| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
Department of Comparative Biosciences, School of Veterinary Medicine, University of Wisconsin, Madison, Wisconsin 53706
 |
ABSTRACT |
|---|
 TOP
 ABSTRACT
 INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
|
|---|
Episodic hypoxia elicits a long-lasting augmentation of phrenic inspiratory activity known as long-term facilitation (LTF). We investigated the respective contributions of carotid chemoafferent neuron activation and hypoxia to the expression of LTF in urethane-anesthetized, vagotomized, paralyzed, and ventilated Sprague-Dawley rats. One hour after three 5-min isocapnic hypoxic episodes [arterial PO2 (PaO2) = 40 ± 5 Torr], integrated phrenic burst amplitude was greater than baseline in both carotid-denervated (n = 8) and sham-operated (n = 7) rats (P < 0.05), indicating LTF. LTF was reduced in carotid-denervated rats relative to sham (P < 0.05). In this and previous studies, rats were ventilated with hyperoxic gas mixtures (inspired oxygen fraction = 0.5) under baseline conditions. To determine whether episodic hyperoxia induces LTF, phrenic activity was recorded under normoxic (PaO2 = 90-100 Torr) conditions before and after three 5-min episodes of isocapnic hypoxia (PaO2 = 40 ± 5 Torr; n = 6) or hyperoxia (PaO2 > 470 Torr; n = 6). Phrenic burst amplitude was greater than baseline 1 h after episodic hypoxia (P < 0.05), but episodic hyperoxia had no detectable effect. These data suggest that hypoxia per se initiates LTF independently from carotid chemoafferent neuron activation, perhaps through direct central nervous system effects.
plasticity; respiratory control; carotid body; episodic hypoxia
| |
INTRODUCTION |
|---|
|
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
|
|---|
EPISODIC HYPOXIA OR ELECTRICAL stimulation of carotid chemoafferent neurons elicits a long-lasting augmentation of respiratory motor output known as long-term facilitation (LTF; 44, 54). When repeated carotid sinus nerve (CSN) stimulation is used to elicit LTF in cats (40, 41) and rats (23, 33), thus bypassing chemotransduction in the carotid body, central neural mechanisms must be involved. LTF is also observed after episodic hypoxia in anesthetized rats (23; additional references in Ref. 44), sleeping humans (2), and awake dogs (10), goats (60), ducks (43), rats (52), and mice (31); the pattern of hypoxia is critical because episodic hypoxia elicits LTF whereas a single bout of continuous hypoxia does not (5, 6, 14, 52, 60). It has been proposed that hypoxia stimulates carotid body chemoreceptors and that the associated intermittent chemoafferent neuron activation elicits LTF by central neural mechanisms without the need for hypoxia per se (3). Although this model is intuitively appealing because it links LTF induced by intermittent hypoxia and electrical CSN stimulation, there is no clear evidence that the same neural mechanisms underlie both forms of LTF.
LTF induced by episodic hypoxia and electrical CSN stimulation both require serotonin receptor activation (3, 7, 18, 28, 39, 41). Moreover, in anesthetized rats, LTF in phrenic nerve motor output after episodic hypoxia requires serotonin receptor activation and protein synthesis in the cervical spinal cord (7), confirming that hypoxia-induced LTF results from central nervous system (CNS) mechanisms (44, 53). On the other hand, there are indications that LTF involves at least two distinct mechanisms, one associated with carotid chemoafferent neuron activation and the other involving direct effects of hypoxia on the CNS. For example, LTF expression in anesthetized rats differs qualitatively after CSN stimulation and episodic hypoxia (44). CSN stimulation elicits decrementing phrenic LTF of relatively short duration (<1 h; 23, 33). In contrast, hypoxia-induced LTF starts small immediately after hypoxia but increases progressively over the next hour (3, 44). These contrasting manifestations of LTF could arise from differences in chemoafferent activation by electrical vs. hypoxic stimulation or from differences in the balance of inhibitory and facilitatory mechanisms that they evoke (6, 29). Alternatively, hypoxia-induced LTF may arise from a fundamentally different mechanism, perhaps through direct effects of hypoxia on CNS neurons. Consistent with this hypothesis, a form of LTF has been reported after brief hypoxic exposures in peripherally chemodenervated, anesthetized cats (20). Thus hypoxia may contribute to LTF by mechanisms that do not involve carotid chemoafferent neuron activation.
There is also some question as to the relative roles of hypoxia and hyperoxia in the initiation of LTF by episodic hypoxia. In previous studies on hypoxia-induced LTF in anesthetized rats (e.g., 19, 23) and cats (20), animals were ventilated with hyperoxic gas mixtures [inspired oxygen fraction (FIO2) = 0.5-1.0] before and after hypoxia, ostensibly to improve preparation stability. Consequently, each hypoxic episode ended with a rapid oxygen increase to hyperoxic levels. The repeated oxygen rise after the hypoxic bouts may alter cell signaling and gene expression, perhaps through the formation of reactive oxygen species (11, 13, 26, 56). Thus it is unclear whether hypoxia per se is the stimulus for LTF or whether the repeated rise in oxygen (to hyperoxic levels) elicits this form of respiratory plasticity.
In this study, we investigated the role of hypoxia in phrenic LTF in anesthetized rats. In one set of experiments, we tested the hypothesis that episodic hypoxia, and not episodic hyperoxia, is a proximate stimulus for LTF. Specifically, we compared the effects of episodic hypoxia vs. hyperoxia in otherwise normoxic rats. After confirming that episodic hypoxia was the relevant stimulus, we studied carotid-denervated (CSNX) rats to test the hypothesis that episodic hypoxia elicits LTF independently from carotid chemoafferent neuron activation.
| |
METHODS |
|---|
|
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
|
|---|
Experiments were conducted on 3- to 5-mo-old male Sprague-Dawley rats (311-414 g; Charles River Laboratories, Wilmington, MA; rat colony K-62, Kingston, NY). The general experimental preparation and protocol have been described previously (3, 19). All experimental procedures were approved by the Animal Care and Use Committee of the School of Veterinary Medicine at the University of Wisconsin-Madison.
Experimental Preparation
Anesthesia was induced with isoflurane and maintained (2.5% isoflurane, FIO2 = 0.2 or 0.5, balance N2) first through a nose cone and then via a tracheal cannula placed to allow pump ventilation (Rodent Respirator, model 683, Harvard Apparatus, Holliston, MA). A catheter was placed in the femoral vein, and the rats were slowly converted to urethane anesthesia (1.6 g/kg iv). Adequacy of anesthesia was monitored throughout the experiment by testing blood pressure responses to toe pinch; supplemental urethane was given if a response was observed. A catheter was placed in the femoral artery to monitor blood pressure (P23ID pressure transducer and P122 amplifier, Gould) and to withdraw 0.2-ml blood samples to determine blood gases and pH with a blood analysis system (ABL-500, Radiometer, Copenhagen, Denmark). Arterial blood values were corrected to rectal temperature, which was maintained at 37-38°C with a heated table. Animals received a continuous intravenous infusion (2.5 ml/h) of a 1:11 (vol/vol) solution of sodium bicarbonate (5%) and lactated Ringer solution to maintain fluid and acid-base balance, beginning shortly after urethane administration. Animals were bilaterally vagotomized in the midcervical region and paralyzed with pancuronium bromide (3.25-3.5 mg/kg) to prevent spontaneous breathing movements and entrainment of neurograms with the ventilator. End-tidal carbon dioxide partial pressures (PETCO2) were measured with a rapidly responding flow-through carbon dioxide analyzer (Capnogard, Novametrix, Wallingford, CT) placed in the expired line of the ventilator circuit.The left phrenic nerve was isolated by use of a dorsal approach, cut distally, desheathed, submerged in mineral oil, and placed on a bipolar silver wire electrode. Nerve activity was amplified (×10,000), band-pass filtered (100 Hz to 10 kHz; Model 1700 or 1800, A-M Systems, Carlsborg, WA), and integrated (time constant = 50 ms; Moving Averager, model MA-821RSP, CWE, Ardmore, PA). The integrated signal was digitized and processed with commercially available computer software (WinDaq 2.18, DATAQ Instruments, Akron, OH).
Carotid Body Denervation
In some preparations (series 2; see below), the carotid sinus nerve was isolated and sectioned bilaterally immediately before isolation of the phrenic nerve. After arterial and venous catheters were implanted, the carotid bifurcation was exposed bilaterally via a dorsal-lateral approach. The carotid sinus nerve was then carefully isolated at its junction with the glossopharyngeal nerve. In CSNX rats, the CSN nerve was sectioned and the entire region was covered with a mineral oil-petroleum jelly mixture to prevent desiccation. Sham-operated rats were treated identically, but the isolated carotid sinus nerves were left intact.Completeness of carotid chemodenervation was assessed in a separate group of rats. Whereas intravenous injection of sodium cyanide (25 µg NaCN in 0.2 ml saline) produced a robust increase in phrenic nerve activity in rats with intact carotid bodies (n = 3), this response was completely abolished in rats in which the carotid sinus nerve was sectioned bilaterally (n = 3).
Experimental Protocol
Preparations were allowed to stabilize for ~60 min after surgery (FIO2 = 0.2 or 0.5 in series 1 and 2, respectively; PETCO2 ~40 Torr). The carbon dioxide apneic threshold was then determined by hyperventilating the rat until phrenic nerve activity ceased and then slowly raising PETCO2 by decreasing ventilator rate or increasing inspired carbon dioxide until rhythmic activity reappeared. Baseline neural activity was standardized among preparations by maintaining PETCO2 3 Torr above the PETCO2 at which activity resumed. After baseline phrenic activity was established (30 min), an arterial blood sample (0.2-0.3 ml in a heparinized syringe) was drawn; subsequent blood samples were compared with this baseline value.Series 1: episodic hypoxia vs. episodic hyperoxia, normoxic background. Baseline phrenic nerve activity was established under normoxic and normocapnic conditions [FIO2 = 0.2, arterial PO2 (PaO2) 90-100 Torr; PETCO2 3 Torr above apneic threshold]. Rats were then exposed to 1) three 5-min bouts of hypoxia (episodic hypoxia; PaO2 = 40 ± 5 Torr; n = 6), 2) three 5-min bouts of hyperoxia (episodic hyperoxia; FIO2 ~1.0, PaO2 > 470 Torr; n = 6), or 3) maintained baseline conditions (time control; n = 6); PaCO2 was maintained within 2 Torr of baseline during hypoxic and hyperoxic episodes. Blood samples were collected during the final 30 s of the first treatment bout, and, if blood gases were not within the acceptable range, FIO2 and/or inspired carbon dioxide fraction were adjusted during the next bout. Rats were returned to normoxia for 5 min between bouts of hypoxia or hyperoxia. After the last bout of hypoxia or hyperoxia, or the equivalent period of normoxia, phrenic nerve activity was monitored for 1 h under baseline conditions. Blood samples were collected at 15, 30, and 60 min to ensure that blood gases remained normoxic and isocapnic (PaCO2 within 1 Torr of baseline). At the conclusion of experiments, rats were killed via urethane overdose.
Series 2: episodic hypoxia in CSNX rats. Baseline phrenic nerve activity was established under hyperoxic and normocapnic conditions (FIO2 = 0.5, PaO2 > 190 Torr; PETCO2 3 Torr above apneic threshold). CSNX or sham-operated rats were then exposed to three 5-min bouts of hypoxia (episodic hypoxia; PaO2 = 40 ± 5 Torr; n = 8 CSNX, 7 sham) or maintained under baseline conditions (time control; n = 8 CSNX, 8 sham); PaCO2 was maintained within 2 Torr of baseline during hypoxic episodes. Blood samples were collected during the final 30 s of the first hypoxic bout, and, if blood gases were not in the acceptable range, FIO2 and/or inspired carbon dioxide fraction were adjusted during the next bout. Rats were returned to hyperoxia for 5 min between bouts of hypoxia. After the last bout of hypoxia or the equivalent period of hyperoxia, phrenic nerve activity was monitored for 1 h under baseline conditions. Blood samples were collected at 15, 30, and 60 min to ensure that blood gases remained isocapnic (PaCO2 within 1.5 Torr of baseline). At the conclusion of experiments, rats were killed via urethane overdose.
Data Analysis
Phrenic activity was averaged in 30-s bins (immediately preceding blood sampling) under baseline conditions; during the fifth minute of hypoxia, hyperoxia, or normoxia; and 15, 30, and 60 min after episodic treatment. Variables measured included peak amplitude of integrated phrenic activity, phrenic burst frequency, and their product, the minute phrenic activity. Changes from baseline in burst amplitude and minute activity were normalized as a percentage of baseline values (% baseline).Apneic thresholds were compared among treatment groups by use of
one-way ANOVA. Acute hypoxic and hyperoxic responses and time-dependent
changes in phrenic activity, blood gases, and blood pressure were
compared among treatment groups by use of two-way repeated-measures
ANOVA followed by Student-Newman-Keuls post hoc tests. In the
series 2 experiments, acute hypoxic responses were
reanalyzed by using one-sample t-tests to improve
statistical power. Linear regression was used to investigate the
potential influence of arterial blood pressure on the expression of
LTF. Effects were considered statistically significant at
P 
0.05. Unless otherwise noted, statistical tests
were conducted by using SigmaStat 2.03, SPSS, Chicago, IL). All data
are presented as means ±SE.
| |
RESULTS |
|---|
|
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
|
|---|
Series 1: Episodic Hypoxia Vs. Episodic Hyperoxia
Blood gases and mean arterial blood pressure.
Baseline PaO2, PaCO2, and mean
arterial pressure (MAP) were similar among the three treatment groups
used in these experiments, and rats remained isocapnic
(PaCO2 within 1 Torr of baseline) throughout the
protocol (Table 1). Acute hypoxic
challenges (mean PaO2 = 40 Torr) decreased MAP
(
= 
49 ± 5 mmHg; P < 0.001), but MAP
returned to baseline in normoxia and did not change significantly throughout the remainder of the protocol in the episodic hypoxia rats.
Increasing FIO2 to nearly 1.0 (mean
PaO2 > 495 Torr) caused a small but
statistically significant increase in MAP ( = 16 ± 3 mmHg; P = 0.011) that disappeared on return to
normoxia. PaO2, PaCO2 and MAP were
stable throughout the protocol in rats maintained under normoxic
conditions.
|
Responses to acute hypoxia and hyperoxia.
Acute hypoxia elicited a brisk increase in phrenic activity. Peak burst
amplitude increased significantly (87 ± 11% baseline; P < 0.001) with no change in burst frequency ( = 1 ± 3 bursts/min; P = 0.769) during the fifth
minute of hypoxia; consequently, minute phrenic activity also increased
significantly during hypoxia (93 ± 22% baseline;
P = 0.008). Acute hyperoxia caused no change in burst
amplitude (8 ± 5% baseline; P = 0.154) or minute
phrenic activity (0 ± 7% baseline; P = 0.980),
although there was a small decrease in burst frequency ( = 4 ± 1 bursts/min; P = 0.048).
LTF of phrenic activity.
Episodic hypoxia elicited significant LTF in the peak amplitude of
integrated phrenic bursts and in phrenic minute activity but not in
burst frequency (Fig. 1). LTF was evident
as an increase in phrenic amplitude by 30 min posthypoxia (30 ± 9% increase from baseline; P = 0.001) that remained
elevated 60 min posthypoxia (44 ± 10%; P < 0.001). Burst frequency was somewhat reduced 15 min posthypoxia
( = 4 ± 2 bursts/min; P = 0.008) but
returned to baseline by 30 min posthypoxia. As a result, minute phrenic activity was elevated above baseline at the 30 (32 ± 12%;
P = 0.005)- and 60 (47 ± 11%; P < 0.001)-min time points. The progressive facilitation of phrenic
amplitude and minute activity between 15 and 30 min posthypoxia was
statistically significant (P = 0.006 and 0.002, respectively). Although phrenic activity continued increasing between
30 and 60 min posthypoxia, the changes in amplitude and minute activity
were not statistically significant during this time (P = 0.088 and 0.117, respectively). There was no relationship between the
magnitude of LTF and the blood pressure (MAP and/or MAP) at
baseline, during hypoxia, or 60 min posthypoxia (all P > 0.05).
|
Series 2: Episodic Hypoxia in CSNX rats
Apneic threshold, blood gases, and MAP.
Acute CSN section produced a marginally significant increase in carbon
dioxide apneic threshold [PETCO2 = 42 ± 1 vs. 40 ± 1 Torr in CSNX (n = 16) and
sham-operated (n = 15) rats, respectively; P = 0.051], but baseline PaCO2 did
not differ between groups (45.1 ± 0.7 vs. 44.2 ± 0.7;
P = 0.345). Consequently, baseline
PaCO2, as well as baseline PaO2 and
MAP, were similar among the four treatment groups (Table
2). Rats remained isocapnic (average PaCO2 within 1 Torr of baseline) throughout the
protocol (Table 2).
|
MAP = 72 ± 6 vs. 32 ± 6 mmHg in
sham-operated rats; P < 0.001); MAP returned to
baseline after episodic hypoxia in both groups. Although MAP tended to
fall (5-15 mmHg) over the course of the protocol in all four
treatment groups, this drop reached statistical significance only in
CSNX rats 60 min after episodic hypoxia ( MAP = 14 ± 5;
P = 0.003 vs. baseline). PaO2, PaCO2, and MAP were stable in CSNX and sham-operated
rats maintained under hyperoxic conditions throughout the protocol.
Phrenic responses to acute hypoxia in CSNX and sham-operated rats. Acute CSN section had no effect on peak amplitude of integrated phrenic bursts or phrenic burst frequency under baseline, hyperoxic conditions [for CSNX (n = 16) vs. sham-operated (n = 15) rats: amplitude 2.4 ± 0.3 vs. 2.4 ± 0.2 volts, P = 0.651; frequency 47 ± 1 vs. 46 ± 2 bursts/min, P = 0.624].
Acute hypoxia (PaO2 ~40 Torr) elicited a brisk increase in minute phrenic activity in sham-operated rats, but this response was greatly reduced in CSNX rats (Figs. 2 and 3; P < 0.001). In the fifth minute of hypoxia, sham-operated rats exhibited significantly elevated minute phrenic activity (128 ± 24% increase from baseline; P < 0.001), resulting from an increase in burst amplitude (111 ± 24% increase from baseline; P < 0.001) without change in burst frequency ( = 3 ± 2 bursts/min; P = 0.353). Despite apparent increases in phrenic amplitude and minute activity during acute hypoxia, changes in phrenic amplitude, burst frequency, and minute activity were not statistically significant for CSNX rats when analyzed
with sham-operated rats by two-way repeated-measures ANOVA ( = 15 ± 4% baseline, 1 ± 1 bursts/min, and 18 ± 7%
baseline, respectively; all P > 0.05). This lack of
significance is likely the result of unequal variance between groups;
indeed, when hypoxic phrenic responses are considered for CSNX rats
alone (one sample t-test; SPSS 10.1, SPSS), the small
increases in phrenic amplitude and minute activity reach statistical
significance (both P < 0.05). Thus CSNX rats typically
experienced some residual increase in phrenic activity during hypoxia,
although the increase was greatly reduced and considerably delayed
(20-40 s) relative to sham-operated rats (Fig. 2).
|
|
LTF of phrenic activity.
Phrenic activity did not change over the course of the experiment in
either sham-operated or CSNX rats that were not exposed to episodic
hypoxia (i.e., time controls; Fig. 4; all
P > 0.05 vs. baseline).
|
4 ± 2 bursts/min; P = 0.003 vs. baseline) but returned to
baseline for the remainder of the experiment (P > 0.05). Consequently, minute phrenic activity was significantly greater
than baseline at 30 (28 ± 14%; P = 0.003) and 60 min (60 ± 8%; P < 0.001) posthypoxia. Increases
in phrenic amplitude and minute activity were significantly greater at
60 min posthypoxia than at 30 min and greater at 30 min than at 15 min
(all P < 0.05). Moreover, the time-dependent changes
in phrenic activity discussed above (vs. baseline) were significant
compared with sham-operated rats that were not exposed to episodic
hypoxia (Fig. 4; all P < 0.05). An eighth
sham-operated rat was originally studied with this group; however, this
rat responded to episodic hypoxia with a long-lasting depression of
phrenic activity (34 and 38% decrease from baseline in phrenic
amplitude and minute activity, respectively). Using the same protocol,
our laboratory has never observed a long-lasting respiratory depression
(decrease from baseline > 15%) in comparable rats after episodic
hypoxia (n > 100 healthy, male rats); for example, in
a recent meta-analysis from our laboratory (19), only 2 of
63 rats exhibited phrenic amplitudes less than baseline at 60 min (1
and 13%). Consequently, this anomalous rat was excluded from the
analyses. Although including this rat lowers the mean value for LTF at
60 min posthypoxia (to 44 ± 14 and 47 ± 14% increase from
baseline in phrenic amplitude and minute activity, respectively), LTF
in both phrenic amplitude and minute activity would remain
statistically significant in sham-operated rats (both P < 0.001).
Episodic hypoxia also elicited LTF in CSNX rats, although the
expression of LTF differed somewhat from sham-operated rats (Fig. 4).
As in sham-operated rats, the magnitude of integrated phrenic burst
amplitude increased after episodic hypoxia in CSNX rats and was
significantly greater than baseline at 30 (26 ± 6%; P = 0.002) and 60 min (27 ± 7%, range
1-50%; P = 0.002) posthypoxia; these increases
in phrenic amplitude were also significant compared with CSNX rats that
were not exposed to episodic hypoxia (P = 0.002 and
P < 0.001 at 30 and 60 min, respectively). However, rather than having reduced burst frequencies 15 min posthypoxia, CSNX
rats exhibited a small (3 ± 1 bursts/min), but statistically significant, increase in burst frequency relative to baseline at all
posthypoxia time points (all P < 0.05). Compared with
CSNX rats that were not exposed to episodic hypoxia, this frequency LTF
was significant at 30 min posthypoxia (P = 0.018) and
marginally significant at 60 min (P = 0.054). As a
result of increases in phrenic amplitude and burst frequency, minute
phrenic activity was significantly greater than baseline at 15 (20 ± 10%; P = 0.021), 30 (37 ± 8%;
P < 0.001), and 60 min (38 ± 9%;
P < 0.001) posthypoxia. Likewise, minute phrenic
activity was significantly greater at 30 and 60 min compared with CSNX
rats that were not exposed to episodic hypoxia (both P < 0.001). Overall, phrenic activity tended to increase between 15 and
30 min after episodic hypoxia in CSNX rats due to a gradual increase in
phrenic amplitude, although this change did not reach statistical
significance. There were no changes in phrenic amplitude, burst
frequency, or minute activity between 30 and 60 min posthypoxia (all
P > 0.05). Although phrenic amplitude and minute
activity were similar between sham-operated and CSNX rats at 15 and 30 min posthypoxia (P > 0.05), the leveling off of
phrenic amplitude between 30 and 60 min in CSNX rats (vs. progressive
increase in phrenic amplitude in sham-operated rats) resulted in
significantly less LTF of phrenic amplitude (P = 0.002) and minute activity (P = 0.043) at 60 min posthypoxia
compared with sham-operated rats.
There was no statistically significant relationship between the
magnitude of LTF and the blood pressure (MAP and/or MAP) at
baseline, during hypoxia, or 60 min posthypoxia in CSNX or sham-operated rats (all P > 0.05). However, there was
a nonsignificant trend toward diminished LTF with lower blood pressures
during hypoxia within the CSNX group (r2 = 0.46, P = 0.064).
| |
DISCUSSION |
|---|
|
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
|
|---|
This study demonstrates that LTF is initiated by episodic hypoxia in otherwise normoxic rats but not by episodic hyperoxia. Moreover, the LTF observed in CSNX rats indicates that carotid chemoafferent neuron activation is not essential for hypoxia-induced LTF in anesthetized rats. However, the magnitude of LTF was somewhat reduced in CSNX rats, providing suggestive evidence that chemoafferent neuron activation and hypoxia per se both contribute to the expression LTF. Together, these results suggest a specific role for CNS hypoxia in hypoxia-induced LTF.
Hyperoxia and LTF
Time-dependent effects of episodic hypoxia on respiratory motor output have been studied in anesthetized rats and cats with hyperoxic baseline conditions (e.g., 19, 20, 23); in these studies, the animals were ventilated with hyperoxic gas mixtures (FIO2 > 0.5) to prolong viability of the experimental preparation. Even in the absence of overt hyperoxic toxicity, hyperoxia has the potential to influence cellular processes through the generation of reactive oxygen species (ROS; Refs. 11, 13, 26, 56). Previous studies have shown that hyperoxia can induce plasticity in the control of breathing in humans and other mammals (e.g., 21, 22, 24, 32). However, the results of the present study suggest that hyperoxia does not influence the expression of LTF in anesthetized rats. When superimposed on normoxic arterial blood, episodic hypoxia elicited an LTF of phrenic nerve activity that was qualitatively similar to the LTF reported for hyperoxic rats (19, 44, present study): the amplitude of integrated phrenic bursts returned toward baseline after the third hypoxic episode and then gradually increased over the ensuing 1 h (Fig. 1). The magnitude of LTF in these normoxic rats was somewhat lower than typically reported for anesthetized rats (19), but considerable interstudy variation makes quantitative comparisons among studies difficult. Recent studies in awake rats (52) and mice (31) also revealed LTF after episodic hypoxia in otherwise normoxic, awake animals. Our findings are consistent with hypoxia serving as the proximate stimulus for LTF in anesthetized rats.One consequence of using a background of hyperoxia in episodic hypoxia studies is that, in addition to hypoxic episodes, animals are exposed to rapid rises in oxygen to hyperoxic levels at the end of each hypoxic bout. Brief hyperoxia has been shown to induce plasticity in respiratory control (22, 24). For example, 10-min preexposure to hyperoxia leads to a nitric oxide-dependent augmentation of the hypoxic ventilatory response in rats (22). To our knowledge, the effects of repeated, brief exposures to hyperoxia on resting respiratory motor output have not been investigated. In contrast to episodic hypoxia, three 5-min bouts of hyperoxia failed to elicit LTF of phrenic activity (Fig. 1), thus providing additional evidence that the expression of LTF is unique to hypoxia. Indeed, there was no evidence for any time-dependent changes in phrenic activity after episodic hyperoxia, at least in the time domain studied here, although this does not preclude the possibility that episodic hyperoxia and/or ROS exposures initiate other forms of plasticity (56). We produced hyperoxia by raising FIO2 from normoxia to nearly 100% oxygen (supplemental carbon dioxide was used to maintain isocapnia), and it is not known how the ROS produced by our protocol compare with ROS generated during episodic hypoxia. Therefore, we cannot rule out causative roles of ROS produced during hypoxia or during arterial reoxygenation (i.e., hypoxia to normoxia).
Carotid Denervation and the Short-Term Hypoxic Response
In adult rats, the brisk increase in respiratory motor output during hypoxia is normally mediated through the activation of carotid body chemoreceptors, although other oxygen sensitive tissues contribute to a lesser extent (37). As expected, carotid denervation caused a substantial reduction in the phrenic response to hypoxia (PaO2 = 40 Torr) in the present study. However, residual hypoxic sensitivity remained after bilateral CSN section, as evidenced by a small increase in phrenic amplitude during the fifth minute of hypoxia (Fig. 3; see also Fig. 2). Residual hypoxic responses have been reported previously in awake (37, 51) and anesthetized rats (34) after carotid denervation and most likely reflect extracarotid chemosensitivity or barosensitivity (see below). Although extracarotid peripheral chemoreceptors (e.g., aortic and abdominal chemoreceptors) contribute to hypoxic responses in rats (9, 37), these inputs were eliminated in the present study by bilateral cervical vagotomy. It is also possible that additional oxygen-sensitive chemoreceptors in the cervical region remain intact after CSN transection (37, 38) or that regions of the carotid chemoafferent pathway downstream of the carotid body possess innate hypoxic sensitivity (e.g., neurons of the petrosal ganglion or nucleus tractus solatarius; Refs. 36, 45, 59); however, the existence and functional significance of such residual hypoxic sensitivity are questionable (1, 42, 55). The residual hypoxic response in the present study more likely resides in the CNS.Central hypoxic sensitivity has been implicated in the hypoxic responses in rats (37) and other mammals (e.g., Ref. 12). Many areas of the brain known to modulate respiratory activity have been shown to possess hypoxic sensitivity and could stimulate breathing during central hypoxia (15, 25, 57, 58). Systemic hypoxia has also been shown to produce acidosis in the CNS, including the ventral medulla (50, 63), although this acidosis may not have a stimulatory effect on breathing (49). Alternatively, changes in phrenic activity during hypoxia could be secondary to changes in blood pressure in CSNX rats, either directly through a respiratory baroreflex (61) or through CNS hypoxia, hypercapnia, and acidosis resulting from inadequate perfusion. Consistent with the latter hypothesis, the increase in phrenic amplitude during hypoxia was delayed (20-40 s) in CSNX rats relative to sham-operated rats and consistently followed a drop in arterial blood pressure (see Fig. 2). Moreover, as previously reported for awake rats (8), hypoxic hypotension was more pronounced in CSNX rats than in sham-operated rats. Because brain blood flow regulation is compromised at the low arterial blood pressures achieved in CSNX rats (27), brain PO2 and acid-base homeostasis may be compromised in CSNX rats during hypoxia and ultimately stimulate respiratory activity. The present study was not designed to differentiate between these possibilities (i.e., direct or secondary effects of hypoxia on CNS neurons).
Carotid Denervation and LTF
Episodic hypoxia induced a long-lasting (>1 h) facilitation in phrenic motor output in CSNX rats, demonstrating that the carotid body itself is not necessary for the expression of LTF. This result is consistent with previous evidence indicating that LTF is primarily a CNS mechanism (44). Indeed, Peng and colleagues (53) found no evidence of LTF in carotid chemoafferent activity after episodic hypoxia in the rat, unless the rat was pretreated with 10 days of intermittent hypoxia (8 h/day). Chronic intermittent hypoxia enhances the expression of phrenic LTF in anesthetized rats (35), apparently evoking additional central and peripheral plasticity (35, 44, 53, 56).In carotid body-intact rats (series 1 and 2), phrenic motor output increased progressively throughout the hour after episodic hypoxia, and this facilitation was entirely attributable to changes in integrated phrenic burst amplitude. In contrast, phrenic LTF in CSNX rats involved augmentation of both amplitude and frequency of respiratory bursts. Frequency LTF has been observed in anesthetized (3, 5, 7) and awake (52) rats with functional carotid bodies after episodic hypoxia, although this finding has not been consistent among studies (17, 18, 65, 66, this study). Accordingly, it is difficult to interpret the differential expression of frequency LTF in CSNX and intact rats. A progressive increase in phrenic amplitude for at least 1 h after episodic hypoxia is typical in anesthetized-rat studies (44). Over the first 30 min, this "winding up" pattern was observed in CSNX rats (Fig. 4) as opposed to the decrementing pattern of LTF observed in CSN stimulation experiments (23, 33), suggesting that LTF induced by hypoxia is similar in CSNX and intact rats; additional studies are needed to verify that this LTF has the same underlying mechanisms. However, phrenic amplitude failed to continue increasing between 30 and 60 min in CSNX rats, and CSNX rats ultimately exhibited only one-half to two-thirds as much LTF as sham-operated rats (Fig. 4). It is possible, therefore, that chemoafferent neuron activation is necessary for the full expression of LTF.
Expression of LTF after episodic hypoxia appears to be localized to regions near respiratory motor nuclei (44). For example, phrenic LTF is abolished after intrathecal injection of serotonin receptor antagonists or protein synthesis inhibitors in the cervical spinal cord (7), suggesting that LTF results from plasticity within the phrenic motor nucleus. Recent models of phrenic LTF (6, 19, 29, 44) propose that episodic hypoxia leads to episodic release of serotonin in the region of respiratory motoneurons from projections of the caudal raphe nuclei. Repeated activation of serotonergic 5-HT2 receptors on phrenic motor neurons is expected to initiate an intracellular signaling cascade that ultimately potentiates glutamate-dependent activation of phrenic motor neurons in response to bulbospinal respiratory drive. In this model, as in earlier incarnations (e.g., Ref. 3), it has been hypothesized that hypoxia initiates the release of serotonin at the phrenic motor nucleus through the activation of carotid chemoafferent pathways. Consistent with this hypothesis, both electrical stimulation of the CSN and hypoxia elicit c-fos expression in serotonergic raphe neurons (16); activation of carotid chemoafferent pathways has also been shown to influence raphe discharge in cats (46-48, 64). Assuming that hypoxia-induced LTF has a similar serotonin dependence in CSNX and intact rats, the results of the present study suggest that hypoxia may activate serotonergic cells through multiple, potentially overlapping, pathways. For example, central hypoxia could directly activate raphe cells leading to the release of serotonin near phrenic motor neurons; serotonergic cells from the rat medullary raphe are stimulated by changes in PCO2 and pH in vitro (62), but we are unaware of any studies on the hypoxic sensitivity of these cells. Alternatively, hypoxia could modulate the activity of raphe neurons through direct or indirect interactions of raphe cells with peripheral and/or central hypoxia sensitive tissues remaining after CSN transection and vagotomy (see Carotid Denervation and the Short-Term Hypoxic Response).
Our laboratory recently reported a positive, linear relationship
between the short-term hypoxic phrenic response and posthypoxic LTF in
anesthetized rats (19); rats exhibiting greater short-term hypoxic responses had significantly greater LTF at 60 min posthypoxia. This relationship is not a simple function of increased respiratory drive during hypoxia because increasing respiratory activity with nonhypoxic chemical (e.g., hypercapnia) and nonchemical stimuli does
not initiate LTF (4, 6, 40). One possible explanation for
the correlation is that rats stimulated more by hypoxia may also
experience greater activation of serotonergic raphe neurons and greater
serotonin release near phrenic motoneurons, thereby promoting greater
LTF (29, 30). We observed a similar relationship between
the short-term hypoxic response and LTF in sham-operated rats in the
present study (linear regression, phrenic burst amplitude: r2 = 0.93, P < 0.001; Fig.
5 ), but no such relationship was
detected in CSNX rats (r2 < 0.01, P = 0.841). Although the residual short-term hypoxic response does not appear to explain the variation in LTF in CSNX rats
(several CSNX rats had much greater LTF than expected on the basis of
their hypoxic responses), the average value for CSNX rats falls within
the 95% confidence limits for the LTF-hypoxic response relationship
constructed for sham-operated rats (Fig. 5). Thus it is possible that
the mechanisms underlying the correlation of LTF with short-term
hypoxic responses also apply to whatever responsiveness persists after
peripheral chemodenervation.
|
Alternatively, hypoxia could initiate phrenic LTF independent of increases in respiratory motor output during hypoxia. For example, hypoxia could alter intracellular signaling or gene expression directly at the level of the phrenic motor nucleus or perhaps at higher brain centers. Gallman and Millhorn (20) investigated the effects of hypoxic challenges of varying severity in anesthetized, vagotomized, and carotid body-denervated cats. They found that a single 10-min bout of hypoxia (PaO2 = 36-65 Torr) produced a LTF of phrenic motor output that lasted more than 1 h posthypoxia. Phrenic activity was diminished (often to apnea) during hypoxia in these peripherally chemodenervated cats, but phrenic activity returned to baseline by 15 min posthypoxia and increased progressively throughout the remainder of the protocol; this winding up of phrenic activity posthypoxia is qualitatively similar to what is seen in the anesthetized rat after three episodes of hypoxia (44, present study). The hypoxia-induced LTF observed by Gallman and Millhorn (20) was abolished in decerebrate cats, suggesting involvement of supraspinal mechanisms. In contrast, episodic CSN stimulation is equally or more effective at inducing LTF in unanesthetized, decerebrate vs. CNS-intact cats (40). These data suggest that, at least in cats, hypoxia can initiate a long-lasting facilitation of respiratory motor output through direct CNS effects, apparently by mechanisms distinct from carotid chemoafferent neuron activation. Therefore, it is plausible that CNS hypoxia, independent from hypoxic respiratory responsiveness, contributes to LTF induction in rats. Whether these effects of hypoxia contribute to LTF in carotid body intact animals, or whether they are revealed only in the absence of carotid chemoafferent inputs, is unknown.
Although additional studies are clearly needed to investigate their respective roles in the initiation of LTF, the present study suggests that hypoxia may have an important role in respiratory plasticity apart from the activation of chemoafferent neurons.
| |
ACKNOWLEDGEMENTS |
|---|
We thank Dr. D. D. Fuller for assistance during these experiments.
| |
FOOTNOTES |
|---|
This work was supported by National Heart, Lung, and Blood Institute (NHLBI) Grants HL-65383 and HL-53319. R. W. Bavis was supported by NHLBI Training Grant HL-07654.
Address for reprint requests and other correspondence: R. W. Bavis, Dept. of Comparative Biosciences, School of Veterinary Medicine, Univ. of Wisconsin, 2015 Linden Dr., Madison, WI 53706 (E-mail: bavisr{at}svm.vetmed.wisc.edu).
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. Section 1734 solely to indicate this fact.
July 12, 2002;10.1152/japplphysiol.00374.2002
Received 19 April 2002; accepted in final form 8 July 2002.
| |
REFERENCES |
|---|
|
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
|
|---|
1.
Alcayaga, J,
Varas R,
Arroyo J,
Iturriaga R,
and
Zapata P.
Responses to hypoxia of petrosal ganglia in vitro.
Brain Res
845:
28-34,
1999[Web of Science][Medline].
2.
Babcock, MA,
and
Badr MS.
Long-term facilitation of ventilation in humans during NREM sleep.
Sleep
21:
709-716,
1998[Web of Science][Medline].
3.
Bach, KB,
and
Mitchell GS.
Hypoxia-induced long-term facilitation of respiratory activity is serotonin dependent.
Respir Physiol
104:
251-260,
1996[Web of Science][Medline].
4.
Bach, KB,
and
Mitchell GS.
Hypercapnia-induced long-term depression of respiratory activity requires 
2-adrenergic receptors.
J Appl Physiol
84:
2099-2105,
1998
5.
Baker, TL,
and
Mitchell GS.
Episodic but not continuous hypoxia elicits long-term facilitation of phrenic motor output in rats.
J Physiol
529:
215-219,
2000
6.
Baker, TL,
Fuller DD,
Zabka AG,
and
Mitchell GS.
Respiratory plasticity: differential actions of continuous and episodic hypoxia and hypercapnia.
Respir Physiol
129:
25-35,
2001[Web of Science][Medline].
7.
Baker-Herman, TL,
and
Mitchell GS.
Phrenic long-term facilitation requires spinal serotonin receptor activation and protein synthesis.
J Neurosci
22:
6239-6246,
2002
8.
Behm, R,
Mewes H,
DeMuinck Keizer WH,
Unger T,
and
Rettig R.
Cardiovascular and renal effects of hypoxia in conscious carotid body-denervated rats.
J Appl Physiol
74:
2795-2800,
1993
9.
Brophy, S,
Ford TW,
Carey M,
and
Jones JFX
Activity of aortic chemoreceptors in the anaesthetized rat.
J Physiol
514:
821-828,
1999
10.
Cao, KY,
Zwillich CW,
Berthon-Jones M,
and
Sullivan CE.
Increased normoxic ventilation induced by repetitive hypoxia in conscious dogs.
J Appl Physiol
73:
2083-2088,
1992
11.
Chandel, NS,
and
Schumacker PT.
Cellular oxygen sensing by mitochondria: old questions, new insight.
J Appl Physiol
88:
1880-1889,
2000
12.
Curran, AK,
Rodman JR,
Eastwood PR,
Henderson KS,
Dempsey JA,
and
Smith CA.
Ventilatory responses to specific CNS hypoxia in sleeping dogs.
J Appl Physiol
88:
1840-1852,
2000
13.
Dröge, W.
Free radicals in the physiological control of cell function.
Physiol Rev
82:
47-95,
2002
14.
Dwinell, MR,
Janssen PL,
and
Bisgard GE.
Lack of long term facilitation of ventilation after exposure to hypoxia in goats.
Respir Physiol
108:
1-9,
1997[Web of Science][Medline].
15.
Edelman, NH,
Melton JE,
and
Neubauer JA.
The modulation of peripheral chemoreceptor input by central nervous system hypoxia.
Adv Exp Med Biol
337:
345-352,
1993[Web of Science][Medline].
16.
Erickson, JT,
and
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
348:
161-182,
1994[Web of Science][Medline].
17.
Fuller, DD,
Baker TL,
Behan M,
and
Mitchell GS.
Expression of hypoglossal long-term facilitation differs between substrains of Sprague-Dawley rat.
Physiol Genomics
4:
175-181,
2001
18.
Fuller, DD,
Zabka AG,
Baker TL,
and
Mitchell GS.
Phrenic long-term facilitation requires 5-HT receptor activation during but not following episodic hypoxia.
J Appl Physiol
90:
2001-2006,
2001
19.
Fuller, DD,
Bach KB,
Baker TL,
Kinkead R,
and
Mitchell GS.
Long term facilitation of phrenic motor output.
Respir Physiol
121:
135-146,
2000[Web of Science][Medline].
20.
Gallman, EA,
and
Millhorn DE.
Two long-lasting central respiratory responses following acute hypoxia in glomectomized cats.
J Physiol
395:
333-347,
1988
21.
Gelfand, R,
Lambertsen CJ,
Clark JM,
and
Hopkin E.
Hypoxic ventilatory sensitivity in men is not reduced by prolonged hyperoxia (Predictive Studies V and VI).
J Appl Physiol
84:
292-302,
1998
22.
Gozal, D.
Potentiation of hypoxic ventilatory response by hyperoxia in the conscious rat: putative role of nitric oxide.
J Appl Physiol
85:
129-132,
1998
23.
Hayashi, F,
Coles SK,
Bach KB,
Mitchell GS,
and
McCrimmon DR.
Time-dependent phrenic nerve responses to carotid afferent activation: intact vs. decerebellate rats.
Am J Physiol Regul Integr Comp Physiol
265:
R811-R819,
1993
24.
Honda, Y,
Tani H,
Masuda A,
Kobayashi T,
Nishino T,
Kimura H,
Masuyama S,
and
Kuriyama T.
Effect of prior O2 breathing on ventilatory response to sustained isocapnic hypoxia in adult humans.
J Appl Physiol
81:
1627-1632,
1996
25.
Horn, EM,
and
Waldrop TG.
Suprapontine control of respiration.
Respir Physiol
114:
201-211,
1998[Web of Science][Medline].
26.
Jamieson, D,
Chance B,
Cadenas E,
and
Boveris A.
The relation of free radical production to hyperoxia.
Annu Rev Physiol
48:
703-719,
1986[Web of Science][Medline].
27.
Jones, SC,
Radinsky CR,
Furlan AJ,
Chyatte D,
and
Perez-Trepichio AD.
Cortical NOS inhibition raises the lower limit of cerebral blood flow-arterial pressure autoregulation.
Am J Physiol Heart Circ Physiol
276:
H1253-H1262,
1999
28.
Kinkead, R,
and
Mitchell GS.
Time-dependent hypoxic ventilatory responses in rats: effects of ketanserin and 5-carboxyamidotryptamine.
Am J Physiol Regul Integr Comp Physiol
277:
R658-R666,
1999
29.
Kinkead, R,
Bach KB,
Johnson SM,
Hodgeman BA,
and
Mitchell GS.
Plasticity in respiratory motor control: intermittent hypoxia and hypercapnia activate opposing serotonergic and noradrenergic modulatory systems.
Comp Biochem Physiol A Physiol
130:
207-218,
2001.
30.
Kinkead, R,
Zhan WZ,
Prakash YS,
Bach KB,
Sieck GC,
and
Mitchell GS.
Cervical dorsal rhizotomy enhances serotonergic innervation of phrenic motoneurons and serotonin-dependent long-term facilitation of respiratory motor output in rats.
J Neurosci
18:
8436-8443,
1998
31.
Kline, DD,
Overholt JL,
and
Prabhakar NR.
Mutant mice deficient in NOS-1 exhibit attenuated long-term facilitation and short-term potentiation in breathing.
J Physiol
539:
309-315,
2002
32.
Ling, L,
Olson EB, Jr,
Vidruk EH,
and
Mitchell GS.
Developmental plasticity of the hypoxic ventilatory response.
Respir Physiol
110:
261-268,
1997[Web of Science][Medline].
33.
Ling, L,
Olson EB, Jr,
Vidruk EH,
and
Mitchell GS.
Integrated phrenic responses to carotid afferent stimulation in adult rats following perinatal hyperoxia.
J Physiol
500:
787-796,
1997
34.
Ling, L,
Olson EB, Jr,
Vidruk EH,
and
Mitchell GS.
Phrenic responses to isocapnic hypoxia in adult rats following perinatal hyperoxia.
Respir Physiol
109:
107-116,
1997[Web of Science][Medline].
35.
Ling, L,
Fuller DD,
Bach KB,
Kinkead R,
Olson EB, Jr,
and
Mitchell GS.
Chronic intermittent hypoxia elicits serotonin-dependent plasticity in the central neural control of breathing.
J Neurosci
21:
5381-5388,
2001
36.
Lipton, AJ,
Johnson MA,
Macdonald T,
Lieberman MW,
Gozal D,
and
Gaston B.
S-nitrosothiols signal the ventilatory response to hypoxia.
Nature
413:
171-174,
2001[Medline].
37.
Martin-Body, RL,
Robson GJ,
and
Sinclair JD.
Respiratory effects of sectioning the carotid sinus glossopharyngeal and abdominal vagal nerves in the awake rat.
J Physiol
361:
35-45,
1985
38.
McDonald, DM,
and
Blewett RW.
Location and size of carotid body-like organs (paraganglia) revealed in rats by the permeability of blood vessels to Evans blue dye.
J Neurocytol
10:
607-643,
1981[Web of Science][Medline].
39.
Millhorn, DE.
Stimulation of raphe (obscurus) nucleus causes long-term potentiation of phrenic nerve activity in cat.
J Physiol
381:
169-179,
1986
40.
Millhorn, DE,
Eldridge FL,
and
Waldrop TG.
Prolonged stimulation of respiration by a new central neural mechanism.
Respir Physiol
41:
87-103,
1980[Web of Science][Medline].
41.
Millhorn, DE,
Eldridge FL,
and
Waldrop TG.
Prolonged stimulation of respiration by endogenous central serotonin.
Respir Physiol
42:
171-188,
1980[Web of Science][Medline].
42.
Mizusawa, A,
Ogawa H,
Kikuchi Y,
Hida W,
Kurosawa H,
Okabe S,
Takishima T,
and
Shirato K.
In vivo release of glutamate in nucleus tractus solitarii of the rat during hypoxia.
J Physiol
478:
55-66,
1994
43.
Mitchell, GS,
Powell FL,
Hopkins SR,
and
Milsom WK.
Time domains of the hypoxic ventilatory response in awake ducks: episodic and continuous hypoxia.
Respir Physiol
124:
117-128,
2001[Web of Science][Medline].
44.
Mitchell, GS,
Baker TL,
Nanda SA,
Fuller DD,
Zabka AG,
Hodgeman BA,
Bavis RW,
Mack KJ,
and
Olson EB, Jr.
Intermittent hypoxia and respiratory plasticity.
J Appl Physiol
90:
2466-2475,
2001
45.
Mitchell, RA,
Sinha AK,
and
McDonald DM.
Chemoreceptive properties of regenerated endings of the carotid sinus nerve.
Brain Res
43:
681-685,
1972[Web of Science][Medline].
46.
Morris, KF,
Shannon R,
and
Lindsey BG.
Changes in cat medullary neurone firing rates and synchrony following induction of respiratory long-term facilitation.
J Physiol
532:
483-497,
2001
47.
Morris, KF,
Arata A,
Shannon R,
and
Lindsey BG.
Long-term facilitation of phrenic nerve activity in cats: responses and short time scale correlations of medullary neurones.
J Physiol
490:
463-480,
1996
48.
Morris, KF,
Baekey DM,
Shannon R,
and
Lindsey BG.
Respiratory neural activity during long-term facilitation.
Respir Physiol
121:
119-133,
2000[Web of Science][Medline].
49.
Neubauer, JA,
Simone A,
and
Edelman NH.
Role of brain lactic acidosis in hypoxic depression of respiration.
J Appl Physiol
65:
1324-1331,
1988
50.
Neubauer, JA,
Santiago TV,
Posner MA,
and
Edelman NH.
Ventral medullary pH and ventilatory responses to hyperperfusion and hypoxia.
J Appl Physiol
58:
1659-1668,
1985
51.
Olson, EB, Jr,
Vidruk EH,
and
Dempsey JA.
Carotid body excision significantly changes ventilatory control in awake rats.
J Appl Physiol
64:
666-671,
1988
52.
Olson, EB, Jr,
Bohne CJ,
Dwinell MR,
Podolsky A,
Vidruk EH,
Fuller DD,
Powell FL,
and
Mitchell GS.
Ventilatory long-term facilitation in unanesthetized rats.
J Appl Physiol
91:
709-716,
2001
53.
Peng, Y,
Kline DD,
Hamadani A,
and
Prabhakar NR.
Induction of long term facilitation in the carotid body activity by intermittent hypoxia (Abstract).
FASEB J
15:
A153,
2001.
54.
Powell, FL,
Milsom WK,
and
Mitchell GS.
Time domains of the hypoxic ventilatory response.
Respir Physiol
112:
123-134,
1998[Web of Science][Medline].
55.
Prabhakar, NR.
Oxygen sensing by the carotid body chemoreceptors.
J Appl Physiol
88:
2287-2295,
2000
56.
Prabhakar, NR.
Oxygen sensing during intermittent hypoxia: cellular and molecular mechanisms.
J Appl Physiol
90:
1986-1994,
2001
57.
Solomon, IC.
Excitation of phrenic and sympathetic output during acute hypoxia: contribution of medullary oxygen detectors.
Respir Physiol
121:
101-117,
2000[Web of Science][Medline].
58.
Solomon, IC,
Edelman NH,
and
Neubauer JA.
Pre-Botzinger complex functions as a central hypoxia chemosensor for respiration in vivo.
J Neurophysiol
83:
2854-2868,
2000
59.
Sun, MK,
and
Reis DJ.
Dopamine or transmitter release from rat carotid body may not be essential to hypoxic chemoreception.
Am J Physiol Regul Integr Comp Physiol
267:
R1632-R1639,
1994
60.
Turner, DL,
and
Mitchell GS.
Long-term facilitation of ventilation following repeated hypoxic episodes in awake goats.
J Physiol
499:
543-550,
1997
61.
Walker, JK,
and
Jennings DB.
Respiratory effects of pressor and depressor agents in conscious rats.
Can J Physiol Pharmacol
76:
707-714,
1998[Web of Science][Medline].
62.
Wang, W,
Tiwari JK,
Bradley SR,
Zaykin RV,
and
Richerson GB.
Acidosis-stimulated neurons of the medullary raphe are serotonergic.
J Neurophysiol
85:
2224-2235,
2001
63.
Xu, F,
Sato M,
Spellman MJ, Jr,
Mitchell RA,
and
Severinghaus JW.
Topography of cat medullary ventral surface hypoxic acidification.
J Appl Physiol
73:
2631-2637,
1992
64.
Yates, BJ,
Goto T,
and
Bolton PS.
Responses of neurons in the caudal medullary raphe nuclei of the cat to stimulation of the vestibular nerve.
Exp Brain Res
89:
323-332,
1992[Web of Science][Medline].
65.
Zabka, AG,
Behan M,
and
Mitchell GS.
Long term facilitation of respiratory motor output decreases with age in male rats.
J Physiol
531:
509-514,
2001
66.
Zabka, AG,
Behan M,
and
Mitchell GS.
Time-dependent hypoxic respiratory responses in female rats are influenced by age and by the estrus cycle.
J Appl Physiol
91:
2831-2838,
2001
This article has been cited by other articles:
|
|
 |
|
  K.-Z. Lee, P. J. Reier, and D. D. Fuller Phrenic Motoneuron Discharge Patterns During Hypoxia-Induced Short-Term Potentiation in Rats J Neurophysiol, October 1, 2009; 102(4): 2184 - 2193. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 S. Ryan and P. Nolan Episodic hypoxia induces long-term facilitation of upper airway muscle activity in spontaneously breathing anaesthetized rats J. Physiol., July 1, 2009; 587(13): 3329 - 3342. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. Mahamed and G. S. Mitchell Simulated apnoeas induce serotonin-dependent respiratory long-term facilitation in rats J. Physiol., April 15, 2008; 586(8): 2171 - 2181. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A. Tadjalli, J. Duffin, Y. M. Li, H. Hong, and J. Peever Inspiratory activation is not required for episodic hypoxia-induced respiratory long-term facilitation in postnatal rats J. Physiol., December 1, 2007; 585(2): 593 - 606. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 H. Gu, M. Lin, J. Liu, D. Gozal, K. E. Scrogin, R. Wurster, M. W. Chapleau, X. Ma, and Z. Cheng Selective impairment of central mediation of baroreflex in anesthetized young adult Fischer 344 rats after chronic intermittent hypoxia Am J Physiol Heart Circ Physiol, November 1, 2007; 293(5): H2809 - H2818. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 S. R. Reeves, G. S. Mitchell, and D. Gozal Early postnatal chronic intermittent hypoxia modifies hypoxic respiratory responses and long-term phrenic facilitation in adult rats Am J Physiol Regulatory Integrative Comp Physiol, June 1, 2006; 290(6): R1664 - R1671. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
M. McGuire, Y. Zhang, D. P. White, and L. Ling Phrenic long-term facilitation requires NMDA receptors in the phrenic motonucleus in rats J. Physiol., September 1, 2005; 567(2): 599 - 611. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T. A. Day and R. J. A. Wilson Specific carotid body chemostimulation is sufficient to elicit phrenic poststimulus frequency decline in a novel in situ dual-perfused rat preparation Am J Physiol Regulatory Integrative Comp Physiol, August 1, 2005; 289(2): R532 - R544. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K. J. Cummings and R. J. A. Wilson Time-dependent modulation of carotid body afferent activity during and after intermittent hypoxia Am J Physiol Regulatory Integrative Comp Physiol, June 1, 2005; 288(6): R1571 - R1580. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A. G Zabka, G. S Mitchell, and M Behan Ageing and gonadectomy have similar effects on hypoglossal long-term facilitation in male Fischer rats J. Physiol., March 1, 2005; 563(2): 557 - 568. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R. W. Bavis, E. B. Olson Jr, E. H. Vidruk, D. D. Fuller, and G. S. Mitchell Developmental plasticity of the hypoxic ventilatory response in rats induced by neonatal hypoxia J. Physiol., June 1, 2004; 557(2): 645 - 660. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Y. Zhang, M. McGuire, D. P White, and L. Ling Episodic phrenic-inhibitory vagus nerve stimulation paradoxically induces phrenic long-term facilitation in rats J. Physiol., September 15, 2003; 551(3): 981 - 991. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Visit Other APS Journals Online |
, n = 6) or episodic hyperoxia (
, n = 6).
Time-matched normoxic time control rats (
,
n = 6) were treated the same but were maintained in
normoxia throughout the protocol. Values are means ± SE.
* Different from baseline, P < 0.05; +different
from normoxic control group at same time point, P < 0.05; #different from episodic hyperoxia group at same time point,
P < 0.05.
, n = 8) were treated
the same but were maintained in hyperoxia throughout the protocol.
Values are means ± SE. * Different from baseline,
P < 0.05; +different from corresponding hyperoxic
control group (i.e., same surgery) at same time point,
P < 0.05; #different from corresponding sham-operated
group (i.e., same treatment, hypoxia or control) at same time point,
P < 0.05. A: amplitude;
B: frequency; C: minute
activity.
