INTRODUCTION

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RECENT WORK IN HUMANS, RATS, and ducks has demonstrated a time-of-day-dependent change in respiratory responsiveness to hypoxic and hypercapnic stimuli (24, 29, 33, 38), suggesting that the respiratory system is modulated by the circadian timing system.

It is well established that manipulation of many clock-controlled variables will affect clock function (21), which raises the possibility that there may be a reciprocal interaction between the circadian timing and the respiratory control systems. An effect of respiratory stimuli on circadian timing mechanisms could have important implications, for example, by contributing to sleep disruption in disorders, such as sleep apnea syndromes and nocturnal asthma, in which hypoxia and hypercapnia occur (23), as well as for the design of long-term respiratory studies in which daily repetitions of stimuli are used.

Several studies report data that suggest that chemical respiratory stimuli could have long-lasting effects on the circadian clock. A study on simulated high altitude (7,620 m) in a hypobaric chamber, during which the oxygen masks of three human subjects were removed for 2-3 min, concluded that acute exposure to severe hypoxia caused a transient phase shift in several physiological variables (peak expiratory air flow, grip strength, and oral temperature) (2). However, the subjects were maintained in a light-dark (LD) cycle after the test, thus complicating interpretation of apparent phase shifts.

Another study examined the potential role of elevated CO₂ concentration ([CO₂]) in circadian rhythm disturbances (32). Four male subjects were studied in an environmental chamber before and during 24 days of elevated inspired [CO₂] (~2%). Activity (wrist actigraphy), rectal temperature, and urine analyses (cortisol, catecholamines, and 6-hydroxymelatoninsulphate) were reported. Small effects on circadian temperature rhythm amplitudes were observed, but confounding factors, including LD cycle and competing experimental objectives, rendered the data inconclusive.

Recently, two groups (20, 31) concluded that prolonged exposure (7 days and 63 h, respectively) to hypoxia or hypercapnia can reduce the amplitude of circadian temperature rhythms in rats. One of these studies (31) kept the animals on a 12:12-h LD cycle, preventing an analysis of the effects of hypoxia on circadian phase and period. In the other study (20), poststimulus period and phase were found to be unaffected under free running conditions. However, a prolonged hypoxic stimulus could conceivably have had opposing effects on phase at different times of day that resulted in no net phase change. Analyses of phase and period during the stimulus were prevented by the severe attenuation of the rhythm amplitude (20).

To reexamine this question, we studied wheel running behavior in golden hamsters (Mesocricetus auratus), because this has been shown to be a reliable and precise marker of circadian phase in this species. Hamsters were held in an environment free of time cues and were exposed to scheduled pulses of hypoxia or hypercapnia to test the hypothesis that respiratory stimuli can induce long-lasting changes in the phase, period, and quantity of wheel running activity.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Animals and Housing

Eight hamster cages were placed in a single sealed animal chamber. Cage walls were opaque, thus preventing visual contact between experimental subjects. The chamber was impermeable to external light and was equipped with timer-controlled fluorescent lights that provided ~140 lux light intensity at the level of the cage floor. Four fans ensured full mixing of the air in the chamber and cages, and, except during pulses, the animal chamber was ventilated continuously with air at a rate sufficient to maintain [CO₂] < 0.5%. Although ammonia was not measured, the high ventilation rate and mixing fans were assumed to prevent accumulation of this gas as they did for CO₂. The air or test gas was distributed evenly within the chamber via a multiport ventilation duct and four mixing fans. Gas exhaust ports were designed to minimize pressure fluctuations when gas flow rates were adjusted and to prevent light from entering the chamber. Temperature within the closed chamber was 20-22°C.

Respiratory stimuli (pulses) were presented simultaneously to all hamsters by manipulation of the oxygen concentration ([O₂] = 8% for hypoxia) or [CO₂] (11% for hypercapnia) of the air within the animal chamber. Immediately before the pulse, normal chamber air ventilation was terminated. Hypoxia was induced by briefly blowing pure N₂ into the chamber, and hypercapnia was induced by briefly blowing pure CO₂ into the chamber. No attempt was made to maintain isooxic conditions, and elevation of [CO₂] during hypercapnic pulses produced a mild reduction in [O₂] (to ~17%). Gases were supplied from pressurized gas cylinders, and control of [O₂] or [CO₂] was automated. Gas flow to the chamber (air, N₂, or CO₂) was controlled by a series of solenoid valves. Power supply to the solenoid valves was controlled by a timer that was set to a cycle of 3 h on, 21 h off. A paramagnetic O₂ analyzer (model 572, Servomex) and an infrared CO₂ analyzer (model CD-3A, Ametek) was used to monitor chamber [O₂] and [CO₂], respectively. The analog voltage outputs of the analyzers were connected to a custom-built electronic comparator that closed the solenoid valve supplying N₂ or CO₂ when gas concentrations exceeded the user-defined limits (i.e., <8% for [O₂] and >11% for [CO₂] in the present study). A small leak prevented further depletion of O₂ or accumulation of CO₂ due to animal metabolism, and the solenoids were briefly triggered approximately every 20 min to maintain gas concentrations at their target levels. At the end of the 3-h pulse, airflow was reinstated. "Sham" pulses using compressed air instead of N₂ or CO₂, were used as a control for factors such as noise, humidity, or pressure changes (10).

Experiment 1: Effect of Scheduled Respiratory Stimuli on Wheel Running Behavior

Hamsters were kept in a LD cycle (14-h light, 10-h darkness) to align their circadian rhythms. Cages were changed, and then the hamsters were released into DD for at least 7 days or until a stable free-running period was apparent. The hamsters (now aged 250-255 days) were then exposed to daily 3-h pulses of hypoxia ([O₂] = 8%) for 7 days, starting at ~CT6. The respiratory stimuli were given at CT6 because established nonphotic zeitgebers are known to be effective at this time (21). CT6 was estimated for each hamster on the first pulse day by extrapolation of the free running onset times of the previous 5 days; the pulse was initiated at the average CT6 for the group and then at the same clock time each day thereafter. After the last pulse day, hamsters were allowed free run in DD for a minimum of 7 additional days. Cages were then changed, and the animals were returned to LD for at least 7 days. This procedure was repeated in the same animals for pulses of hypercapnia ([CO₂] = 11%, [O₂] = 17%) at age 285-290 days and for sham (compressed air) pulses at age 337-342 days.

In preliminary control studies, two different groups of eight male hamsters were used. In one group, the phases of wheel running onset were separated by entraining the animals to individual 14:10-h LD cycles in which lights off occurred at different times of day. The animals were then placed in the animal chamber in DD, and the sham test procedure was conducted as described above. The first sham pulse began at the following CTs: 0, 1, 5, 7, 8, 12, 18. In this group of animals, the free running period () was >24 h [i.e., pulse period (T) < ]. In another group of hamsters,  was <24 h when sham experiments were conducted (i.e., T > ).

Experiment 2: Physiological Effects of Hypoxic and Hypercapnic Pulses at CT6

Respiratory gas exchange (metabolic rate) was estimated by open-flow respirometry (37). The time constant of the system was 35 s; thus gas concentrations were at steady state by 3 min after a change in flow. [O₂] and [CO₂] of inlet gas and the outlet gases of each of the four animal chambers were measured sequentially for ~15 s each. [O₂] and [CO₂] were measured using O₂ and CO₂ analyzers (models S-3A/1 and 3D-3A, Ametek). The gas passed through a desiccating column (Drierite) immediately before entering the analyzers.

Gas exchange measurements were immediately followed by lung ventilation measurements. This involved closing the inlet and outlet of an animal chamber and recording the pressure deflections induced by lung ventilation within the closed chamber (3, 4). A differential pressure transducer (model DP45-14, Validyne Engineering) was used to measure the difference in pressure between the thermobarometer reference chamber and an animal chamber. Pressure deflections were calibrated in each animal chamber by injections of 1 ml of air with the animals in place. Animal chamber temperatures were assumed to equal that measured in the reference chamber, as confirmed in preliminary studies. Relative humidity in each animal chamber was assumed to equal that measured in the combined outflow gas.

Body temperature (T_b) of each animal was monitored throughout the experiment by calibrated radiotransmitters (model TA10TA-F20, Data Sciences International) that were implanted into the peritoneal cavity while the hamster was under halothane anaesthesia. The animals were given 2 wk to recover from the surgery before experiments began. The radio signal was detected by loop antennae, which were coiled around each Plexiglas cylinder and connected to a telemetry receiver (model RTA-500, Minimitter). The receiver was connected to a digital data acquisition system (Maclab/8, AD Instruments) via a signal conditioner that converted radiofrequency pulses into 3-V square waves. T_b was encoded in the pulse frequency.

The hamsters (now aged 379-384 days) were weighed and then placed into the animal chambers under dim red light ~1 h before CT6. A lid shielded the animal chamber, and a small hole was used for visual observation of the animals under dim red light. The hamsters were exposed to 3-h pulses of air, hypoxia, or hypercapnia on 3 separate days, and at least 2 days separated experiments for each animal.

Gas exchange and ventilatory measurements were taken three times before the pulse. Pulses were initiated at CT6, and, after 5 min, gas exchange and ventilation were measured. Measurements were repeated every 20 min thereafter for 3 h. Air was reintroduced to the system after the last pulse measurement was taken at 3 h, and measurements were then taken at 5, 20, 40, and 60 min of the recovery period. After the experiments, the hamsters were removed from the animal chamber and returned to their home cages.

Data Analysis

T calculation. Although the respiratory stimuli began at the same clock time each day, the time to final pulse concentration was variable due to changes in inlet gas flow rates. Pulse onset was arbitrarily defined as the time that the chamber gas concentration reached 50% of the final pulse concentration. T was calculated using the slope of a least-squares linear regression fitted through the seven sequential daily pulse onset times.

calculation. Activity period () was calculated as the slope of a least-squares linear regression fitted through the wheel running onset times. "Prepulse"  was calculated using the activity onset times on the 5 days immediately preceding the first pulse day. "Pulsed"  was based on data from the last 5 of the 7 days on which respiratory pulses were delivered. "Postpulse"  was calculated for activity onsets on the third through seventh days after the last pulse day. Pulsed  was compared with T and the prepulse  to evaluate whether entrainment had occurred.

Phase shift calculation. Both pre- and postpulse regression lines were extrapolated to the first pulse day. The phase shift was defined as the difference between the onset times predicted by the two regression lines for the first pulse day.

Activity level calculation. Activity level was defined as total wheel revolutions per day. Mean prepulse activity levels were calculated using the 5 days immediately before the first pulse day. Mean pulsed activity levels (total wheel revolutions per day) were based on data from the last 5 of the 7 pulse days. Prepulse and pulsed activity levels were compared to determine whether the respiratory stimuli affected the quantity of wheel running.

Calculation of ventilatory parameters. Inspired ventilation (I) = f × VT, where f is the number of breaths per min, and VT is the tidal volume (ml, BTPS). VT was calculated using the equations developed by Drorbaugh and Fenn (4)

(1)

where P_m is the respiratory pressure deflection; P_cal is the pressure deflection of a known volume of gas; V_cal is the volume of gas injected into the chamber; T_b is body temperature (°K); T_c is animal chamber temperature (°K); PB is barometric pressure in the chamber (mmHg); PA_H2O is saturated water vapor pressure of the gas in the alveoli (mmHg); and PC_H2O is water vapor pressure of the gas in the chamber (mmHg).

Calculation of metabolic rate. Metabolic rate was estimated as the rate of oxygen consumption (O₂, ml/min, STPD), as measured by open flow respirometry (37)

(2)

where is dry gas flow entering the animal chamber (ml/min, STPD); FI_O2 is the fractional concentration of oxygen entering the chamber; and FE_O2 is the fractional concentration of oxygen leaving the chamber.

(3)

Statistical analysis. All statistics were computed using SYSTAT software (SPSS). Significance was tested using a general linear model, with sample time and hamster as factors. A general linear model was used because it accounts for the variability between individuals and allows for missing data points. When a significant difference was obtained, a Tukey's post hoc test was used for multiple pairwise comparisons. Differences are considered significant at the 95% confidence level (P < 0.05).

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Experiment 1: Effect of Scheduled Respiratory Stimuli on Phase, Period, and Quantity of Wheel Running

Controls. Daily wheel revolutions and the phase and period of onset of wheel running were unaffected by 3-h pulses of air beginning at CT6 on 7 consecutive days (Fig. 1). This was also confirmed in two additional groups of hamsters: when the first pulse began at a variety of CTs (0, 1, 5, 7, 8, 10, 12, and 18) and when T >  and T <  (data not shown). Consequently, the prepulse days from each experiment were used to measure the effect of each respiratory stimulus to minimize the effect of possible differences (e.g., age-related) between experiments. In addition, a separate group of hamsters was exposed to a novel wheel for 3 h, beginning at CT6, which induced phase advances of 3.2 ± 0.1 h. This positive control (12) demonstrated that large phase shifts are possible under the present laboratory conditions.

View larger version (18K): [in this window] [in a new window]   Fig. 1.   Cumulative phase shifts (mean ± SE; n = 8) after 7 consecutive days of 3-h pulses [delivered at circadian time (CT)6-CT9] of hypoxia (O2 concentration = 8%), hypercapnia (CO2 concentration = 11%), and control tests in which air was substituted for test gas. * Significantly different from control.

Hypoxia. Seven consecutive days of 3-h hypoxic pulses ([O₂] = 8%) beginning at CT6 caused a cumulative phase delay of 46.4 ± 6.9 min (Figs. 1 and 2). Hamsters exhibited a prepulse  of 23.88 ± 0.01 h. Postpulse  (23.86 ± 0.02 h) was not significantly different from prepulse . In the last five hypoxic pulse days,  (23.93 ± 0.02 h) was not significantly different from T (23.96 ± 0.00 h). Daily wheel revolutions on days with hypoxic pulses (6,395 ± 901 wheel revolutions, 3.71 ± 0.53 km) were significantly reduced from the prepulse values (9,180 ± 1,378 wheel revolutions, 5.32 ± 0.80 km).

View larger version (54K): [in this window] [in a new window]   Fig. 2.   Actograms of representative animals illustrating phase change of wheel running activity due to hypoxic (A) and hypercapnic (B) pulses. Animals were free running in constant darkness. Each line of the actogram represents a 24-h day, and consecutive days are plotted beneath each other. The ordinate for each day is plotted as a histogram of distance run (m) per 2 min, vertical scale 0-150 m. Horizontal, double-headed arrows indicate the time of hypoxic or hypercapnic pulses. Dashed regression lines illustrate the wheel running rhythm periods over 5 days before (prepulse) and after (postpulse) the respiratory pulses. The cumulative phase delay induced by the pulses was calculated by extrapolation of the regressions to the first pulse day.

Hypercapnia. Seven consecutive days of 3-h hypercapnic pulses at CT6 caused a cumulative but nonsignificant (P = 0.143) phase delay of 25.9 ± 12.3 min (Figs. 1 and 2). Postpulse  (23.87 ± 0.02 h) was not significantly different from prepulse  (23.89 ± 0.01 h). In the last five hypercapnic pulse days,  (23.94 ± 0.02 h) was not significantly different from T (23.95 ± 0.00 h). Daily wheel revolutions on days with hypercapnic pulses (6,504 ± 1,127 wheel revolutions, 3.77 ± 0.67 km) were significantly lower than the prepulse values (8,255 ± 1,703 wheel revolutions, 4.79 ± 0.99 km).

Experiment 2: Physiological Effects of Hypoxic and Hypercapnic Pulses at CT6

Controls. Figure 3 shows that a 3-h pulse of air (sham pulse) at CT6 had no significant effect on lung ventilation, O₂, or T_b. Furthermore, for the group as a whole, the sham pulse values were not significantly different from those measured during the prepulse interval in the hypoxia and hypercapnia experiments. Consequently, to minimize the effect of possible differences between days in individual animals, the prepulse measurements from each experiment were used to measure the effect of each respiratory stimulus.

View larger version (60K): [in this window] [in a new window]   Fig. 3.   Respiratory and metabolic responses to 3-h pulses of air (), hypoxia (; O2 concentration = 8%), and hypercapnia (; CO2 concentration = 11%) beginning at CT6. Pulses were initiated at 60 min and concluded at 240 min. A: body temperature (Tb). B: minute inspired ventilation (I, BTPS). C: rate of oxygen consumption (O2). D: metabolism-specific ventilation (I/O2).Values are means ± SE; n = 8. * Significantly different from mean prepulse value (P < 0.05).

Hypoxia. Visual observations indicated that hamsters were initially aroused from sleep by introduction of 8% O₂ into the plethysmograph. After 3-4 min, the hamsters curled up and appeared to resume sleeping. Sleep was occasionally interrupted by periods of feeding and grooming. It is assumed that similar behavioral events occurred during pulses in experiment 1, but no attempt was made to account for changes in behavioral state in the data analyses. Hamsters started to shiver immediately after the return to air. Shivering was visible for several minutes.

Hypercapnia. Behavioral responses of the hamsters to 11% [CO₂] were similar to those during hypoxia, with the exception that shivering was not observed during posthypercapnic recovery in air.

Correlation Analysis

View larger version (29K): [in this window] [in a new window]   Fig. 4.   Regressions of wheel-running activity (A), Tb (B), O2 (C), and I (D) vs. phase change in 8 hamsters using pooled data (n = 22; 2 points were removed from analysis due to running wheel malfunction) from the 3 test conditions. , Air; , hypoxia; , hypercapnia.

	DISCUSSION

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The results of this study support the hypothesis that chemical respiratory stimuli can influence the output of the circadian pacemaker, as measured by the timing and intensity of wheel running activity in the golden hamster.

Quantity of Wheel Running

Saiki and Mortola (29) have shown that acute hypoxia can abolish the day-to-night differences in lung ventilation, metabolic rate, and T_b in 6-day-old rat pups. Furthermore, prolonged (7 days) hypoxia and hypercapnia reduced the amplitudes of circadian rhythms in T_b, O₂, and general body movements in adult rats (20). Salloum et al. (31) also observed a suppression of T_b rhythm amplitude in rats exposed to 63 h of continuous hypoxia. The present study differs from these studies in that our hamsters were normoxic and normocapnic for >3 h before activity onset during each of the pulse days, but wheel running activity was still reduced. It is of interest to note that, in a preliminary study, our laboratory found that hamsters began running on the wheel ~2-4 h after the termination of a 3-h hypoxic pulse initiated at CT11 (12). This suggests (but does not prove) that the daily reduction in wheel running activity may be a masking effect of the stimulus on activity, rather than an affect on the amplitude of the output of the circadian pacemaker. Further research is needed to resolve this question.

Period of Wheel Running Rhythm

Phase of Wheel Running Rhythm

Nonphotic stimuli presented at CT6 usually evoke phase advances in the hamster wheel running rhythm (21), whereas we have found that respiratory stimuli presented at this time cause phase delays. The majority of nonphotic stimuli that have been shown to affect long-term changes in circadian phase, including access to a novel wheel, social interactions, and bolus injections of benzodiazepines (26, 35), have been shown to induce an acute increase in locomotor activity during the stimulus. It is unclear whether activity per se, or a correlated variable such as arousal, novelty, motivation, or reward, is critical in producing the phase shift (21). Significantly, Van Reeth et al. (34) found that enforced immobility induced phase delays in hamsters when imposed in subjective night but not in subjective day. In the present study, both hypoxia and hypercapnia had delayed effects on activity, reducing wheel running intensity during subjective night. This presents a possible mechanism for the hypoxia- and hypercapnia-induced phase delays, a suggestion that is supported by the significant correlation between the magnitude of the phase shift and the depression of wheel running (Fig. 4).

Physiological Responses to Pulses and Relationships to Circadian Phase Shifts.

T_b. Several studies have found that, whereas the circadian system is temperature compensated, temperature pulses can nevertheless induce phase shifts and ambient temperature cycles can entrain circadian rhythms both in vivo and in vitro (5, 28). In the present study, hypoxia caused a significant reduction in T_b, and hypercapnia induced a transient, but nonsignificant, reduction in T_b. Acute profound hypothermia has been shown to induce phase shifts in hamsters and other rodents (8, 9, 25, 27), suggesting that acute T_b changes may have mediated the phase responses to respiratory gas pulses. However, the studies cited above used intense environmental cooling to induce hypothermia and are therefore not directly comparable to the present study. Furthermore, these earlier studies did not control for confounding behavioral correlates. For example, examination of the published representative actograms (9, 25, 27) reveals that hypothermic pulses usually reduced subsequent nocturnal activity levels, even when hypothermia occurred during subjective day. Thus phase delays generated by cold-induced hypothermia (9, 25, 27) and hypoxia or hypercapnia (present study) could both be an indirect effect of reduced levels of activity (34).

O₂. The hypoxic hypometabolism observed in the present experiment has previously been observed in numerous species (6). The reduction in O₂ during acute hypoxia has been attributed to an inhibition of thermogenesis (19). In the present study, the decrease in T_b during hypoxia and the threefold increase in O₂ coincidental with visible shivering after the return to air are consistent with this hypothesis.

I. Hypoxia has been reported to evoke a sustained increase in I/O₂ in hamsters and other rodents (6, 18, 19), and this was confirmed in the present experiment. The ventilatory response to hypercapnia was also consistent with responses previously reported for the hamster (3, 11, 18, 36). The increase in I found in the present study was primarily caused by changes in VT. Respiratory frequency remained relatively constant in hypoxia and hypercapnia. The ventilatory responses to pulses were not correlated with phase shifts.

Perspectives

A similar argument can be made regarding the potential role of hypoxia and hypercapnia in disruption of circadian organization of sleep-wake cycles in sleep-disordered breathing syndromes. Any effect of respiratory gas fluctuations on circadian timing is likely to be inconsequential, unless the human circadian timing system is considerably more sensitive to these stimuli than that of the hamster.

Finally, because respiratory chemoreflex responsiveness varies with time of day (22, 33) and manipulation of respiratory gases causes circadian phase delays, a respiratory stimulus on one day might alter the response to the same stimulus (at the same clock time) on the next day. This "feedback" effect would be of significance to investigators who use respiratory stimuli on successive days in chronic animal models of respiratory control. Again, however, we must conclude that this is likely to be of little or no practical importance, because the phase shift induced by a relatively intense stimulus was <5 min/day, and phase shifts of several hours would be required to appreciably change ventilatory chemoreflexes (22, 33). Furthermore, our preliminary work showed that maintaining hamsters in a 14:10-h LD cycle during three consecutive days of hypoxia did not alter the phase angle of entrainment to the LD cycle (12). Thus a normal LD cycle can negate the small effects of respiratory stimuli.

In conclusion, this study has confirmed that hypoxia and hypercapnia can influence the phase of the circadian pacemaker of the hamster and thus raises interesting questions about the mechanisms involved. However, the effect was modest and is unlikely to represent a confounding factor in experimental design or in the etiology of respiratory-related sleep disorders.