiocam MR cooled CCD monochrome digital camera (431–146; Zeiss). Counts of DCF/PAG expressing cells were compiled from six separate images from six separate slices of the ventrolateral region at x40 magnification for each experiment.
Oxygen measurements
Tissue pO2 was recorded using a Clark-style oxygen-sensitive microelectrode (10 µm tip) and Chemical Microsensor polarographic amplifier (Diamond General) as described (Neff et al. 2004). Tissue pO2 was measured at a 400-µ depth throughout the intermittent hypoxia protocol.
Statistical analysis
Analysis of spontaneous synaptic currents was performed using MiniAnalysis (Synaptosoft) with minimal acceptable amplitude determined by the amplitude at which all synaptic events were blocked in the presence of AP5 and CNQX. Excitatory postsynaptic current (EPSC) frequency in CVNs was grouped into 1-s bins and cross-correlated with the onset of inspiratory-related hypoglossal activity. Frequency cross-correlations were analyzed from all bursts throughout the entire experiment. Frequency histograms were averaged from all of the respiratory bursts for each administration; for hypoxias, all frequency histograms in the entire 5-min exposure were averaged. Results are presented as means ± SE. Statistical comparisons were performed between control and hypoxic conditions as well as between the first second of the inspiratory burst and 5 s before burst onset using ANOVA with repeated measures. Significant difference was set at P < 0.05. For time-lapse experiments, 350-µ regions of DCF fluorescence images in the brain stem were divided into 50-µ regions, and increases in fluorescence intensity for every cell in each region were normalized. The normalized intensities for all cells were averaged for each experiment. Final data represent the average of all cells in all experiments for each condition. Data are represented in Fig. 4C as P values for each 50-µ section analyzed. Comparisons of fluorescence intensity increase were performed for the third hypoxic exposure between average time dependent control and each region of the slice examined using unpaired t-test. Significant difference was set at P < 0.05.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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To examine the respiratory modulation of excitatory neurotransmission to CVNs during acute intermittent hypoxia, we employed a thick in vitro brain stem slice that retains spontaneous respiratory-related rhythms and allows simultaneous whole cell recordings of CVNs. Acute intermittent hypoxia was applied by exposing the preparation to three episodes of hypoxia of 5-min duration, followed by a 15-min recovery period after each hypoxia. Prior to hypoxia, CVNs possessed no respiratory-related modulation of excitatory neurotransmission (Fig. 1; n = 10; P > 0.05). Likewise, a single exposure of hypoxia elicited no respiratory-related change in excitatory neurotransmission to CVNs (P > 0.05). However, during the second administration of hypoxia, CVNs exhibited a significant increase in glutamatergic neurotransmission elicited during respiratory bursts (P < 0.01). Furthermore, during the third hypoxic exposure, CVNs received a more robust increase in excitatory neurotransmission simultaneous with inspiratory bursts (P < 0.01). Respiratory-related increases in excitatory neurotransmission persisted during the recovery after the second and third hypoxia; however, these increases were not significantly different from the preceding hypoxia (P > 0.05); additionally, increases observed in recovery periods were not progressively more robust (P > 0.05).
Central cardiorespiratory responses to continuous hypoxia
Intermittent hypoxia evokes physiological responses distinct from those elicited by continuous hypoxia (Baker and Mitchell 2000; Blitz and Ramirez 2002). To test whether an equivalent duration of continuous hypoxia would recruit an excitatory neurotransmission to CVNs, we exposed the preparation to 15 min of continuous hypoxia. As with a single 5-min exposure to hypoxia, CVNs received no significant increase in respiratory-related glutamatergic neurotransmission throughout the 15-min exposure to hypoxia, see Fig. 2 (P > 0.05; n = 8).
ROS mediate hypoxia-evoked increases in excitatory neurotransmission
Because ROS are strongly implicated in physiological adaptations after episodic hypoxia (Das et al. 1999; Giordano 2005; Peng and Prabhakar 2003), we tested whether the hypoxia-evoked increase in glutamatergic neurotransmission to CVNs was also mediated by a ROS-dependent mechanism. This hypothesis was examined using the stable nitroxide Tempol, which efficiently catalyzes the dismutation of free radicals (Bonini et al. 2002; Dikalov et al. 1997; Thiemermann et al. 2001). Application of Tempol (1 mM) blocked the hypoxia-evoked increase in glutamatergic neurotransmission during inspiratory activity (Fig. 3, A and B; n = 8). Neither intermittent hypoxia (n = 4; P > 0.05) nor Tempol (n = 4; P > 0.05) alone altered CVN responses to exogenous glutamate application (Fig. 3, C and D).
To test whether there are specific brain stem neurons that incrementally generate ROS, we used the indicator 5-(and-6)-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate. This compound fluoresces when converted to DCF after being oxidized by ROS (Hempel et al. 1999; Munzel et al. 2002). Time-lapse recordings revealed a significant stepwise generation of ROS during episodic hypoxia in specific neurons localized to the ventrolateral region (VLM) of the brain stem (Fig. 4, A and C; n = 5; hypoxia 3 P < 0.05). The responsive neurons were clustered and located 100–250 µ ventral to the external formation of the nucleus ambiguus. CVNs in the external formation of the nucleus ambiguus, as well as other neurons outside the VLM area, however, did not exhibit significant increases in DCF fluorescence during acute episodic hypoxia (Fig. 4C; n = 6 for each area examined; P > 0.05). Surprisingly, we found that ROS were significantly generated during the second and third hypoxias but not during reperfusion as reported in other tissues (Bolli et al. 1989; Paller and Neumann 1991; Terada 1996) (Fig. 4, A and B). In contrast, time-lapse recordings of DCF oxidation during continuous hypoxia revealed no significant increase in ROS formation either during hypoxia or throughout the recovery period (data not shown; n = 6; P > 0.05). Tempol (1 mM) effectively blocked the incremental increase in ROS formation in VLM neurons during episodic hypoxia (Fig. 4A). To determine whether the specific cells in the VLM that generate high levels of ROS during intermittent hypoxia were glutamatergic and could therefore be the source of the glutamatergic pathway recruited to CVNs during hypoxia, we colocalized these neurons with an antibody to phosphate-activated glutaminase, a reliable marker for glutamatergic neurons (Kaneko and Mizuno 1994; Kaneko et al. 1987). Quantification of the population of DCF-positive neurons that coexpressed phosphate-activated glutaminase revealed that 17.5–23.3% of the neurons that generate ROS in the VLM were glutamatergic (Fig. 4, D–F; n = 7, SE = 0.8%).
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DISCUSSION |
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20% of which are glutamatergic neurons, and may constitute the source of excitatory neurotransmission to CVNs.
Consistent with work both in vivo (Baker and Mitchell 2000; Dwinell et al. 1997; Turner and Mitchell 1997) and in vitro (Blitz and Ramirez 2002), this study establishes distinctly different physiological responses to continuous hypoxia and a comparable duration of intermittent hypoxia. As reported previously (Evans et al. 2005; Huang et al. 2005), CVNs exhibited no change in glutamatergic neurotransmission during a single exposure to hypoxia. However, during the second and third hypoxic insults, significant respiratory-related glutamatergic excitatory neurotransmission to CVNs was recruited. Whereas studies examining physiological responses to intermittent hypoxia often focus on long-term effects of this insult, i.e., the long-term increase (
90 min) in respiratory activity after intermittent hypoxia, (Blitz and Ramirez 2002; Peng and Prabhakar 2003) this study extends the physiological distinction between continuous and intermittent hypoxia to include acute alterations in excitatory neurotransmission during each hypoxic insult.
A distinct difference between intermittent and continuous hypoxia is the recovery period that occurs between each hypoxic insult with intermittent hypoxia. This study indicates ROS generation during each hypoxic period mediates the enhancement of respiratory-related excitatory neurotransmission to CVNs. However, ROS generation is reported to occur both during hypoxia and the recovery period in different systems. Indeed, ROS generation has been described during hypoxia in cardiomyocytes (Vanden Hoek et al. 1997), pulmonary artery smooth muscle cells (Killilea et al. 2000), and PC12 cells (Hohler et al. 1999). In contrast, ROS generation is reported during recovery from hypoxia in the forebrain (Lievre et al. 2000) nodose ganglion neurons (Yamamoto et al. 2003), and endothelial cells (Rieger et al. 2002; Therade-Matharan et al. 2004). Differences in the period during which ROS are generated may be due to differential expression of cellular antioxidants and/or hypoxia response elements (Li and Jackson 2002; Prabhakar 2001).
In addition to the incremental increase in DCF with intermittent hypoxia, there was also a time dependent increase in DCF fluorescence throughout the time course of experiments. Although ROS generation may occur due to the high oxygen levels of the perfusate during control conditions, a more likely explanation for time-dependent increases in DCF fluorescence in this work is photo-oxidation (Afzal et al. 2003). Tempol, a stable nitroxide that efficiently catalyzes the dismutation of free radicals, blocked the incremental increases in ROS generation observed during intermittent hypoxia without significantly altering the time-dependent increases in DCF fluorescence. This suggests the incremental increase in DCF fluorescence with intermittent hypoxia is due to ROS generation, whereas the time-dependent changes in DCF fluorescence are Tempol resistant and likely ROS-independent photo-oxidation.
In the central cardiorespiratory responses to intermittent hypoxia, ROS generation may alter neuronal excitability by activating intracellular signaling mechanisms and modulating intracellular calcium concentrations or directly altering protein or ion channel function. ROS enhance intracellular calcium concentrations by directly mobilizing calcium release from internal stores (Gen et al. 2001; Wang et al. 2004). Further, ROS oxidize amino acid residues on signaling proteins associated with calcium signaling. For example, superoxide activates protein kinase C (Knapp and Klann 2000) and extracellular-signal-regulated kinase 2 (Kanterewicz et al. 1998) and inactivates calcium/calmodulin-dependent phosphatase calcineurin (Wang et al. 1996). Oxidation of signaling proteins by ROS results in changes in protein function and may increase protein turnover by increasing the susceptibility to proteolysis (Grune et al. 1997). In addition to facilitating intracellular signaling through altering cytoplasmic proteins, ROS are also capable of modulating the activity of ionic channels. Formation of disulfide bonds by oxidation of sulfhydryl groups of cysteine residues on calcium channels reduces their activity (Chiamvimonvat et al. 1995). ROS directly activate L-type calcium channels (Wang et al. 2004), alter the open probability of potassium channels (Duprat et al. 1995), and reduce the activity of GABAA-gated chloride channels (Sah et al. 2002). One or more of these mechanisms may be functioning within the central cardiorespiratory network to recruit inspiratory-evoked excitatory neurotransmission to CVNs during intermittent hypoxia.
Intermittent hypoxia enhances respiratory-related glutamatergic neurotransmission, which is dependent on ROS generation; however, CVN responses to exogenous glutamate application are not altered during intermittent hypoxia. This indicates that intermittent hypoxia does not enhance excitatory inputs to CVNs by altering postsynaptic glutamate receptor activity or distribution but rather increasing the activity of glutamatergic neurons that synapse on CVNs. Consistent with this, we have identified specific cells localized in the ventrolateral medulla in which ROS are incrementally formed during intermittent hypoxia. Importantly, these neurons in the VLM are in the same localized region that contains, among others, respiratory neurons in the pre-Botzinger complex, the proposed kernel of respiratory rhythm (Ramirez et al. 2002; Rekling and Feldman 1998; Smith et al. 1991). Approximately 20% of the neurons have been identified as glutamatergic and therefore may be the trigger of the excitatory pathway to parasympathetic cardiac neurons that is incrementally recruited by intermittent hypoxia. Alternatively, there may be chemosensitive neurons in close proximity but nearer to the ventral surface of the brain stem that trigger the respiratory responses to hypercapnia and/or hypoxia (Weston et al. 2004). The candidate neurons identified as glutamatergic and which generate ROS in response to hypoxia may be an essential link but not the source of the initial response to hypoxia in the CNS. It would be interesting to determine in future studies if selective ablation of the ROS-generating neurons abolishes the cardiorespiratory responses to intermittent hypoxia.
Perhaps paradoxically, acute intermittent hypoxia can facilitate beneficial physiological adaptations. For example, within both the CNS and myocardium, a single hypoxic episode "preconditions" and strengthens cellular defenses to subsequent hypoxia (Lasley and Mentzer 1995; Samoilov et al. 2003). Likewise, acute intermittent hypoxia evokes the long-term facilitation of respiratory activity, which is postulated to increase upper airway patency and reduce the frequency and severity of airway obstructions (Baker and Mitchell 2000; McKay et al. 2004). However, chronic exposure to intermittent hypoxic episodes can lead to harmful physiological outcomes, depending in part on the gradual accumulation of ROS over time. For example, chronic intermittent hypoxia is associated with oxidative stress and increased neuronal death likely resulting from damage by ROS (Row et al. 2003; Xu et al. 2004). Further, within the hippocampus, low ROS concentrations (200 µM H2O2) enhance synaptic transmission (Kamsler and Segal 2003), whereas high ROS concentrations (0.5–10 mM H2O2) depress synaptic neurotransmission and plasticity (Avshalumov et al. 2000). It is likely that the central cardiorespiratory plasticity observed in this study during acute intermittent hypoxia is beneficial, evoking a bradycardia and decreasing the metabolic demands of the heart by increasing parasympathetic drive during hypoxia. It is also worth noting the results in this study were obtained in young postnatal animals, and the beneficial responses to CVNs that would evoke a bradycardia may not be as powerful in older animals. As in other systems, the chronic effects of intermittent hypoxia result in distinct and damaging effects on the autonomic nervous system; chronic intermittent hypoxia depresses baroreflex function and significantly reduces vagal efferent projections to the heart (Soukhova-O'Hare et al. 2006).
The cardiorespiratory interactions characterized in this study may also be involved in the potentially life-threatening arrhythmias associated with sleep apnea. Sleep apnea is associated with bradycardia (Garrigue et al. 2002; Gilbey et al. 1984), and this effect is at least partly mediated by increases in parasympathetic CVN activity as atropine partially blocks bradycardia both during and after apneas (Tilkian et al. 1977). Importantly, sleep apnea is postulated to result from oxidative stress accumulated during intermittent hypoxia (Prabhakar 2002). Although the cause of bradycardia during sleep apnea is unknown, this study suggests these heart rate changes may be mediated by intermittent periods of hypoxia and respiratory-related excitation of CVNs.
In summary, this study demonstrates the incremental recruitment of an excitatory neurotransmission to CVNs during acute intermittent hypoxia mediated by ROS generation. In addition, we have identified a brain stem region containing candidate glutamatergic neurons in which ROS are incrementally generated during intermittent hypoxia, suggesting this area may contain cells necessary for the critical cardiorespiratory responses to intermittent hypoxia.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Address for reprint requests and other correspondence: K.J.S. Griffioen, Dept. of Pharmacology and Physiology, George Washington University, 2300 Eye St. N.W., Washington, D.C. 20037 (E-mail: kgriff{at}gwu.edu)
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