Cardiovascular Pulmonary Research Laboratory, Department of Medicine, University of Colorado Health Sciences Center, Denver, Colorado 80262
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ABSTRACT |
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Studies in humans indicate genetic effects on the ventilatory response to hypoxia, but the site of these effects is unknown. The present study explores the question of whether there are genetically directed effects on the intrinsic hypoxic chemosensitivity of the carotid body. The approach was to study these responses in two inbred rat strains [spontaneously hypertensive rats (SHR) and Fischer 344 (F-344)] and to measure in vivo carotid chemosensitivity as the change in carotid sinus nerve (CSN) activity during progressive, isocapnic hypoxia and the isolated, in vitro responses of excised superfused carotid bodies, loaded with the fluorimetric indicator fura 2, measured as the cytosolic calcium response to moderate hypoxia (PO2 = 55 mmHg). CSN responses in F-344 rats (n = 12) were uniformly low, with a shape parameter A of 13.8 ± 6.59 (SE), whereas responses in SHR (n = 15) were sevenfold higher (108 ± 24.1; P < 0.002) and showed greater variation. In vitro, intracellular calcium responses of superfused carotid bodies estimated from the fluorimetric ratio (340/380 nm) showed a greater peak increase during hypoxia in carotid bodies from SHR (140 ± 4.7%) than from F-344 rats (114 6.0%; P < 0.01). Our results indicate strain-related differences in hypoxic chemosensitivity that are intrinsic to the carotid body and that could mediate genetic effects on ventilatory responsiveness to hypoxia.
chemoreceptor; calcium; genetic; hypertension; spontaneously hypertensive rat; Fischer rat; carotid sinus nerve
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INTRODUCTION |
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VENTILATORY RESPONSES to hypoxia are subject to marked variation both within and among individuals (23), and this variation has important effects on ventilatory adaptation to high altitude (20, 30, 34, 40, 47), and to lung disease (13, 27), and are correlated with performance in athletics (4, 6, 39) and mountain climbers (40). Early studies indicated that some of this variation is acquired as in the decreased response produced by chronic hypoxia (19, 42, 58) and the increased sensitivity seen with increased metabolic rate (61) or with short-term hypoxia (38, 60).
However, it is also clear that these responses show considerable variation among basal, "normal" individuals, with nearly an eightfold range of responses found in humans (23) and in cats (57). These differences are reproducible with repeated testing and thus seem to constitute stable interindividual differences. It is also evident that this variable hypoxic sensitivity is nonrandomly distributed. The first indications were the finding of clusters of low responders in families of patients with unexplained hypoventilation (24, 33) or obstructive lung disease (13, 27, 29, 35) and in endurance athletes (41). That such variation might have a genetic basis is suggested by concordance of these responses within pairs of identical twins (7, 28) and by differences in ventilatory pattern and responses to hypoxia and hypercapnia in inbred strains of mice (50-52).
Little is known about the nature of this apparent genetic effect on the hypoxic ventilatory response. In particular, it is unclear whether such influences act at the peripheral chemoreceptor (the carotid body), within the central nervous system, or on the peripheral ventilatory apparatus. The present study was undertaken to explore the question of whether there are genetically directed effects on the intrinsic hypoxic chemosensitivity of the carotid body. The approach was to study these responses in two inbred rat strains selected for apparently divergent responses in a preliminary survey and to measure in vivo carotid chemosensitivity as carotid sinus nerve (CSN) activity. To determine whether the findings reflect differences intrinsic to the carotid body, we compared in vitro responses of excised superfused carotid bodies from the two strains. We measured the response to hypoxia as the cytosolic calcium because free intracellular calcium is thought to be integral to stimulus-response coupling in the carotid body.
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METHODS |
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Experimental Animals
We studied rats weighing between 280 and 450 g, with ages ranging from 95 to 310 days. They were obtained from colonies of spontaneously hypertensive rats (SHR) and Fischer 344 rats (F-344) maintained at the same breeding site and had been inbred through >28 generations by the provider (Harlan Sprague Dawley, Indianapolis, IN). All protocols were approved by the Institutional Animal Care and Use Committee.In Vivo Studies
Rats were anesthetized with an initial dose of chloralose (90 mg/kg) and urethan (450 mg/kg) intraperitoneally, followed by intravenous doses of 30 and 150 mg/kg, respectively, which we have found to produce consistent surgical levels of anesthesia that are stable over 4 h. Core temperature was maintained at 37-38°C with a heating pad servo-controlled by a rectal temperature probe. Catheters were inserted in the femoral vein for supplemental administration of anesthetic and in the femoral artery for monitoring of arterial pressure. After tracheostomy, a tracheal cannula was inserted into the distal trachea and connected to a small-animal ventilator (model 683; Harvard) set to a rate and tidal volume to produce an end-tidal PCO2 of 30-34 Torr. Animals were paralyzed with gallamine (15 mg/kg). In some cases, an expiratory water column was added to maintain positive end-expiratory pressure (10 cmH2O). Respiratory gases were measured by O2 fuel cell (Applied Technical Products, Denver, CO; see Ref. 59) and a CO2 infrared analyzer (LB-2; Sensor Medics, Anaheim, CA), which were calibrated with gases previously analyzed by the Scholander technique. Signals were recorded on a four-channel strip-chart recorder (model 2400; Gould, Cleveland, OH).The reliability of end-tidal O2 tension (PETO2) as an indicator of arterial values was tested in six animals by comparison with simultaneous values produced by an O2 electrode (no. 16-730A; Microelectrodes), calibrated with Scholander-analyzed gases, and placed in a femoral arteriovenous loop, with continuous flow maintained by a roller pump at 0.6 ml/min. The system has a total dead space of 1 ml, and with abrupt changes, values lagged end tidal by 5-10 s. This lag was measured in each case, and data were time adjusted for comparison. End-tidal and arterial O2 tensions were tightly correlated over the entire range (Fig. 1). Because of the shape of the response curves, the range of O2 tensions from 40 to 80 mmHg is particularly important in accurate definition of curve shape. Over this range, 78 measurements in four rats produced an average end-tidal arterial difference of 1.2 ± 4.35 (SD) mmHg. We were unable to perform these measurements during neural recording because the impaired hemostasis produced by the anticoagulation needed for the flow-through system produced bleeding that interfered with CSN signal acquisition.
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CSN recording. After the proximal trachea and esophagus were ligated and transected to expose the carotid sinus region, the CSN was carefully stripped of surrounding tissue. The discrete nerve leading from the carotid sinus that carries baroreceptor afferents was transected (37); we have found that, after this procedure, the residual CSN signal contains none, or only very slight, cardiosynchronous activity (<10% total signal in hyperoxia and much less in hypoxia). Carotid body neural output was recorded by platinum bipolar electrodes from the whole desheathed nerve bundle (28). The amplified signal was filtered (100-3,000 Hz), sampled at a rate of 1 kHz, and processed to measure the variance of the amplitude of the whole nerve signal, as previously described (8). This measurement reflects the summation of action potentials and is proportional to both the activity of independently firing individual fibers and the number of active fibers. The width of this distribution (amplitude variance) provides a useful index of whole nerve activity (8, 45). CSN activity was measured during progressive reduction of the inspired O2 fraction by nitrogen dilution to produce a fall in end-tidal tension from a starting value of ~300 to a final level of 30-35 mmHg over 3-5 min. A minimum of two, but most often three to five, runs separated by 5-10 min were averaged in each animal. To minimize variation due to nerve preparation and nerve-electrode contact, the initial CSN activity in hyperoxia was arbitrarily set to equal 1.0, with subsequent values expressed as relative change. These responses were measured under isocapnic conditions stabilized by constant ventilator settings adjusted to set the PCO2 in each animal at levels measured initially before gallamine-induced paralysis.
CSN responses to hypoxia were measured as the shape parameter A, which
describes the hyperbolic relationship between
PETO2 and CSN activity
described by the equation CSN = CSN0 + A/(PETO2 26), where CSN is
the observed sinus nerve activity,
CSN0 is the horizontal asymptote
of nerve activity, and A is a measure of the curvature of the
relationship such that higher values for A indicate greater response.
The constant 26 represents the
PETO2 at which the slope approaches
infinity, which was determined empirically in previous studies to
produce an optimum curve fit for rats. The shape parameters for each
run were averaged for individual animals, and comparisons were made
across the two strains by two-sample t-test for unequal variance.
In Vitro Studies
Animals and anesthesia used in these studies were identical to those of the in vivo protocols. After tracheostomy and institution of ventilatory support, the carotid arterial bifurcation was exposed, and loose ligatures were placed around all major arterial branches. Two rats, one of each strain, were prepared in this fashion. The CSN was then exposed, secured with a ligature, and transected. After preparation of both animals, heparin (1,000 units) was administered intravenously via a femoral venous catheter. The nerves and carotid bifurcation were then dissected free of surrounding tissue, all vascular ligatures were tied, and the carotid bodies with nerve were dissected free of the bifurcation and immediately transferred to an iced, oxygenated superfusate bath. The superfusate contained (in mM) 120 NaCl, 5 KCl, 0.5 MgCl2, 2.2 CaCl2, 10 glucose, 5 HEPES, 1.9 NaHCO3, and 1 glutathione, pH 7.4. Under a dissecting microscope, the carotid bodies were cleaned of connective tissue and placed on the stage of the microfluorimetry apparatus. The process of removal/cleaning was accomplished within 10-15 min, and the sequence of removal was alternated across the two strains.The pair of carotid bodies was then loaded with the fluorimetric calcium indicator fura 2-AM (3 mM) with pluronic F-127 (5 mM) for 30 min at room temperature, and excess dye was then removed by continuous superfusion with a solution equilibrated with 55% O2-5% CO2. A 20-min period was allotted for deesterification of the internalized indicator, which proved sufficient to produce stable baseline (hyperoxic) values.
Fluorimetric measurements were made by sequential imaging of paired carotid bodies excited at alternate wavelengths of 340 and 380 nm using a computer-controlled filter wheel (Ludl). Epifluorescence at 510 nm was measured by an intensified charge-coupled device camera, and data were stored to a 286/16 MHz personal computer (Everex). Imaging, intensity measurements, and ratioing were performed with a parallel processor and software (Quantex Fluorescent Microscopy, Sunnyvale, CA).
Pairs of carotid bodies were imaged on a coverslip positioned in the well of a stage-mounted heating device (Warner Instrument RC-30L) servo-regulated by a thermocouple and maintained at 37°C immersed in the superfusate. Superfusate was supplied by gravity flow from two reservoirs bubbled with either 55% O2-5% CO2 (hyperoxia baseline) or 5% O2-5% CO2 (hypoxia) at a flow rate of 1 ml/min. The superfusate was routed through a flow-through PO2 electrode (Microelectrodes no. 16-730A) positioned immediately next to the stage. Under hyperoxic baseline conditions, the superfusate PO2 was 315-320 mmHg, and during hypoxic perfusion, the PO2 was 50-55 mmHg. The stage was covered with an acrylic dome, and the dead space above the preparation was perfused with gas of identical composition to that used in the bubbler supplying the superfusate.
Each run consisted of exposure of the loaded pair of carotid bodies to the hyperoxic perfusate for a baseline period of 10-15 min during which three to five images were acquired. The superfusate was then switched to the hypoxic mixture, which produced a fall in PO2 from initial values of 315-320 mmHg to a stable plateau of 50-55 mmHg, which was nearly complete by 5 min. Paired images of each carotid body were then acquired at 5, 10, and 15 min of hypoxia followed by a final pair of images after 20 min of hyperoxia. Time-control measurements were performed in carotid bodies of each strain and maintained in hyperoxia for a period corresponding to the duration of baseline hypoxic exposure in the experimental group.
Image Analysis
Images of whole carotid bodies were divided into five to seven discrete regions of interest and were measured as the intensity ratio at 340/380 nm. Because of uncertainties concerning the precise relationship of fluorescence ratio to absolute intracellular calcium concentrations, we used the ratios as indicators of relative changes in free calcium ions. To minimize day-to-day variation, we expressed values during hypoxia as changes from baseline. Our predetermined goal was to test for within-pair differences in responses to hypoxia of carotid bodies from the two strains. The ratios for individual regions were averaged to produce a composite value for each carotid body at each experimental condition. Values for the paired carotid bodies were tested for the effects of experimental condition and strain-associated differences by two-way ANOVA with repeated measures. Results are expressed as means ± SE. ![]() |
RESULTS |
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Baseline data for rats studied in the in vivo protocols are summarized in Table 1. There were small but significant differences in age and body weight between strains. As expected, mean arterial blood pressure was substantially higher in the SHR.
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In Vivo Studies
CSN responses are shown in Fig. 2 for typical individual rats of each strain, with steeper response for the SHR rat and the usual hyperbolic shape for the responses of both. Responses, summarized as fitted curves, for the entire series are shown in Fig. 3, which indicates that rats of the F-344 strain show uniformly low responses, whereas those of the SHR are greater and more variable.
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Results analyzed as the shape parameter A, a measure of steepness of response, are shown in Fig. 4, which indicates that SHR had on average a nearly eightfold steeper CSN response to hypoxia (P < 0.002) but also showed much greater within-strain variability.
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As mentioned, there were significant differences in age and body weight between groups that could have influenced our findings, so we tested for a within-group relationship of shape parameter A vs. each of these variables and found no significant correlation for either that indicated these age and weight differences were unlikely contributors to the findings.
We also found no correlation, within strains, of the hypoxic response with either arterial partial pressure of CO2, arterial pH, hematocrit or mean arterial pressure, which suggested that these variables were unlikely contributors to among- or within-strain differences.
In Vitro Results
Loading with fura 2 produced homogenous uptake of the indicator over the entire carotid body. With subsequent exposure to hypoxia, there was a diffuse increase in the fluorescence ratio, indicating a rise in cytosolic calcium (Fig. 5).
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Results with seven pairs of carotid bodies showed that hypoxia (PO2 = 55 mmHg) induced a substantially greater increase in fluorescence ratio in carotid bodies of SHR vs. F-344 rats (Fig. 6).
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When averaged over a time interval of 5-15 min, responses were 34% greater in carotid bodies of SHR than of F-344 rats (P < 0.01). Because there was some variation in the timing of the peak response for individual carotid bodies, we also compared the peak responses of carotid bodies of the two strains independent of the time of occurrence and again found that responses of SHR carotid bodies exceeded those of F-344 rats (Fig. 7).
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Control measurements in carotid bodies observed for 15 min under baseline conditions showed a peak change in the fluorescence ratio, which averaged 7 ± 0.49% in F-344 rats (n = 6) and 23.4 ± 8.78% in SHR (n = 6). This difference among strains was of borderline significance (P < 0.08) and may represent continued low-level deesterification of unhydrolyzed fura 2-AM (ester).
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DISCUSSION |
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These studies present evidence of differences in hypoxic carotid body chemosensitivity among two inbred rat strains. Recordings of CSN responses to progressive, isocapnic hypoxia produced uniformly low responses in rats of the F-344 strain, whereas those in SHR were more variable and on average sevenfold higher. Parallel findings were evident in hypoxic responses of isolated, superfused carotid bodies of the two strains, measured as fluorimetric estimates of the increase in free, cytosolic calcium. Carotid bodies of the SHR showed a nearly threefold greater increase than those of the F-344 strain.
Our rationale for the use of parallel in vivo and in vitro approaches was that the former would address the question of whether such differences exist in intact, living animals, whereas the latter would indicate whether these differences could be traced to an effect intrinsic to the carotid body. Our findings point to the existence of in vivo strain-related differences in chemoreceptor hypoxic chemosensitivity, and the in vitro findings indicate intrinsic differences in hypoxic responsiveness of the carotid body of the two strains.
The combined use of in vivo and in vitro approaches also presents the advantage of offsetting pitfalls and artifacts inherent in each approach. Data from in vivo recordings of CSN activity have a number of potential difficulties, including possible effects of anesthesia, signal sampling error, mode of signal analysis, and contamination of chemoreceptor signal by baroreceptor and efferent traffic. There are also potential confounding effects via sympathetic innervation, which was left intact in our preparation. It seems unlikely that anesthetic effects contributed to our findings because, although ventilatory responses to hypoxia are clearly depressed by anesthesia, we have previously demonstrated that CSN responses are only slightly affected by large changes in depth of anesthesia (57), and in our study the average dose of anesthetic and apparent depth of anesthesia were similar for the two groups.
Other problems with the in vivo studies include the use of recordings from the intact, uncut CSN, with the result that afferent chemoreceptor activity may have been contaminated by descending, efferent traffic. We have previously found in cats that CSN responses recorded in intact nerves and in neural segments distal to proximal transection produce similar hypoxic responses and that interindividual differences in the CSN response are similar for both the intact and sectioned nerve preparations (53, 56). Our use of relative, rather than absolute, increase in sinus nerve activity is intended to minimize the effect of variation due to differences in electrode nerve contact and other systematic differences among individual preparations, but such an approach could be misleading if there were differences in absolute neural activity for groups being compared. In our study, there were no systematic differences in basal (hyperoxic) CSN variance for the two strains.
Additional potential confounding variables in the in vivo studies include differences in age, weight, hematocrit, and systemic arterial blood pressure, but the absence of within-group correlations for these factors suggests that they are unlikely to explain observed differences among strains. Finally, the findings in the in vivo preparation do not indicate whether the observed differences among strains reflect effects intrinsic to the carotid body or were due to humoral influences such as circulating catecholamines, potassium levels, or to other unknown factors. The complementary use of the in vitro studies was intended to address these reservations.
Measurements in isolated, superfused carotid bodies have the advantage of freedom from confounding effects of humoral and CNS influences. The use of superfusion also avoids effects due to hypertension-induced changes in the vasculature of the carotid body (2, 17). We used estimated changes in cytosolic calcium as the indicator of hypoxic chemosensitivity because of broad evidence suggesting that calcium mediates the coupling of the stimulus, hypoxia, to the response, CSN activity (3, 15, 43). Tight correlation of a rise in cytosolic calcium with neural response hypoxia is found by most (3, 15, 43), but not all, investigators (11), and the neural response is attenuated by antagonists and enhanced by agonists of calcium channels (43).
Our approach presented several problems, including those inherent in a superfused preparation, which include difficulty in estimating the magnitude of the internal (tissue) hypoxic stimulus. The responses were also delayed relative to the prompt increase in neural activity, which likely reflects the time for washout of the reservoir and the slow lowering of internal tissue O2 tension by the superfusate. Nonetheless, we used this approach because several studies have indicated that, compared with superfused carotid bodies, perfused preparations are generally nonresponsive to hypoxic stimulation (12, 26, 36), although one study suggests responsiveness in a perfused preparation (25). Nonetheless, superfusion complicates the interpretation of the actual, internal stimulus to the carotid body, and because of this, we intentionally used a mild degree of hypoxia (PO2 = 55 mmHg) and high perfusion flow rates. We were also careful to remove all connective tissue surrounding the carotid body by dissection under a microscope.
Another problem is the uncertainty in translation of the observed changes in fluorescence measurements into precise values for tissue free calcium because of difficulties in accurate calibration encountered in a tissue mass that is more uncertain than techniques used for individual cells. For this reason, we have expressed our results only as fluorimetric ratios and analyzed responses as relative change. There is also uncertainty about the specific cellular origin of the measured responses. Our data only indicate that these responses were intrinsic to the whole carotid body but cannot be localized to a specific cell type. Although it is likely that these changes reflect events in chemosensory cells, considering the linkage of the rise in calcium to the fall in O2 tension, there could be contributions from downstream secondary elements such as neural conduit cells. However, we noted that fluorimetric signals of the CSN, often seen in our images, were uniformly unaltered during hypoxia, suggesting that the observed fluorimetric changes were not nonspecific. Although the observed relative changes in fluorimetric ratio with hypoxia were less than those seen in the CSN traffic, this probably reflected the mild extent of the stimulus, which was intentionally chosen to maximize the physiological relevance of the findings. Indeed end-tidal and arterial O2 tensions comparable to those of our superfusate produce only minor increases in in vivo CSN activity. It is also likely that our observed calcium changes were a blend of increases in responsive cells with unchanged levels in nonchemosensory cells, which would attenuate the net response of the overall carotid body. Finally, although there is clear concordance of our in vivo and in vitro findings, this does not prove that the latter are the cause of the former.
Although these studies point to strain-dependent differences in carotid body sensitivity to hypoxia, they do not indicate the factors within the organ responsible for this effect. Recent work points to variation in the calcium response to hypoxia among individual cells isolated from single carotid bodies (5), and strain-associated variation in carotid body responses could thus reflect differential respresentation of cell subpopulations of different sensitivity. Alternatively, these different carotid body responses might be attributed to homogenous alterations in the intrinsic hypoxic-sensing mechanisms. Comparative studies of single cells from high and low responder strains would be useful in resolving this question.
Our finding of interstrain differences in carotid body chemosensitivity may reflect broader, fundamental differences between these rat strains. Specifically, several observations suggest the existence of a linkage of chemosensitivity and systemic arterial blood pressure. Several strains of SHR show both increased vasoreactivity and resting hyperventilation, which is abolished by hyperoxia (reviewed in Refs. 16 and 54). Conversely, F-344 rats have been shown to have diminished pulmonary vasoreactivity (21). Hypertensive rats are found to have enlarged carotid bodies with increased content of norepinephrine, but not dopamine (1, 16), and enhanced peripheral chemoreceptor sensitivity (14). However, in none of these studies could the influence of systemic neural or humoral effects be separated from differences intrinsic to the carotid body. A similar association of systemic hypertension and increased chemosensitivity is noted in humans with essential hypertension who demonstrate resting hyperventilation, which is quenched by hyperoxia, and a markedly increased (4-fold) hypoxic ventilatory response with no increase in hypercapnic response (48, 49, 55).
Enhanced chemosensitivity could either represent a primary genetic feature of hypertension or be acquired, perhaps as a secondary consequence of hypertension. Indeed, changes in small glomic arteries, including wall thickening and luminal narrowing, have been found in older SHR with established hypertension, but changes in ventilatory and carotid body hypoxic sensitivity and size are most pronounced in young animals with new onset hypertension (16, 18). Similarly, in humans, increased chemosensitivity is most pronounced in young patients with recent onset of hypertension (48, 55) and is also found in their normotensive relatives (49). Such differences are absent in renal hypertension, suggesting a specific link to essential hypertension.
Thus, on balance, findings in both rats and humans suggest that altered hypoxic ventilatory control and carotid body function are genetically directed attributes rather than secondary consequences of hypertension. Some authors have suggested that increased chemosensitivity may play a role in the pathogenesis of hypertension in view of the observations that chemoreceptor stimulation increases blood pressure (16). Indeed, brief hyperoxia produces an exaggerated fall in systemic blood pressure and vascular resistance in hypertensive patients and in normotensive relatives of such patients (49), and hypoxia evokes a greater rise in peripheral sympathetic nerve activity in borderline hypertensive patients than in normal subjects (44). On the other hand, high-altitude exposure, which should produce tonic chemoreceptor stimulation, seems to blunt rather than augment hypertension in SHR (22, 31), which suggests that increased chemoreceptor activity is probably not a primary cause of the hypertension.
Although reasons for the apparent linkage of essential or spontaneous hypertension to enhanced hypoxic chemosensitivity are unclear, it is evident that patients with essential hypertension and SHR show fundamental alterations in cell membranes that alter ion fluxes (9) and membrane fluidity (10), which have been implicated in altered intracellular calcium homeostasis. These changes are transmitted as a single autosomal dominant gene (32) and include an increased inward sodium leak possibly linked to reversed sodium ion-calcium ion exchange. Calcium binding to plasma cell membranes has a "membrane-stabilizing effect" that reduces calcium influx (9). A decrease in calcium binding sites on plasma membranes has been found in patients with essential hypertension, which could promote calcium influx and contribute to increased intracellular calcium levels (9). Together, these effects may explain the observed increases in basal and stimulated levels of cytosolic calcium in arterial smooth muscle (9). These changes are not seen in renal or other forms of hypertension, persist through multiple passages in culture (46), and are thus thought to be primary and genetically determined rather than secondary features of essential or spontaneous hypertension. Furthermore, it is clear that these differences are not confined to vascular smooth muscle, are evident in erythrocytes, platelets, neutrophils, and lymphocytes, and thus might involve membranes of all cells (9).
These global effects could, by addition or synergy, augment the calcium response to hypoxia in the carotid body and thus explain a potential linkage of hypertension and enhanced chemosensitivity. However, this idea that hypertension and chemosensitivity are mechanistically linked remains speculative. The present work provides no direct support for such a linkage because our limited strain comparison cannot exclude the existence of normotensive strains with high chemosensitivity or hypertensive strains with low sensitivity.
In summary, our studies point to the presence of strain-associated influences on the intrinsic chemosensitivity of the carotid body that could contribute to the genetic effects on the hypoxic ventilatory response. Further work to explore the molecular basis of genetically directed influences on chemosensitivity is needed to better understand the mechanism of this effect and may clarify the nature of hypoxic sensing mechanisms.
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ACKNOWLEDGEMENTS |
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This work was supported by National Institutes of Health (NIH) Program Project Grant HL-5P01-HL-14985, Training Grant 5T32-HL-07171, and Shared Instrumentation Grant S10 RR-05803. M. G. Dickinson was supported by NIH Short-Term Training Grant 5T35 DK-07496-08.
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FOOTNOTES |
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Address for reprint requests: J. V. Weil, Cardiovascular Pulmonary Research Laboratory, B-133, Univ. of Colorado Health Sciences Center, Denver, CO 80262.
Received 14 October 1997; accepted in final form 9 January 1998.
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