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J Appl Physiol 88: 1496-1508, 2000;
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Vol. 88, Issue 4, 1496-1508, April 2000

HIGHLIGHTED TOPICS
Blunted respiratory responses to hypoxia in mutant mice deficient in nitric oxide synthase-3

David D. Kline1, Tianen Yang1, Daniel R. D. Premkumar1, Agnes J. Thomas2, and Nanduri R. Prabhakar1

Department of 1 Physiology and Biophysics and 2 Medicine, School of Medicine, Case Western Reserve University, Cleveland, Ohio 44106


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

In the present study, the role of nitric oxide (NO) generated by endothelial nitric oxide synthase (NOS-3) in the control of respiration during hypoxia and hypercapnia was assessed using mutant mice deficient in NOS-3. Experiments were performed on awake and anesthetized mutant and wild-type (WT) control mice. Respiratory responses to 100, 21, and 12% O2 and 3 and 5% CO2-balance O2 were analyzed. In awake animals, respiration was monitored by body plethysmography along with O2 consumption (VO2) and CO2 production (VCO2). In anesthetized, spontaneously breathing mice, integrated efferent phrenic nerve activity was monitored as an index of neural respiration along with arterial blood pressure and blood gases. Under both experimental conditions, WT mice responded with greater increases in respiration during 12% O2 than mutant mice. Respiratory responses to hyperoxic hypercapnia were comparable between both groups of mice. Arterial blood gases, changes in blood pressure, VO2, and VCO2 during hypoxia were comparable between both groups of mice. Respiratory responses to cyanide and brief hyperoxia were attenuated in mutant compared with WT mice, indicating reduced peripheral chemoreceptor sensitivity. cGMP levels in the brain stem during 12% O2, taken as an index of NO production, were greater in mutant compared with WT mice. These observations demonstrate that NOS-3 mutant mice exhibit selective blunting of the respiratory responses to hypoxia but not to hypercapnia, which in part is due to reduced peripheral chemosensitivity. These results support the idea that NO generated by NOS-3 is an important physiological modulator of respiration during hypoxia.

nitric oxide; nitric oxide synthase enzyme; nitric oxide synthase deficiency; carotid body


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

NITRIC OXIDE (NO) is generated by the enzyme nitric oxide synthase (NOS). Three isoforms of NOS have been characterized, including neuronal (NOS-1), inducible (NOS-2), and endothelial (NOS-3) (36). Of the three isoforms, NOS-1 and NOS-3 are constitutively expressed and are responsible for basal generation of NO. It is being increasingly recognized that NO is involved in many physiological processes, including neurotransmission in the nervous system, control of blood pressure, and immune responses (26). Recent evidence also suggests that NO modulates breathing during hypoxia.

The NOS-1 isoform is expressed in several neuronal structures associated with respiration. For instance, NOS-1-like immunoreactivity is localized in the sensory nerve fibers innervating the carotid body, the primary sensory organ that monitors arterial oxygen (18, 42, 56), as well as in the neurons in the nucleus tractus solitarius (NTS), the region of the brain stem that receives and integrates afferent inputs from the peripheral chemoreceptors (20, 54). NOS inhibitors augment the respiratory responses to hypoxia, an effect that appears to be due to blockade of NOS in the carotid body and central neural structures that regulate breathing during hypoxia (17, 52). NOS inhibitors also augment the basal sensory discharge of the carotid body chemoreceptors during normoxia (42, 57) and enhance the sensory response to hypoxia (9, 52, 57). On the other hand, NO donors (9, 57) inhibit the sensory discharge. Microinjections of NO donors into the NTS neurons, on the other hand, have varied effects on breathing. Reported responses include inhibition (55) and excitation (37) of breathing. Recently, we have demonstrated that mutant mice deficient in the NOS-1 protein (NOS-1 mutant mice) exhibit augmented respiratory responses to hypoxia, which are due, in part, to enhanced peripheral chemosensitivity (28). Together, these studies provide compelling evidence that NO generated by NOS-1 is a potential modulator of breathing during hypoxia.

Although NOS-1 is predominately confined to neuronal structures, NOS-3 is primarily localized to the endothelium of many blood vessels, including the vasculature supplying the carotid body and cerebral blood vessels (33, 56). NO generated from NOS-3 regulates blood flow by way of controlling vascular tone (14, 53). Consequently, NO produced from NOS-3 may modulate carotid body activity via regulation of blood flow and subsequent changes in tissue PO2 within the chemoreceptors (18, 41, 57). NO generated by NOS-3 may also regulate brain stem neuronal activity by altering the blood flow to the neurons (14). Therefore, the purpose of the present study is to determine the specific contribution of NO generated from NOS-3 in the control of breathing during hypoxia and hypercapnia. However, to delineate the selective contribution of NOS-3, the use of NOS inhibitors is inadequate because many of these compounds cannot distinguish between the NOS-1 and NOS-3 isoforms. The development of mutant mice deficient in the NOS-3 protein offers an excellent animal model for assessing the importance of NO generated by NOS-3 independent of NOS-1. Therefore, in the present study, we examined the respiratory responses to hypoxia, as well as to hypercapnia, in NOS-3 mutant mice. The results demonstrate that the respiratory responses to hypoxia, but not to hypercapnia, are markedly attenuated in mice deficient in NOS-3. The blunted respiratory responses to hypoxia appear, in part, to be due to reduced drive from the peripheral chemoreceptors.


    METHODS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

General Preparation of Animals

Experiments were performed on age-matched wild-type (WT) and NOS-3 mutant mice of either sex. The average weights of the animals were as follows: WT mice were 22.6 ± 0.6 g and NOS-3 mutant mice were 23.5 ± 0.5 g (P > 0.05, t-test). Mutant mice were obtained from Dr. P. L. Huang (22). Hybrids of the 129/SV and C57BL/6 strains of mice, the parental strains of the mutant mice, were used as WT controls. Experiments were performed on anesthetized, as well as awake, unrestrained mice.

Protocols were approved by the Institutional Animal Care and Use Committee of Case Western Reserve University. Animals were anesthetized with intraperitoneal injections of urethan (1.2 g/kg; Sigma Chemical). The choice of anesthesia was based on reports that acid-base status is well maintained under urethan in experimental animals (7). Supplemental doses of anesthesia (15% of initial dose) were given when corneal reflexes and responses to toe pinch persisted.

Routine surgical procedures included tracheal intubation and catheterization of the femoral artery and vein. The femoral artery catheter was used to monitor arterial blood pressure, whereas systemic administration of fluids and/or drugs was accomplished through the femoral vein catheter. Blood pressure was measured by a Grass pressure transducer (model PT300). Animals were allowed to breathe spontaneously. Core body temperature was monitored by a rectal thermistor probe and was maintained at 37 ± 1°C by a heating pad. At the end of the experiment, the animal was killed by intracardiac injection (0.1 ml) of euthanasia solution (Beuthanasia-D Special, Schering-Plough Animal Health, Kenilworth, NJ).

Measurements of Respiratory Variables

In anesthetized animals, integrated efferent phrenic nerve activity (<LIM><OP>∫</OP></LIM>Phr) was monitored as an index of central respiratory neuronal output. For this purpose, the phrenic nerve was isolated unilaterally at the level of the cervical 3 and 4 spinal segments. The nerve was cut distally and placed on bipolar stainless steel electrodes. The electrical activity was filtered (band pass 0.3-1.0 kHz), amplified, and passed through Paynter filters (time constant of 100 ms; CWE, Ardmore, PA) to obtain a moving average signal.

In unanesthetized animals, respiration was monitored by whole body plethysmograph as described previously (28). Briefly, animals were placed in a 600-ml Lucite chamber containing an inlet port for the administration of test gases. The animal chamber, as well as a reference chamber, was connected to a high-gain differential pressure transducer (Valydine MP45, Validyne Engineering, Northridge, CA). As the animal breathed, small changes in pressure were converted to signals representing tidal volume (VT) (5). The signals were amplified (BMA 830; CWE) and recorded on a strip-chart recorder (Dash 10; Astro-Med, West Warwick, RI). The signals were also stored in a computer with respiratory acquisition software for analysis off-line. O2 consumption (VO2) and CO2 production (VCO2) was determined by the open-circuit method (46) using Beckman OM-14 and LB-2 analyzers.

Measurements of cGMP Levels by Competitive Enzyme Immunoassay

Anesthetized mice (n = 9 each of WT and mutant) were exposed to 100, 21, or 12% inspired oxygen for 5 min. At the end of the gas challenge, brain stems were removed and placed in 50 mM sodium acetate (pH 4.0) containing the phosphodiesterase inhibitor IBMX (3 µg/ml). Brain stems were frozen in liquid nitrogen and kept at -80°C until further analysis. Tissues were thawed, minced, and sonicated into 50 mM sodium acetate (pH 4.0) containing IBMX. The homogenate was centrifuged at 10,000 g for 15 min at 4°C. Acetylated cGMP levels were determined in 50 µl of supernatant by a cGMP enzyme immunoassay kit (EIA; Cayman Chemical, Ann Arbor, MI). Protein was assayed using a protein analyzing kit (Bio-Rad Technologies) with BSA as standard. All assays were in duplicate, and the values of cGMP are expressed as picomoles per milligram of protein.

Carotid Body Morphology

Mice (n = 3 WT and n = 3 NOS-3 mutant; 6 carotid bodies in each group) were anesthetized with urethan (1.2 g/kg ip). After the chest was opened with a midline incision, a 25-gauge needle was inserted into the left ventricle for perfusion. To allow drainage of the fluid, an incision was made in the right atrium. Animals were perfused with heparinized PBS, pH 7.4, for 10 min at 10 ml/min with a peristaltic pump (Masterflex, Cole-Parmer). This was followed by freshly prepared 4% paraformaldehyde-PBS for an additional 10 min.

Carotid artery bifurcations were removed and immersed in 4% paraformaldehyde-PBS for postfixation for 1 h at 4°C. The tissue was washed three times for 10 min in PBS and cryoprotected in 30% sucrose-PBS at 4°C for 24 h. Specimens were dissected free of excess connective tissue, frozen in Tissue Tek (OCT; VWR), and stored at -80°C until they were sectioned. The specimens were cut serially at 15 µm on a cryostat (Bright Instruments) and mounted on Vectabound (Vector Laboratories, Burlingame, CA)-treated slides.

Tissue sections were washed three times for 15 min in PBS. After they were washed, sections were exposed to 20% normal goat serum (NGS)-0.2% Triton X-100-PBS for 2 h. Endogenous biotinylated proteins were blocked with avidin and biotin (Vector Laboratories). Sections were incubated at 4°C for 16 h with either anti-chromogranin A (CGA, 1:1,000; Instar, Stillwater, MN), anti-NOS-3 (1:50; Transduction Laboratories, Lexington, KY), or anti-tyrosine hydroxylase (TH, 1:100; Pel-Freeze, Rogers, AK) in 1% NGS-0.2% Triton X-100-PBS. CGA- and TH-positive staining was used to identify glomus cells (29, 58). Parallel experiments were performed on sections without the primary antibody. After a 15-min PBS wash (3 times), sections were incubated for 120 min with biotinylated anti-rabbit IgG (1:200; Vector Laboratories) in 1% NGS-0.2% Triton X-100-PBS at room temperature. Immunostaining was visualized by the Vectastain Elite avidin-biotinylated enzyme complex method (Vector Laboratories) using diaminobenzidine peroxidase substrate. CGA- and TH-immunoreactive cells were counted manually in each tissue section (n = 5 sections/carotid body, n = 6 carotid bodies from each group). Data are presented as means ± SE of positively stained cells per tissue section, and statistical significance was evaluated by unpaired t-test.

Experimental Protocols

Anesthetized mice. The effects of three levels of inspired O2 (100, 21, and 12% O2-balance nitrogen) on efferent phrenic nerve activity were determined in anesthetized, spontaneously breathing mice (n = 11 WT, n = 10 NOS-3 mutant). Baseline respiratory activity and blood pressure were monitored while the animals breathed 100% O2. Subsequently, inspired gas was switched to 21% followed by 12% O2. Each gas challenge was maintained for 5 min. After 12% O2, inspired air was switched back to 100% O2. Gases were administered through a needle placed near the tracheal cannula, and gas flow was controlled by a flow-meter.

The blood volume of an average mouse weighing 20 g is ~1.2 ml (25). Because of this limitation, in a given experiment, repeated withdrawal of arterial blood (200 µl/sample) was found to be lethal to the animal. Therefore, in parallel experiments on WT and mutant mice (n = 10 WT and 12 NOS-3 mutant mice), arterial blood gases were analyzed at the end of 100, 21, and 12% O2 gas challenge. Arterial blood was sampled via a catheter placed in the descending abdominal aorta near the iliac region for quick removal of the blood sample. Care was taken that blood flow to major organs, such as the liver and kidney, was not compromised. Arterial blood PO2, PCO2, and pH were analyzed by a blood-gas analyzer (Radiometer Instruments).

Unanesthetized mice. In the experiments involving unanesthetized, unrestrained mice, all measurements were made between 9:00 AM and 1:00 PM. Animals were placed in the plethysmograph chamber containing aspen bedding and allowed to acclimate to the environment for 60 min while room air flowed through the chamber. Subsequently, animals were challenged with varying levels of inspired O2 or CO2 as described below.

In the first group of experiments, mice (n = 8 WT and n = 8 NOS-3 mutant) were exposed to 100, 21, and 12% O2-balance nitrogen. Each gas challenge was given for 5 min. The protocols were repeated three times in each animal, with a 20-min interval between each protocol. VO2 and VCO2 were measured at the end of each 5-min gas challenge.

Respiratory responses to hyperoxic hypercapnia were determined in the second group of experiments (n = 12 WT and n = 8 NOS-3 mutant). Mice were allowed to breathe 100% O2 for 5 min followed by 3 and 5% CO2-balance oxygen. The protocols were repeated three times, with a 20-min interval between each protocol.

Peripheral Chemoreceptor Sensitivity

Hyperoxic challenge. The effects of brief hyperoxic challenge on respiration (12) were examined on anesthetized WT (n = 6) and NOS-3 mutant (n = 6) mice. Baseline respiration was recorded while animals breathed 12% O2 for 45 s; 100% O2 was added to the inspired air for 20 s. Respiratory rate (RR) was analyzed for 20 s during 12% O2 and during the last 15 s of hyperoxia. Breathing during the initial 5 s of hyperoxia was excluded from analysis because of the dead space of the tubing.

Sodium cyanide. The effects of intravenous administration of sodium cyanide (Fisher Scientific) on respiration were examined in anesthetized WT and NOS-3 mutant mice (n = 7 each) breathing room air. The dose of cyanide was 50 µg/kg. The volume of the injectate was 50 µl of saline (0.9% NaCl). This dose was based on the dose-response curve reported by us previously (28). The same volume of saline (50 µl) without cyanide served as a control. Stock solutions of cyanide were prepared fresh before each experiment. Respiration was measured 1 min before and 1 min immediately after the injection of sodium cyanide.

Data Analysis

In anesthetized mice, the following variables were analyzed: RR (number of phrenic bursts per minute), amplitude of the <LIM><OP>∫</OP></LIM>Phr [arbitrary units (AU)], and minute neural respiration (MNR, AU/min, RR × <LIM><OP>∫</OP></LIM>Phr). Respiratory variables (RR and <LIM><OP>∫</OP></LIM>Phr) and blood pressure (in mmHg) were averaged over a 5-min period of each gas challenge. Amplitude of the <LIM><OP>∫</OP></LIM>Phr and MNR were normalized to the body weight of the animal.

The following variables were analyzed in unanesthetized mice: RR (breaths/min), inspiratory VT (µl), and minute ventilation (VE; ml/min, RR × VT). Respiratory variables (RR and VT) were averaged for 15 consecutive breaths over 5 min of inspired O2 and CO2 challenge. Sighs or sniffs were excluded in the analysis. VT and VE were normalized to the body weight of the animal. Metabolic variables were measured at the end of each 5-min inspired O2 challenge. Each data point in a given animal, for a given gas challenge, represents the average of three trials.

All results are expressed as means ± SE. Paired t-tests were used to evaluate if each animal responded with significant increases in respiration during 21 and 12% O2 compared with 100% O2. Significance of changes between WT and mutant mice was determined by an unpaired t-test. P values <0.05 were considered significant.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Respiratory Responses to Changes in Inspired Oxygen in Anesthetized Mice

An example illustrating the effect of changing inspired oxygen on respiration in an anesthetized WT and a NOS-3 mutant mouse is shown in Fig. 1A. As can be seen, lowering the inspired oxygen from 100 to 21 and 12% O2 resulted in a marked stimulation of respiration in WT mice, whereas mutant mice responded only with a modest increase in breathing. Average results are summarized in Table 1. The basal RR was lower in mutant mice breathing 100% O2. In response to 21% O2, respiration increased significantly in WT mice, which was due to increases in the amplitude of <LIM><OP>∫</OP></LIM>Phr as well as RR. During 12% O2, only <LIM><OP>∫</OP></LIM>Phr increased. As a consequence, MNR was significantly augmented during 21 and 12% O2 in WT mice. In contrast, respiration was unaffected in mutant mice subjected to either 21 or 12% O2 (P < 0.05, t-test; Fig. 1B).


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  		Fig. 1.   Respiratory responses to varying levels of inspired oxygen in anesthetized wild-type (WT) and nitric oxide synthase (NOS)-3 mutant mice. A: representative tracings of an experiment illustrating respiratory responses during 100, 21, and 12% inspired oxygen in an anesthetized WT and NOS-3 mutant mouse. <LIM><OP>∫</OP></LIM>Phr, integrated phrenic nerve activity; 100% O2, 21% O2, and 12% O2 indicate inspired oxygen levels. Respiration [respiratory rate (RR) and <LIM><OP>∫</OP></LIM>Phr] increased in response to 21% O2 in WT mice. Respiration continued to increase in the WT mouse during 12% O2 due to increases in <LIM><OP>∫</OP></LIM>Phr. Respiration during 21 and 12% O2 did not increase dramatically in NOS-3 mutant mice. B: comparison of the respiratory responses during 21 and 12% O2 in anesthetized WT and mutant mice. Results are expressed as percentage of 100% O2 and are presented as means ± SE of WT (n = 11) and mutant (n = 10) mice. * P < 0.05 (t-test). Note that the respiratory responses [RR, <LIM><OP>∫</OP></LIM>Phr, and minute neural respiration (MNR)] were greater in WT than in NOS-3 mutant mice during 21 and 12% O2.


                              
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  		Table 1.   Changes in respiratory variables in anesthetized and unanesthetized wild-type and NOS-3 mutant mice during 100, 21, and 12% O2

Changes in Arterial Blood Pressure and Blood Gases

The changes in arterial blood pressure were analyzed during three levels of inspired oxygen in both groups of mice. Basal arterial blood pressure was significantly higher in NOS-3 mutant than in WT mice [65 ± 5 mmHg for NOS-3 mutant (n = 10) vs. 53 ± 4 mmHg for WT (n = 11); P < 0.05, t-test]. Blood pressure decreased in both groups of mice during 21 and 12% O2. The magnitude of the decrease in blood pressure was not statistically different between both groups of mice (P > 0.05, t-test; Table 2). Arterial PO2 and pH levels were comparable between WT and NOS-3 mice at any given level of inspired oxygen, except that arterial PCO2 during 100% O2 was significantly higher in NOS-3 compared with WT mice (P < 0.05, t-test).

                              
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  		Table 2.   Wild-type and NOS-3 mutant mice arterial blood gases and arterial blood pressure during 100, 21, and 12% O2

Respiratory Responses to Changes in Inspired Oxygen in Awake Mice

The results obtained in anesthetized mice demonstrate that NOS-3 mutant mice do not respond with increases in respiration during hypoxia. To determine whether the blunting of respiratory stimulation by hypoxia is due to anesthesia, experiments were performed on awake mice and respiration was recorded by plethysmographic techniques. As shown in the example depicted in Fig. 2A, 12% O2 stimulated breathing in WT mice, primarily due to increases in RR. In contrast, NOS-3 mutant mice responded only with a modest increase in RR during 12% O2. Average results are summarized in Table 1. As in anesthetized mice, basal RR during breathing 100% O2 was significantly lower in mutant mice. In response to 21% O2, respiration was unaffected in both groups of mice; however, there was a significant increase in VE in both groups of animals during 12% O2. In WT mice, the increase in respiration was due to increases in RR as well as VT. On the other hand, only RR increased in NOS-3 mutant mice. The relative increases in RR and VE during 12% O2 were significantly greater in WT than in NOS-3 mutant mice (P < 0.05, t-test; Fig. 2B).


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  		Fig. 2.   Respiratory responses to varying levels of inspired oxygen in unanesthetized (awake) WT and NOS-3 mutant mice. A: representative tracings of respiratory responses to 3 levels of inspired oxygen in an unanesthetized WT and mutant mouse; 100% O2, 21% O2, and 12% O2 indicate inspired oxygen levels. VT, tidal volume. B: comparison of the respiratory responses during 21 and 12% O2 in WT and mutant mice. Note the greater increases in respiration [RR and minute ventilation (VE)] in WT mice during 12% O2 compared with NOS-3 mutant mice. Results are expressed as percentage of 100% O2 and are presented as means ± SE of 8 each of WT and NOS-3 mutant mice. * P < 0.05 (t-test).

Comparison of Changes in VO2 and VCO2

It has been well established that hypoxia causes a reduction in body metabolism, as evidenced by changes in VO2 and VCO2 (15). To assess whether changes in these variables contributed to the ventilatory responses to hypoxia, VO2 and VCO2 were determined in the same experiments as above. The results are summarized in Fig. 3. Under basal conditions (i.e., 100% O2), the values of VO2 were similar between both groups of mice (2.6 ± 0.2 ml/min for WT vs. 2.9 ± 0.2 ml/min for mutant; P > 0.05, t-test). VO2 decreased significantly in both groups of mice in response to 21% O2 (P < 0.05, paired t-test); however, VO2 decreased to a greater extent in NOS-3 mutant than in WT mice (P < 0.05, t-test). VO2 continued to decrease during 12% O2 in WT and NOS-3 mice (P < 0.05, paired t-test), yet the relative changes in VO2 during 12% O2 were comparable between both groups of mice (P > 0.05, t-test; Fig. 3A). Likewise, there was no significant difference in VCO2 values under basal conditions (i.e., 100% O2) between WT and NOS-3 mice (1.4 ± 0.1 ml/min for WT vs. 1.1 ± 0.07 ml/min for mutant; P > 0.05, t-test). VCO2 was unaffected during 21% O2 in both groups of mice (P > 0.05, paired t-test). There was a significant reduction in VCO2 during 12% O2 in WT and NOS-3 mutant mice (P < 0.05, paired t-test); however, the magnitude of decrease was comparable (P > 0.05, t-test; Fig. 3B).


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  		Fig. 3.   Comparison of changes in O2 consumption (VO2; A) and CO2 production (VCO2; B) during 21 and 12% O2 in unanesthetized WT and NOS-3 mutant mice. Results are presented as percentage of 100% O2. Data were analyzed from the same mice as in Fig. 2 and are presented as means ± SE. * P < 0.05 (t-test) between both groups of mice. Note that the reductions in VO2 and VCO2 were comparable between WT and NOS-3 mice during hypoxia. NS, not significant.

Respiratory Responses to Hypercapnia in WT and NOS-3 Mutant Mice

Respiratory responses to two levels of hyperoxic hypercapnia (3 and 5% CO2-balance O2) were recorded in WT and NOS-3 mutant mice. These experiments were performed on unanesthetized mice because respiratory responses to CO2 were markedly suppressed in mice under urethan anesthesia (28). An example showing the respiratory responses to 3 and 5% CO2 in a WT and NOS-3 mutant mouse is presented in Fig. 4A. As can be seen, both mice responded with increased breathing during 3 and 5% CO2. The augmentation of respiration in both groups of mice was due to significant increases in RR as well as VT. However, there were no significant differences in the magnitude of respiratory responses to hypercapnia between both groups of mice (P > 0.05, t-test; Fig. 4B).


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  		Fig. 4.   Respiratory responses to hyperoxic hypercapnia (3 and 5% CO2-balance oxygen) in unanesthetized WT and NOS-3 mutant mice. A: representative tracings of respiratory responses to 2 levels of inspired CO2 in an unanesthetized WT and NOS-3 mutant mouse; 3% CO2 and 5% CO2 indicate inspired CO2 (balance O2) levels. Note that both WT and mutant mice responded to 3 and 5% CO2 in a comparable manner. B: comparison of the respiratory responses during 3 and 5% CO2 in WT and mutant mice. Results are expressed as percentage of 100% O2 and are presented as means ± SE of 8 each of WT and mutant mice. Note that the respiratory responses were similar between both groups of mice (P > 0.05, t-test).

Peripheral Chemoreceptor Sensitivity in NOS-3 Mutant and WT Mice

The results described thus far indicate that NOS-3 mutant mice exhibit reduced respiratory responses to hypoxia but not to hypercapnia. The blunting of the hypoxic responses may in part be due to changes in peripheral chemoreceptor sensitivity. The following experiments were performed on anesthetized mice to test this possibility.

Effects of brief hyperoxia on respiration (Dejour's test). The decrease in RR in response to brief hyperoxia (100% O2) is commonly used as an index of peripheral chemoreceptor sensitivity (12). If carotid body sensitivity is altered in NOS-3-deficient mice, it should be reflected in the magnitude of decrease in the RR during hyperoxia. To test this possibility, we compared the respiratory response to brief hyperoxia between NOS-3-mutant and WT mice. O2 (100%) was added to the inspired air for 20 s while the animals breathed 12% O2. As shown in Fig. 5A, hyperoxia resulted in a prompt reduction in RR in WT mice. In contrast, the respiratory response to brief hyperoxia was nearly absent in NOS-3 mutant mice (Fig. 5A). Averaged results showed that, in response to brief hyperoxia, RR decreased by 16.0 ± 1.5% in WT mice, whereas it was reduced only by 9.0 ± 1.0% in NOS-3 mutant mice (WT vs. mutant, P < 0.05, t-test; Fig. 5B).


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  		Fig. 5.   Respiratory responses to brief hyperoxia in anesthetized WT and NOS-3 mutant mice. A: representative tracings of an experiment illustrating the effect of brief hyperoxia (Dejour's test) on phrenic nerve activity in an anesthetized, spontaneously breathing WT and NOS-3 mutant mice. Animals breathed 12% O2. At arrow, 100% O2 was added to the inspired air; 100% O2 caused prompt decreases in respiration in both mice, but the response was more pronounced in WT mice. B: averaged data for changes in RR. RR was analyzed during the last 15 s of hyperoxia (for details, see text). Changes are expressed as percentage of 12% O2 controls and are shown as means ± SE from 6 WT and 6 NOS-3 mutant mice. * P < 0.05 (t-test) between WT and NOS-3 mutant mice. Note the greater decreases in RR by hyperoxia in WT mice compared with NOS-3 mutant mice.

Effects of sodium cyanide on respiration. In another series of experiments, we analyzed the effects of systemic administration of cyanide (50 µg/kg), a potent stimulant of the carotid body, on breathing. As shown in Fig. 6A, cyanide resulted in a prompt respiratory stimulation in both groups of mice, primarily due to increases in RR. The respiratory stimulation was more pronounced in the WT mouse than in the NOS-3 mutant mouse. Average results showed that increases in RR in response to cyanide were significantly greater in WT than in mutant mice (P < 0.05, t-test; Fig. 6B). Administration of the same volume of saline (i.e., vehicle) had no effect on respiration. Bilateral sectioning of the carotid sinus nerves abolished cyanide-induced stimulation of breathing, indicating that respiratory stimulation is due to excitation of the carotid body chemoreceptors.


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  		Fig. 6.   Respiratory responses to sodium cyanide in anesthetized WT and NOS-3 mutant mice. A: representative tracings of an experiment illustrating the effect of systemic administration of sodium cyanide on phrenic nerve activity in anesthetized, spontaneously breathing WT and NOS-3-deficient mice. Cyanide (CN; 50 µg/kg iv) was injected at the arrow. Sodium cyanide caused prompt increases in RR in both mice, but the response was more pronounced in WT mice. B: averaged data for changes in RR. Results are expressed as percentage of preinjection controls and are shown as means ± SE from 7 animals in each group. * P < 0.05 (t-test). Systemic administration of sodium cyanide resulted in a substantial increase in RR in WT mice, whereas it did not in mutant mice.

Carotid Body Morphology in NOS-3 Mutant Mice

The following histochemical experiments were performed to determine whether decreased carotid body sensitivity is due to a reduced number of glomus cells, the putative O2-sensing cells in the peripheral chemoreceptor tissue. Glomus cells were identified by the presence of CGA, a synaptic vesicle protein, or TH, the rate-limiting enzyme in catecholamine synthesis. Both are well-established markers of glomus cells (29, 58). Examples depicting CGA-like immunoreactivity in the carotid bodies of WT and NOS-3 mutant mice are shown in Fig. 7, C and D. Figure 7 shows that carotid bodies from both groups of mice displayed CGA-positive cells. Quantitative analysis revealed that NOS-3 mutant mice had 32% more CGA-positive cells than WT mice (94 ± 7 in WT vs. 124 ± 22 CGA-positive cells/section in mutant, P > 0.05, t-test; Fig. 7E) The results were essentially the same when TH was used as a glomus cell marker (95 ± 2 in WT vs. 128 ± 12 TH-positive cells/section in mutant, P > 0.05, t-test; Fig. 7E). These results demonstrate that the number of glomus cells were not decreased in mutant mice. Rather, they showed a tendency toward an increase, compared with carotid bodies from WT mice.


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  		Fig. 7.   Carotid body morphology in NOS-3 mutant mice. A and B: immunolocalization of the NOS-3 protein in a WT mouse (A) and a NOS-3 mutant mouse (B). Carotid artery of WT mouse exhibited NOS-3-positive staining within the endothelial lining. Such staining was not observed in NOS-3 mutant mice. C and D: examination of glomus cells, as evidenced by chromogranin A (CGA)-like immunoreactivity, in a WT mouse (C) and a NOS-3 mutant mouse (D). Note that glomus cells were observed in both groups of mice. Scale bars = 50 µm. E: comparison of WT and mutant mouse carotid body morphology. Note there was a tendency for a greater number of glomus cells, as evidenced by greater CGA- and tyrosine hydroxylase-like immunoreactivity, in NOS-3 mutant mice compared with WT mice.

Changes in cGMP Levels in the Brain Stem in Response to Changes in Inspired Oxygen

To determine whether the blunted respiratory responses to acute hypoxia in mutant mice is due, in part, to reduced NO production in the brain stem, we monitored cGMP levels as an index of NO generation (28). Brain stems were removed from anesthetized WT and NOS-3 mutant mice exposed to three levels of inspired oxygen for 5 min, and cGMP levels were analyzed as described in METHODS. The results are summarized in Fig. 8. In response to lowering the inspired oxygen from 100 to 21 and 12% O2, there was a progressive increase in cGMP levels in both groups of mice. The magnitude of cGMP increases during 12% O2, however, was greater in NOS-3 mutant mice compared with WT mice (P < 0.05, t-test).


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  		Fig. 8.   Analysis of cGMP levels in the brain stems of WT and NOS-3 mutant mice during 3 levels of inspired oxygen (100, 21, and 12% O2) in WT and mutant mice. Data are presented as means ± SE from 9 animals in each groups. Note that, in both groups of mice, cGMP levels increased during 21 and 12% O2. However, cGMP levels were greater (* P < 0.05, t-test) in NOS-3 mutant than in WT mice during 12% O2.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

In the present study, we examined the role of endogenous NO generated by the endothelial NOS isoform, NOS-3, in the control of respiration during hypoxia and hypercapnia. For this purpose, we used mutant mice deficient in NOS-3. The advantage of using mutant mice is that it allows the study of NO produced by this isoform independently of other NOS isoforms (i.e., NOS-1 and NOS-2). Our results provide evidence that NO derived from NOS-3 modulates respiration during hypoxia but not during hypercapnia. Furthermore, the results suggest reduced peripheral chemoreceptor sensitivity in NOS-3 mutant mice.

Respiratory Responses to Hypoxia, but not to Hypercapnia, Are Blunted in NOS-3 Mutant Mice

Experiments were conducted on both unanesthetized as well as anesthetized mice. The advantages of using unanesthetized mice are that it alleviates the effects anesthesia may have on breathing and allows us to monitor changes in body metabolism that are known to occur during hypoxia (15). Changes in blood pressure and blood gases, however, could not be monitored in unanesthetized preparations due to technical restraints, but this was accomplished in anesthetized animals. It is clear from both preparations that the respiratory responses to 21 and 12% O2 are significantly blunted in mutant mice deficient in NOS-3 compared with WT control mice. Neither the changes in arterial blood pressure nor the changes in blood gases seem to account for the reduced response to hypoxia because the magnitude of changes in these variables during 21 and 12% O2 was comparable between both groups of mice. However, during hyperoxia, NOS-3 mutant mice hypoventilated compared with control mice, and this hypoventilation is reflected in higher arterial PCO2 values (see Table 2).

Previous studies have suggested that endogenously generated NO affects VO2 by inhibiting mitochondrial cytochrome-c oxidase (6). It is also known that changes in body metabolism influence the ventilatory response to hypoxia (15). Therefore, it is possible that the blunted ventilatory response to hypoxia seen in NOS-3 mutant mice is secondary to alterations in body metabolism. However, the following lines of evidence indicate that this may not be the case. First, the resting VO2 values are comparable between NOS-3 mutant and WT mice. This is not surprising because NO, once released from the endothelial cells, rapidly binds to myoglobin and hemoglobin (10). Consequently, levels of NO near the mitochondria may not be adequate to affect cytochrome-c oxidase activity. The findings by Crystal et al. (11) support this idea: they showed that whole body metabolism is unaffected by NOS inhibition. Second, changes in VO2 and VCO2 were comparable between both groups of mice during hypoxia (12% O2). In fact, when VE was normalized to their VCO2 (VE/VCO2), the magnitude of respiratory stimulation during hypoxia was found to be still significantly less in mutant compared with that in WT mice (151 ± 11% in NOS-3 mutant vs. 177 ± 13% in WT during 12% from 100%; P < 0.05, t-test). Thus changes in metabolic variables seem not to account for the blunted ventilatory response to hypoxia.

Unlike hypoxia, the ventilatory responses to CO2 were comparable between NOS-3 mutant and WT mice. These finding are similar to those observed in NOS-1 mutant mice (28) as well as those reported with NOS inhibitors in rats, suggesting that NO does not play a significant role in the hypercapnia-induced hyperventilation or hypometabolism (4, 17, 20, 39). However, Teppema et al. (51) reported that, after systemic administration of NOS inhibitors, the ventilatory response to CO2 was reduced in anesthetized cats (whereas our experiments were conducted in mice). Recent studies have shown that the density of NOS-positive neurons in the NTS and ventrolateral medulla (the neuronal substrate associated with CO2 responses) varies between the cat and mouse (60). Therefore, the discrepancy between the two studies could be due to species difference. Alternatively, the attenuated ventilatory response to CO2 observed by Teppema et al. (51), who used normoxic CO2, might be secondary to CO2-O2 interactions at the peripheral chemoreceptors. On the other hand, we used hyperoxic CO2, for which any interaction between O2 and CO2, if any, would be minimal. Nonetheless, the present investigation, together with previous observations (28), suggests that endogenously generated NO either from NOS-1 or NOS-3 does not modulate breathing during hyperoxic hypercapnia. These observations further indicate that the absence of respiratory stimulation by hypoxia observed in NOS-3-deficient mice is not due to the inability of the respiratory apparatus to respond to excitatory stimuli. Rather, it may be due to reduced peripheral chemoreceptor sensitivity and/or altered processing of the chemoreceptor inputs at the brain stem neurons.

Evidence for Blunted Carotid Body Sensitivity: Possible Mechanisms

Peripheral chemoreceptors, especially the carotid bodies, are necessary for stimulation of breathing during hypoxia. It has been established that NOS-1 and NOS-3 are present in the carotid body and that NO is inhibitory to chemosensory activity (41). Endothelium-derived NO may play a role in the regulation of blood flow to the chemoreceptor tissue and therefore may modulate carotid body activity. Two lines of evidence from the present experiments support the idea that the peripheral chemoreceptor, especially the carotid body sensitivity, is reduced in NOS-3 mutant mice. First, the respiratory response to cyanide, a potent stimulant to the carotid body, is reduced in mutant mice. Second, the magnitude of respiratory depression to brief hyperoxia (i.e., Dejour's test, from hypoxia) is less pronounced in mutant mice. Previous studies have reported that the ventilatory responses to hypoxia and to cyanide were unaffected after systemic administration of a putative NOS-1 inhibitor, whereas a general NOS inhibitor potentiated the stimulatory effects of hypoxia and cyanide (16, 17). On the basis of these studies, it has been proposed that NO from NOS-3 is inhibitory to the carotid body activity. However, our results showed a clear blunting of the respiratory response to hypoxia and cyanide in mutant mice deficient in NOS-3. It may be that the relative contribution of NO derived from NOS-3 varies from mice to rats. Alternatively, the acute physiological consequences of blockade of NOS-3 may differ from chronic deficiency of the NOS-3 protein (see below).

How might NO from NOS-3 affect chemoreceptor activity? Because NOS-3 is distributed primarily in the blood vessels and NO is a potent vasodilator, it has been proposed that NO regulates chemoreceptor activity by way of regulating blood flow to the carotid body (41). Acute blockade of NOS-3 stimulates chemoreceptor activity presumably due to vasoconstriction and reduced local PO2 in the chemoreceptor tissue. In mutant mice, the gene encoding NOS-3 protein is functionally defective since birth. As a consequence, it is to be expected that the carotid bodies receive less blood flow and are subjected to persistent tissue hypoxia. It is known that persistent tissue hypoxia, such as that which occurs in the later stages of chronic hypertension, renders the carotid body insensitive to changes in oxygen (43-45, 49). Consistent with such an idea, NOS-3 mutant mice were found to have higher blood pressures than control mice. Other investigators have also reported higher blood pressures in mutant mice deficient in NOS-3 (22). Furthermore, we found increased numbers of glomus cells in mutant mice, compared with controls (Fig. 7). This hyperplasia of glomus cells seen in NOS-3 mutant mice is reminiscent of that reported in the carotid bodies of hypertensive rats (19). Together, the blunted peripheral chemosensitivity in NOS-3 mutant mice might be secondarily due to chronic hypertension. The cellular mechanisms underlying the blunted chemosensitivity, such as alterations in ion channels and transmitters, however, remain to be investigated.

The respiratory control system has been shown to possess dramatic plasticity. It has been suggested that postnatal development of the peripheral chemoreceptors is important for the maturation of adult respiratory behavior. For example, brief hypoxia during the neonatal period affects adult ventilatory control, altering resting breathing patterns and causing attenuation of the hypoxic ventilatory response (38). This effect has been suggested to be due to reduced peripheral chemosensitivity (30, 47). Therefore, the blunting of the hypoxic ventilatory response in NOS-3 mutant mice may be due to altered postnatal development of the peripheral chemoreceptors. Consistent with this idea are the data showing that NOS-3 mutant mice exhibit decreased peripheral chemosensitivity as well as glomus cell hyperplasia, a condition that is also seen in chronic hypoxic animals (13, 34, 50).

It is possible that endogenous NO may also modulate chemosensitivity via its actions on the petrosal ganglion, where the somata of the sensory neurons of the carotid sinus nerve are located (1). However, the petrosal neurons contain NOS-1 but not NOS-3 (56). Therefore, the blunting of the hypoxic ventilatory response may not be secondary to the inhibitory actions of NO from NOS-3 on the sensory afferent fibers.

Possible Involvement of Central Mechanisms in the Blunted Respiratory Responses to Hypoxia in NOS-3 Mutant Mice

In addition to the carotid body, NO can also modulate breathing during hypoxia at the brain stem neurons. Ogawa et al. (37) reported that L-citrulline levels (an index of NO generation) increase during hypoxia in the NTS. In the present study, we monitored brain stem cGMP levels as an index of NO generation during hypoxia and found that cGMP levels were higher in NOS-3 mutant than in WT control mice. These observations are consistent with the idea that hypoxia increases NO generation in the brain stem. Furthermore, they also indicate that NOS-3 does not contribute to the increased NO because this isoform is absent in the mutant mice used in the present study (Fig. 7B). Most likely, increased NO is from NOS-1. This notion is supported by our previous study, in which we found that cGMP levels were unaltered in NOS-1 mutant mice during hypoxia (28). Although NO levels are increased during low oxygen, the central effects of NO on breathing are uncertain. For example, systemic administration of NOS inhibitors decreases the hypoxic ventilatory response, suggesting that NO plays an excitatory role in the central component of respiratory behavior (17). However, administration of NOS inhibitors via the forth ventricle in conscious dogs resulted in an increase in respiration, suggesting that NO may play an inhibitory role in the control of breathing (40). Ogawa et al. found that NO donors microinjected into the NTS regions augment ventilatory response to hypoxia. On the other hand, Vitaglianno et al. (55) found inhibition of breathing in response to microinjection of NO donors into the NTS. Within the pons, NO has been suggested to modulate inspiratory termination (32) as well as to depress RR (31). Although in vitro experiments have demonstrated selectivity of pharmacological NOS inhibitors to individual NOS isoforms, in vivo experiments have indicated that many of these inhibitors cannot distinguish among the different NOS isoforms (59) and that the degree of NOS inhibition may vary within a given physiological system (27). Hence, it is difficult to attribute the actions of NOS inhibitors in the in vivo condition to one or the other NOS isoform. As eluded to above, NOS-3 mutant mice are chronically deficient in the NOS-3 isoform, and the effects of chronic absence of NOS isoforms on breathing may differ from acute inhibition of NOS. Thus, from the present results, we cannot ascertain whether NO from NOS-3 exerts an inhibitory or excitatory effect at the central neurons that regulate breathing during hypoxia.

Because it is produced from the vascular endothelium, NO generated from NOS-3 may also influence breathing via regulating cerebral blood flow (CBF). Inhibitors of NOS produce vasoconstriction of cerebral blood vessels and reduce CBF during normoxia (14). Furthermore, it has been suggested that NO may modulate the hypoxic vasodilation observed during hypoxia (3, 14). Consequently, a reduction in CBF may result in ventilatory depression due to brain hypoxia. However, it has been demonstrated that only severe (50%) CBF reductions result in a reduction of the hypoxic ventilatory response, whereas moderate reductions (30%) increase the hypoxic ventilatory response (8). In the present study, CBF was not examined in the mutant mice; therefore, it is not known to what extent the CBF is altered in mutant mice during hypoxia and to what extent this affects the hypoxic ventilatory response.

The blunting of the chemoreceptor sensitivity could also be due to central interaction of baroreceptor and chemoreceptor inputs in the NTS (35). Heistad et al. (21) has demonstrated that an elevation of systemic arterial pressure depresses respiratory augmentation resulting from carotid body stimulation. Likewise, Attinger et al. (2) reported that increasing the carotid sinus pressure reduces the ventilatory response to low PO2. Given that the NOS-3 mutant mice have higher blood pressure, their baroreceptor discharge is expected to be higher than control mice. Therefore, the decrease in the ventilatory response to hypoxia in NOS-3 mice may, in part, be also due to central interaction of the baroreceptor and chemoreflex pathways.

Physiological Significance of a NO Generated by NOS-3

Recent studies have suggested that the biological effects of NO depend on the source of its production. For example, during focal cerebral ischemia, NO generated from NOS-1 has been shown to exert a toxic effect on neurons, whereas NO produced by NOS-3 confers toxic resistance to neurons (23, 24). The fact that NOS-1 mutant mice have augmented (28), whereas the NOS-3 mutant mice exhibit blunted, respiratory responses to hypoxia supports the idea that the modulatory effect of NO on breathing depends on the source of its production. For instance, both NOS-1 and NOS-3 are located within the carotid body, the sensory organ responsible for the ventilatory response to hypoxia. However, these isoforms may modulate carotid body activity by different mechanisms. On the one hand, NO generated by NOS-3 may modulate peripheral chemosensitivity through regulation of vascular tone and thus the local PO2 in the carotid body (41). On the other hand, NO produced from NOS-1 may inhibit carotid body activity primarily through its action on the chemoreceptor glomus cells and nerve fibers (41, 48). In summary, the results of the present study demonstrate that the respiratory responses to hypoxia are selectively blunted in NOS-3-deficient mice. The attenuated hypoxic responses are due, in part, to blunted carotid body chemosensitivity. These observations are consistent with the idea that NO derived from NOS-3 plays an integral role in the respiratory responses to hypoxia but not during hypercapnia.


    ACKNOWLEDGEMENTS

We are grateful to Dr. Paul L. Huang of Harvard Medical School for supplying the NOS-3 mutant mice and to Dr. Ronald Walenga and the Cystic Fibrosis Core Center (National Institute of Diabetes and Digestive and Kidney Diseases Grant P30-DK-27651) for help with cGMP analysis. Our sincere thanks to Dr. Jeffery Overholt for valuable suggestions.


    FOOTNOTES

This work is supported by National Heart, Lung, and Blood Institute Grant HL-25830 (N. R. Prabhakar); D. D. Kline is supported by training grant T32-HL-07887.

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.

Original submission in response to a special call for papers on "Hypoxia Influence on Gene Expression."

Address for reprint requests and other correspondence: N. R. Prabhakar, Dept. of Physiology and Biophysics, School of Medicine, 10900 Euclid Ave., Case Western Reserve Univ., Cleveland, OH 44106 (E-mail: nrp{at}po.cwru.edu).

Received 16 August 1999; accepted in final form 29 November 1999.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Alcayaga, J, Barrios M, Bustos F, Miranda G, Molina MJ, and Iturriaga R. Modulatory effect of nitric oxide on acetylcholine-induced activation of cat petrosal ganglion neurons in vitro. Brain Res 825: 194-198, 1999[ISI][Medline].

2.   Attinger, FM, Attinger EO, Cooperson D, and Gottschalk W. Interactions between carotid sinus mechanoreceptor and chemoreceptor reflex loops. Pflügers Arch 363: 255-261, 1976[ISI][Medline].

3.   Audibert, G, Saunier CG, Siat J, Hartemann D, and Lambert J. Effect of the inhibitor of nitric oxide synthase, NG-nitro-L-arginine methyl ester, on cerebral and myocardial blood flows during hypoxia in the awake dog. Anesth Analg 81: 945-951, 1995[Abstract].

4.   Barros, RC, and Branco LG. Effect of nitric oxide synthase inhibition on hypercapnia-induced hypothermia and hyperventilation. J Appl Physiol 85: 967-972, 1998[Abstract/Free Full Text].

5.   Bartlett, D, and Tenney SM. Control of breathing in experimental anemia. Respir Physiol 10: 384-395, 1970[ISI][Medline].

6.   Brown, GC. Nitric oxide and mitochondrial respiration. Biochim Biophys Acta 1411: 351-369, 1999[Medline].

7.   Buelke-Sam, J, Holson JF, Bazare JJ, and Young JF. Comparative stability of physiological parameters during sustained anesthesia in rats. Lab Anim Sci 28: 157-162, 1978[ISI][Medline].

8.   Chapman, RW, Santiago TV, and Edelman NH. Effects of graded reduction of brain blood flow on chemical control of breathing. J Appl Physiol 47: 1289-1294, 1979[Abstract/Free Full Text].

9.   Chugh, DK, Katayama M, Mokashi A, Bebout DE, Ray DK, and Lahiri S. Nitric oxide-related inhibition of carotid chemosensory nerve activity in the cat. Respir Physiol 97: 147-156, 1994[ISI][Medline].

10.   Cooper, CE. Nitric oxide and iron proteins. Biochim Biophys Acta 1411: 290-309, 1999[Medline].

11.   Crystal, GJ, Zhou X, Halim AA, Alam S, El Orbany M, and Salem MR. Nitric oxide does not modulate whole body oxygen consumption in anesthetized dogs. J Appl Physiol 86: 1944-1949, 1999[Abstract/Free Full Text].

12.   Dejours, P. Chemoreceptors in breathing. Physiol Rev 42: 335-358, 1962[Free Full Text].

13.   Edwards, C, Heath D, Harris P, Castillo Y, Kruger H, and Arias-Stella J. The carotid body in animals at high altitude. J Pathol 104: 231-238, 1971[ISI][Medline].

14.   Faraci, FM, and Heistad DD. Regulation of the cerebral circulation: role of endothelium and potassium channels. Physiol Rev 78: 53-97, 1998[Abstract/Free Full Text].

15.   Gautier, H. Interactions among metabolic rate, hypoxia, and control of breathing. J Appl Physiol 81: 521-527, 1996[Abstract/Free Full Text].

16.   Gozal, D, Gozal E, Gozal YM, and Torres JE. Nitric oxide synthase isoforms and peripheral chemoreceptor stimulation in conscious rats. Neuroreport 7: 1145-1148, 1996[ISI][Medline].

17.   Gozal, D, Torres JE, Gozal YM, and Littwin SM. Effect of nitric oxide synthase inhibition on cardiorespiratory responses in the conscious rat. J Appl Physiol 81: 2068-2077, 1996[Abstract/Free Full Text].

18.   Grimes, PA, Mokashi A, Stone RA, and Lahiri S. Nitric oxide synthase in autonomic innervation of the cat carotid body. J Auton Nerv Syst 54: 80-86, 1995[ISI][Medline].

19.   Habeck, JO, Honig A, Pfeiffer C, and Schmidt M. The carotid bodies in spontaneously hypertensive (SHR) and normotensive rats---a study concerning size, location and blood supply. Anat Anz 150: 374-384, 1981[ISI][Medline].

20.   Haxhiu, MA, Chang CH, Dreshaj IA, Erokwu B, Prabhakar NR, and Cherniack NS. Nitric oxide and ventilatory response to hypoxia. Respir Physiol 101: 257-266, 1995[ISI][Medline].

21.   Heistad, D, Abboud FM, Mark AL, and Schmid PG. Effect of baroreceptor activity on ventilatory response to chemoreceptor stimulation. J Appl Physiol 39: 411-416, 1975[Abstract/Free Full Text].

22.   Huang, PL, Huang Z, Mashimo H, Bloch KD, Moskowitz MA, Bevan JA, and Fishman MC. Hypertension in mice lacking the gene for endothelial nitric oxide synthase. Nature 377: 239-242, 1995[Medline].

23.   Huang, Z, Huang PL, Ma J, Meng W, Ayata C, Fishman MC, and Moskowitz MA. Enlarged infarcts in endothelial nitric oxide synthase knockout mice are attenuated by nitro-L-arginine. J Cereb Blood Flow Metab 16: 981-987, 1996[ISI][Medline].

24.   Huang, Z, Huang PL, Panahian N, Dalkara T, Fishman MC, and Moskowitz MA. Effects of cerebral ischemia in mice deficient in neuronal nitric oxide synthase. Science 265: 1883-1885, 1994[Abstract/Free Full Text].

25.   Jacoby, RO, and Fox JG. Biology and diseases of mice. In: Laboratory Animal Medicine, edited by Fox JG, Cohen BJ, and Loew FM.. New York: Academic, 1984, p. 31-89.

26.   Jaffrey, SR, and Snyder SH. Nitric oxide: a neural messenger. Ann Rev Cell Dev Biol 11: 417-440, 1995[ISI][Medline].

27.   Kalisch, BE, Connop BP, Jhamandas K, Beninger RJ, and Boegman RJ. Differential action of 7-nitro indazole on rat brain nitric oxide synthase. Neurosci Lett 219: 75-78, 1996[ISI][Medline].

28.   Kline, DD, Yang T, Huang PL, and Prabhakar NR. Altered respiratory responses to hypoxia in mutant mice deficient in neuronal nitric oxide synthase. J Physiol (Lond) 511: 273-287, 1998[Abstract/Free Full Text].

29.   Kusakabe, T, Matsuda H, Harada Y, Hayashida Y, Gono Y, Kawakami T, and Takenaka T. Changes in the distribution of nitric oxide synthase immunoreactive nerve fibers in the chronically hypoxic rat carotid body. Brain Res 795: 292-296, 1998[ISI][Medline].

30.   Landauer, RC, Pepper DR, and Kumar P. Effect of chronic hypoxaemia from birth upon chemosensitivity in the adult rat carotid body in vitro. J Physiol (Lond) 485: 543-550, 1995[ISI][Medline].

31.   Leonard, TO, and Lydic R. Pontine nitric oxide modulates acetylcholine release, rapid eye movement sleep generation, and respiratory rate. J Neurosci 17: 774-785, 1997[Abstract/Free Full Text].

32.   Ling, L, Karius DR, Fiscus RR, and Speck DF. Endogenous nitric oxide required for an integrative respiratory function in the cat brain. J Neurophysiol 68: 1910-1912, 1992[Abstract/Free Full Text].

33.   Lo, EH, Hara H, Rogowska J, Trocha M, Pierce AR, Huang PL, Fishman MC, Wolf GL, and Moskowitz MA. Temporal correlation mapping analysis of the hemodynamic penumbra in mutant mice deficient in endothelial nitric oxide synthase gene expression. Stroke 27: 1381-1385, 1996[Abstract/Free Full Text].

34.   McGregor, KH, Gil J, and Lahiri S. A morphometric study of the carotid body in chronically hypoxic rats. J Appl Physiol 57: 1430-1438, 1984[Abstract/Free Full Text].

35.   Mifflin, SW. Inhibition of chemoreceptor inputs to nucleus of tractus solitarius neurons during baroreceptor stimulation. Am J Physiol Regulatory Integrative Comp Physiol 265: R14-R20, 1993[Abstract/Free Full Text].

36.   Moncada, S, Palmer RM, and Higgs EA. Nitric oxide: physiology, pathophysiology, and pharmacology. Pharmacol Rev 43: 109-141, 1991[ISI][Medline].

37.   Ogawa, H, Mizusawa A, Kikuchi Y, Hida W, Miki H, and Shirato K. Nitric oxide as a retrograde messenger in the nucleus tractus solitarii of rats during hypoxia. J Physiol (Lond) 486: 495-504, 1995[ISI][Medline].

38.   Okubo, S, and Mortola JP. Control of ventilation in adult rats hypoxic in the neonatal period. Am J Physiol Regulatory Integrative Comp Physiol 259: R836-R841, 1990[Abstract/Free Full Text].

39.   Patel, GM, Horstman DJ, Adams JM, and Rich GF. Nitric oxide synthase inhibitors alter ventilation in isoflurane anesthetized rats. Anesthesiology 88: 1240-1248, 1998[ISI][Medline].

40.   Pelligrino, DA, Laurito CE, and VadeBoncouer TR. Nitric oxide synthase inhibition modulates the ventilatory depressant and antinociceptive actions of fourth ventricular infusions of morphine in the awake dog. Anesthesiology 85: 1367-1377, 1996[ISI][Medline].

41.   Prabhakar, NR. NO and CO as second messengers in oxygen sensing on the carotid body. Respir Physiol 115: 161-168, 1999[ISI][Medline].

42.   Prabhakar, NR, Kumar GK, Chang CH, Agani FH, and Haxhiu MA. Nitric oxide in the sensory function of the carotid body. Brain Res 625: 16-22, 1993[ISI][Medline].

43.   Przybylski, J, Tafil-Klawe M, Trzebski A, and Klawe J. Increased resistance to acute anoxia of the respiratory and cardiac function in spontaneously hypertensive rats. Acta Physiol Pol 36: 42-50, 1985[Medline].

44.   Przybylski, J, Trzebski A, Czyzewski T, and Jodkowski J. Responses to hyperoxia, hypoxia, hypercapnia and almitrine in spontaneously hypertensive rats. Bull Eur Physiopath Respir 4: 145-154, 1982.

45.   Przybylski, J, Trzebski A, and Przybyszewski A. Circulatory responses to acute hypoxia in spontaneously hypertensive and normotensive rats. Acta Physiol Pol 31: 463-468, 1980[Medline].

46.   Schlenker, EH, and Farkas GA. Endogenous opoids modulate ventilation in the obese Zucker rat. Respir Physiol 99: 97-103, 1995[ISI][Medline].

47.   Sterni, LM, Bamford OS, Wasicko MJ, and Carroll JL. Chronic hypoxia abolished the postnatal increase in carotid body type I cell sensitivity to hypoxia. Am J Physiol Lung Cell Mol Physiol 277: L645-L652, 1999[Abstract/Free Full Text].

48.   Summers, BA, Overholt JL, and Prabhakar NR. Nitric oxide inhibits L-type Ca2+ current in glomus cells of the rabbit carotid body via a cGMP-independent mechanism. J Neurophysiol 81: 1449-1457, 1999[Abstract/Free Ful