INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

THE GROWTH OF OUR KNOWLEDGE about the membrane potentials in living cells owes much to the development of fluorescent probes, fluorescence measurements, and imaging technologies (14, 17, 40, 45). Glomus cells (type I) of the carotid body (CB), which are sensitive to hypoxia, have been the least studied in terms of fluorescence measurements of plasma and mitochondrial membrane potentials (E_m and _m, respectively). Interestingly, the cellular mechanisms of CB O₂ sensing rest mainly on glomus cell E_m and _m measurements (1, 11, 16).

In favor of the ion channel/membrane hypothesis of CB O₂ sensing, various investigators, using the patch-clamp technique, showed in rabbit and rat glomus cells the inhibition of outward K⁺ currents (I_K) and membrane depolarization during hypoxia (12-20 Torr PO₂) (11, 27, 37, 46). However, the same technique, when used in intact rat CBs, produced no significant change in membrane resistance or outward current during anoxia (15). Furthermore, with a different technique using sharp electrode impalement of isolated rat glomus cells, hypoxia (induced by sodium dithionate) caused hyperpolarization in 64%, depolarization in 29%, and no effect in 7% of the cells tested (36). On the other hand, cyanide, which consistently increased cat CB neural discharge (33), caused E_m hyperpolarization and increased cytosolic free Ca²⁺ concentration ([Ca²⁺]_i) in rabbit glomus cells (5). Hence, it is not clear whether glomus cell depolarization is a critical event in hypoxic stimulation. Conventional direct measurement of membrane potential with a patch-clamp electrode is difficult, particularly for small and delicate cells, such as glomus cells (8-15 µm diameter), in which the membrane does not seal properly. Moreover, because the technique is invasive, the risk of electrical and mechanical perturbation remains and could be the reason for distortion of results. It is therefore reasonable to use a noninvasive method to overcome these perturbations and variations of glomus cell E_m. As such, fluorescence measurement using voltage-sensitive dye could be a useful means for indirect evaluation of glomus cell E_m.

The major problem encountered in exploration of glomus cell _m is the minute size of the CB and the yield of mitochondria, which is far too small to permit conventional analysis. Thus an imaging technique using voltage-sensitive dyes could be the mainstay to assess glomus cell _m changes in response to different stimuli. According to the metabolic/mitochondrial hypothesis of CB O₂ sensing, glomus cell mitochondria are depolarized during moderate and severe hypoxia (16). Previously, Duchen and Biscoe (16), using rhodamine 123, measured glomus cell _m photometrically during hypoxia (moderate and severe) and reported graded depolarization in response to graded hypoxia (between 40 and 0 Torr PO₂). The disadvantage of using rhodamine 123 is that all mitochondria would fluoresce with equal intensity, irrespective of their potentials, and would fail to reveal any heterogeneity in fluorescent intensity. Furthermore, rhodamine forms H-aggregates (not J-aggregates), which quench the dye fluorescence (40). Thus an increase in dye uptake as a result of higher _m may not necessarily lead to a brighter fluorescence; it may even reduce the fluorescence to an extent that such mitochondria become undetectable (21). Hence, rhodamine 123 has not been used very widely for continuous assessment of _m, and we were prompted to use a reliable probe (JC-1) to reinvestigate glomus cell _m.

In the present study, we have attempted for the first time the use of two voltage-sensitive fluorescent probes, bis-oxonol and JC-1, to measure indirectly the glomus cell E_m and _m, respectively. Furthermore, we have reinvestigated the relative importance of glomus cell E_m and _m responses to acute hypoxia by a fluorescence microimaging technique. We found that glomus cells that consistently depolarized in response to high K⁺ did not respond (45%), hyperpolarized (35%), or depolarized (20%) in response to acute hypoxia. However, all PC-12 cells depolarized in response to hypoxia. Finally, all glomus cell mitochondria consistently depolarized in response to hypoxia and high CO. Hence, the voltage-sensitive probe bis-oxonol could not always confirm the glomus cell membrane depolarization as a critical event, whereas JC- 1 consistently indicated glomus cell mitochondrial depolarization during hypoxia. Overall, our imaging data support a hypothesis of chemoreception in the CB, which we named the metabomembrane hypothesis (see DISCUSSION).

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Glomus cells. Glomus cells were obtained from adult Sprague-Dawley rats (200-250 g) by enzymatic separation, as described previously (30). Briefly, rats were anesthetized with pentobarbital sodium (60-80 mg/kg ip; Abbott Laboratories) and tracheotomized, and the CBs were surgically removed from the carotid bifurcation and placed in a chamber filled with ice-cold HEPES buffer (pH ~7.4) and bubbled with 100% O₂. After removal of the CB, the animals were killed by an intracardiac injection of pentobarbital sodium (100 mg/kg). The CBs were cleaned of connective tissue under a dissecting microscope and collected in a glass vial containing 0.2% collagenase (type IV, Sigma Chemical) in a Ca²⁺- and Mg²⁺-free Tyrode solution for digestion for 30 min at 37°C. The digested tissue was transferred to a solution of growth medium (85% Ham's F-12 containing HEPES and L-glutamine, 10% fetal calf serum, 5% horse serum, and penicillin-streptomycin), triturated with a fire-polished Pasteur pipette, and then centrifuged at 100-200 g for 5-6 min. The pellet was resuspended in fresh growth medium and plated on sterile 15-mm poly-D-lysine (Sigma Chemical)-coated coverslips on a petri dish. The petri dish containing the separated cells was left undisturbed for 24 h in a humidified incubator (37°C, circulated with 5% CO₂ and air). The glomus cells were identified by using the following criteria: 1) granular birefringent appearance, 2) presence of large nuclei, 3) positive fluorescence of the selected cells stained for catecholamine (by sucrose-phosphate-glyoxalic acid) (30), and 4) significant depolarization of E_m in response to high K⁺.

PC-12 cells. PC-12 cells (rat adrenal pheochromocytoma, CRL-1721, American Type Culture Collection) were acquired from the cell center facility of the University of Pennsylvania. The cells were maintained in growth medium consisting of 85% Ham's F-12 containing 15 mM HEPES and 2 mM L-glutamine, 10% fetal bovine serum, 5% normal horse serum, and penicillin-streptomycin (100 U/ml and 100 µg/ml, respectively). For E_m measurement, PC-12 cells were resuspended in fresh culture medium at low cell density and plated on sterile coated coverslips (15 mm diameter). The cells were allowed to incubate at 37°C under 5% CO₂ and air for 24 h before they were used.

Solutions. Cells were superfused with various solutions. The composition of the basic superfusate, modified Tyrode solution with CO₂-HCO, was as follows: 112 mM NaCl, 4.7 mM KCl, 21.4 mM NaHCO₃, 2.2 mM CaCl₂, 1.1 mM MgCl₂, 22.0 mM sodium glutamate, 5.0 mM glucose, 5 mM HEPES, and 4 g/l dextran (74,200 mol wt). The osmolarity of the buffer was within the normal range of 290-300 mosM. Four separate solutions were initially prepared in 100-ml syringes from the Tyrode buffer: 1) 100% O₂, 2) 100% N₂, 3) 100% CO₂, and 4) 100% CO. Hypoxic solution was prepared by proportionately mixing 100% O₂, 100% N₂, and 100% CO₂ to obtain a required PO₂ of ~10 Torr. Similarly, high-CO (~500 Torr PCO, 125-130 Torr PO₂) solution was prepared by proportionately mixing 100% O₂, 100% N₂, 100% CO₂, and 100% CO. PCO₂ of all the solutions was kept between 33 and 35 Torr and pH at ~7.4. PCO was determined from previously published reports (31), and PO₂ was kept low because the intracellular Ca²⁺ response is evident only at <30 Torr PO₂ (11). All experimental solutions were measured for PO₂, PCO₂, PCO (indirectly), and pH in a blood gas monitor (model PHM73, Radiometer) before use as well as after the experiments. We also measured the pH and gas concentrations of the solutions by sampling from the outlet port of the perfusion chamber to ensure that the parameters did not change during the experiments. For the elevated extracellular K⁺ ([K⁺]_e) solution, equimolar KCl was substituted for NaCl. All test solutions were prepared in airtight glass syringes (50 ml) and stored at 37°C until they were used for superfusion of cells.

Superfusion system. The coverslip containing cells was placed horizontally on a closed-bath imaging chamber (Warner Instrument, Hamden, CT). The small-volume (70-µl) chamber (model RC-20H) ensures a linear flow and fast solution exchange. Perfusate flow rate was maintained by gravity and adjusted to ~1 ml/min. The chamber was mounted on a heated (37°C) platform (Warner Instruments) on an inverted microscope.

Measurement of glomus cell E_m. Optical methods of measuring membrane potentials using voltage-sensitive probes were introduced by Cohen and Salzberg (14). The potential-sensitive probes can be divided into two categories on the basis of their response mechanism: fast- and slow-response probes. In the present study, we selected a slow-response probe, bis-oxonol [bis-(1,3-dibutylbarbituric acid)trimethine oxonol (3), 516.4 mol wt; Molecular Probes, Eugene, OR] to measure cell E_m. The reasons for using this probe are as follows: 1) It has a higher magnitude of optical response: bis-oxonol responds within minutes or seconds and shows a 2% fluorescence change per millivolt, whereas fast-response probes respond within milliseconds but show a 2-10% fluorescence change per 100 mV (19). 2) It detects small potential change: fast dyes are often unable to detect small changes, whereas slow dyes detect small changes. Because glomus cell membrane depolarization due to hypoxia is small (+5 to +13 mV) (46), bis-oxonol is the appropriate dye for our study. 3) Negative charges of bis-oxonol ensure that the dye does not accumulate in mitochondria (29). Therefore, despite its intracellular localization, bis-oxonol is a valuable probe for monitoring changes in E_m. The lipophilic anionic bis-oxonol molecules permeate the cell membrane and undergo a potential-dependent distribution between the cytoplasm and the plasma membrane by a Nernst equilibrium (3). The mechanism underlying the increase in bis-oxonol fluorescence with cellular membrane depolarization is usually ascribed to dye partitioning between extracellular free dye and plasma membrane or cytosol (6) or, probably, orientation of dye molecules within the membrane (2). Repolarization or hyperpolarization of the plasma membrane results in extrusion of the dye and, thus, a decrease in fluorescence (3). Calibration of bis-oxonol fluorescence for indirect assessment of E_m is done with high K⁺ (1). In the present study, aliquots of 100 µM bis-oxonol solution were prepared in DMSO and stored in Eppendorf tubes (20°C). For dye loading, cells in HEPES buffer (pH ~7.4) in the dark and room air at 25°C were incubated in the presence of 1 µM bis-oxonol for 30-45 min. This concentration yielded the greatest signal-to-noise ratio with the glomus and PC-12 cells and also was used previously for other cells (6, 8). For measuring E_m, glomus cell fluorescence was excited at 490 nm and measured at 520 nm.

Measurement of glomus cell _m. Some of the slow dyes form aggregates in certain environments accompanied by a shift of the absorption maximum to a shorter wavelength (H-aggregates) or to a longer wavelength (J-aggregates) (40). Among the various J-aggregate-forming dyes, JC-1 (5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolyl carbocyanine iodide) was used in the present study to probe cell _m. The main reasons are as follows: 1) It is widely used for probing mitochondria in living cells (13). 2) Depending on the difference in _m, JC-1 can exist in monomers as well as in aggregates (40). 3) It is not quenched inside the cell (40). This lipophilic membrane-permeant cation (JC-1) has been shown to be distributed between the cytosol and mitochondria according to Nernstian equilibrium (40). JC-1 selectively enters the mitochondria and exists in a monomeric form, emitting green fluorescence at 527 nm after excitation at 490 nm. However, with the increase in average _m (above 190 mV), JC-1 is able to form J-aggregates emitting red fluorescence with a large shift in emission (590 nm) (40). Because the dye is not significantly quenched in the cell, an increase in JC-1 fluorescence is used to indicate relative hyperpolarization, whereas a decrease in fluorescence intensity is used to indicate mitochondrial depolarization (13, 40). Calibration of the JC-1 fluorescence for relative measurement of _m with carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP) is possible only in isolated mitochondria (40). In the present study, aliquots of JC-1 (100 µM) solution were made in DMSO and stored in Eppendorf tubes (20°C). For effective dye loading, cells were incubated in an experimental concentration of 1 µM JC-1 in HEPES buffer for 30-45 min at 25°C in room air.

Microscopy and fluorescence. Cells were viewed with a Nikon Eclipse TE300 fluorescence microscope (×60 and ×100 oil-immersion objective) and equipped with an optical filter changer (Lambda DG-4, Sutter Instruments, Novato, CA). Excitation of the cells was accomplished with a mercury lamp (150 W) fiber-optic light source, and appropriate filter sets were used as follows: for bis-oxonol, model HQ480/40 exciter, model 505LP dichroic, and model HQ510LP emitter; for JC-1, model HQ500/20 exciter, model 515LP dichroic, and model HQ520LP emitter (Chroma Technology Brattleboro, VT). To prevent rapid photobleaching of the fluorescent preparation, a neutral-density filter (ND 0.3, Chroma Technology) was used to attenuate 50% of the light intensity. The fluorescent images of bis-oxonol- and JC-1-stained cells were acquired during a 10- and a 5-ms exposure time, respectively, with a computer-controlled 12-bit digital cooled charge-coupled device camera (ORCA 100, Hamamatsu), using graphics control software (MetaMorph Imaging System, Universal Imaging). Functional glomus cells were identified as more-or-less round cells with diffuse green fluorescence for bis-oxonol. Glomus cell mitochondria appeared with bright red fluorescence for J-aggregates of JC-1. The regions of interest were digitally marked, and the pixel intensities within the region were then averaged together to obtain a measure of the fluorescence intensity of an individual cell. Changes in fluorescence intensity of the region of interest were acquired before, during, and after stimulation. The background regions outside the cells were also digitally marked, and the average pixel intensity was subtracted from each cell image. All images were in pseudocolor, and the intensity was analyzed with MetaMorph software.

Patch-clamp study. Glomus cell I_K were measured in high PCO by using the whole cell configuration of the perforated patch-clamp technique (Axon Instruments). The coverslip with attached glomus cells was transferred to a recording chamber (Warner Instruments, Hamden, CT) and mounted on a heated plateform (37°C) on an inverted microscope (Nikon). The chamber was perfused by gravity from a 50-ml syringe containing normoxic solution (120-130 Torr PO₂): 140 mM NaCl, 5 mM KCl, 1.8 mM CaCl₂, 1 mM MgCl₂, 10 mM HEPES, 10 mM glucose, and 100 µM ATP, with pH adjusted to ~7.0 with NaOH. Whole cell patch-clamp recordings were made by using patch pipettes (2-6 M resistance) filled with solution composed of 20 mM KCl, 90 mM potassium glutamate, 10 mM HEPES, 1 mM CaCl₂, 2 mM MgCl₂, 10 mM EGTA, 10 mM glucose, and 100-200 µg/ml nystatin, with pH adjusted to 6.8. Previously, it was reported that the CB chemoreceptors to high CO were equally stimulated at outside pH 6.8 with HEPES buffer and at outside pH ~7.4 with CO₂-HCO + HEPES buffer (35). Hence, we used HEPES buffer at pH 6.8 for the patch-clamp study and CO₂-HCO + HEPES buffer at pH 7.4 for the imaging study. Whole cell recordings were performed by using a patch-clamp amplifier (Axopatch 200B, Axon Instruments). The voltage-clamp commands were generated by using DigiData 1200 interface (Axon Instruments) connected to a personal computer (model E-4200, Gateway). The 90-ms depolarizing voltage steps from 80 to +60 mV in 10-mV increments were used to elicit the outward currents. Signals were filtered at 1 kHz and acquired at 10 kHz. Data analyses were performed from the average currents over the range 78-88 ms of the voltage steps. The mean current-voltage relationship was calculated from seven glomus cells. pCLAMP 8.0 (Axon Instruments) was used for data acquisition and analysis.

Statistical analyses. Changes in the fluorescence intensity (arbitrary units) are expressed as percentage of basal response (control), which was considered zero. Values are means ± SE of experiments for each condition. Replicate experiments were carried out by using cells from a separate isolation. Differences among groups were evaluated with one-way ANOVA by using SigmaStat (Jandel Scientific). For the patch-clamp study, significance was determined with a paired t-test. Statistical significance was determined at P < 0.05, and n indicates the number of experiments.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Bis-oxonol fluorescence responses of glomus cell E_m with hypoxia, high [K+]_e, and high PCO. Twenty CBs from 10 rats were used for glomus cell E_m studies. Figure 1A shows the fluorescent images of glomus cells labeled with bis-oxonol (isolated and clustered) during normoxia (~130 Torr PO₂). Switching the perfusion from normoxia to hypoxia (~10 Torr PO₂) did not produce any detectable change of cell fluorescence, which remained stable throughout the period of exposure (0-180 s; Fig. 1A). This was followed by a return to normoxia, which allowed the cells to recover (not shown). With abrupt change of the perfusate to 25 mM K⁺ (Fig. 1A), the fluorescence increased (red color) with time (0-180 s), indicating membrane depolarization. Thus the same glomus cells that were insensitive to low PO₂ responded to high [K⁺]_e with depolarization. In another study, a glomus cell was activated with hypoxia, as indicated by an increase in fluorescence, suggesting membrane depolarization (Fig. 1B). Surprisingly, glomus cells also showed a decrease in fluorescence with hypoxia, indicating that the membrane was hyperpolarized (not shown). Figure 1C shows the time course of average percent change in bis-oxonol fluorescence. Twenty glomus cells were separately studied during normoxia (control) for 240 s to assess the photobleaching effect. Of 40 glomus cells exposed to hypoxia, 18 cells showed no significant change in bis-oxonol fluorescence, which represented 0.3 ± 0.54% (at 180 s; n = 9) of the baseline value, indicating no change in E_m. On the other hand, 14 glomus cells showed a significant decrease in fluorescence, representing 5.6 ± 1.2% (at 180 s; P < 0.05, n = 8) of the control, suggesting membrane hyperpolarization. The remaining eight cells were activated by hypoxia and showed a 6.4 ± 1.05% (at 180 s; P < 0.05, n = 7) increase in fluorescence compared with control, indicating membrane depolarization. Ten glomus cells for each level of [K⁺]_e were used for calibration of the bis-oxonol fluorescence. Figure 1C shows that, with increasing [K⁺]_e from 4.7 mM to 10, 15, and 25 mM, the fluorescence increased linearly, consistent with the expected depolarization of glomus cell E_m. The abscissa represents [K⁺]_e and the corresponding calculated glomus cell E_m using the Mullis-Noda modified constant field equation (12), considering the average resting E_m value for a glomus cell as 55 mV at 4.7 mM [K⁺]_e (46). A 20.3 mM increase (from 4.7 to 25 mM) in [K⁺]_e resulted in an overall 56.6 ± 7% (P < 0.05) increase in fluorescence, indicating an ~28-mV depolarization (from 55 to 27 mV), i.e., an average 2% increase in bis-oxonol fluorescence per millivolt change in glomus cell E_m. This change in bis-oxonol fluorescence is in good agreement with previous published reports on bovine pulmonary arterial endothelial cells (1). The buffer without the cells did not alter the level of bis-oxonol fluorescence.

View larger version (20K): [in this window] [in a new window]   Fig. 1.   Bis-oxonol fluorescence responses of rat glomus cells (isolated and clustered) during hypoxia and high K+. Time (in seconds) for normoxic period (negative numbers) and hypoxic/high-K+ period (positive numbers), which starts at time 0, is shown in each panel. Normoxic images were acquired for 50 s (1 frame/10 s) and hypoxia/high K+ images for 180 s (1 frame/10 s). A: basal fluorescence of glomus cells during normoxia (~130 Torr PO2; top); no detectable change in fluorescence in glomus cells exposed to acute hypoxia (~10 Torr PO2), indicating that plasma membrane potential (Em) was unchanged (middle), and increase in fluorescence (red color) in the same cells exposed to high extracellular K+ concentration ([K+]e, 25 mM) during normoxia, indicating Em depolarization (bottom). Magnification ×600; scale bar, 10 µm. B: 1 glomus cell showing cytosolic distribution of fluorescence (excluding the nucleus) during normoxia (~125 Torr PO2; top) and an increase in fluorescence produced by the same level of hypoxia, indicating depolarization (bottom). Scale bar, 6 µm. C: time course of percent fluorescent changes [arbitrary units (au)] during hypoxia with respect to control (top) and average fluorescent intensity of 30 glomus cells (10 cells for each increase in [K+]e) for calibration of bis-oxonol fluorescence with high K+ (bottom). * Significant difference.

View larger version (26K): [in this window] [in a new window]   Fig. 2.   Bis-oxonol fluorescence in rat glomus cell during high PCO (500 Torr). Three frames are shown from images that were captured for 50 s (1 frame/10 s) during normoxia and 180 s (1 frame/10 s) during high PCO. Time (see Fig. 1 legend) is shown in each panel, with start of high PCO at time 0. A: basal fluorescence response with normoxia (~128 Torr PO2; top), effect of high PCO showing decrease in fluorescence (middle), and images acquired 2 min after return to normoxia to show restoration of basal fluorescence (~90%; bottom). Magnification ×600; scale bar, 7 µm. B: time course of percent change in fluorescence from 10 glomus cells treated with high PCO. * Significant difference.

High PCO and glomus cell I_K. To confirm that the high-PCO-induced E_m hyperpolarization was due to I_K activation, we measured the whole cell I_K of glomus cells. In the absence of high PCO during normoxia (Fig. 3A; ~130 Torr PO₂), the whole cell current was dominated by outward I_K, which were clearly distinguished during steps positive to 20 mV (85-mV holding potential and voltage ramps from 80 to +60 mV in 10-mV increments). Bath perfusion with 500 Torr PCO (~130 Torr PO₂) resulted in activation of I_K, indicating increased K⁺ permeability (Fig. 3B). Figure 3C shows the current-voltage relationship during normoxia and high PCO. Overall, the average current increased significantly from 262 ± 119 (control) to 445 ± 169 pA (high PCO) at +50 mV (n = 7, P < 0.05). A shift in the reversal potential from 31.0 ± 6.15 to 42.57 ± 6.96 mV (Fig. 3D; P < 0.01, n = 7) indicates membrane hyperpolarization.

View larger version (22K): [in this window] [in a new window]   Fig. 3.   Representative recordings of high PCO (~500 Torr) effect on whole outward K+ currents from a rat glomus cell. Membrane was held at 85 mV, with voltage steps from 80 to +60 mV in 10-mV increments. All recordings were from the same patch. A: whole cell outward K+ currents in the absence of high PCO during normoxia (~130 Torr PO2). B: increase in outward currents in the presence of high PCO. C: current-voltage relationship from A and B. D: summary of high-PCO-induced shift in reversal potential. Data were obtained from 7 glomus cells. **P < 0.01 vs. control.

Monitoring PC-12 cell E_m with bis-oxonol. To determine whether the heterogeneity in the glomus cell E_m responses was due to reasons other than an experimental artifact, we conducted a similar study of E_m change of PC-12 cells in response to acute hypoxia, because PC-12 and glomus cells are considered to be of the same embryonic origin and to be O₂ sensitive (47). Figure 4A shows basal fluorescent images from a PC-12 cell captured during normoxic exposure (~125 Torr PO₂). Changing the perfusion to hypoxia (~10 Torr PO₂) resulted in an increase in fluorescence levels as shown at 30 and 120 s (Fig. 4) and then stabilization (not shown). The hypoxia-induced increase in fluorescence represented 8.7 ± 1.6% (P < 0.05, n = 8) of the basal fluorescence and was quite reproducible (Fig. 4B), indicating membrane depolarization and confirming the results of an earlier patch-clamp report (47).

View larger version (46K): [in this window] [in a new window]   Fig. 4.   PC-12 cell Em responses to hypoxia as measured by bis-oxonol fluorescence. Images were acquired for 50 s during normoxia and for 180 s during hypoxia (1 frame/10 s). Time (see Fig. 1 legend) is shown in each panel, with start of hypoxia at time 0. A: bis-oxonol fluorescence with normoxia (~125 Torr PO2; top) and effect of hypoxia showing increase in bis-oxonol fluorescence, indicating depolarization of Em in PC-12 cell (bottom). B: time course of percent change in bis-oxonol fluorescence obtained from 12 PC-12 cells during hypoxia. Magnification ×600; scale bar, 6 µm. * Significant difference.

JC-1 fluorescence responses of glomus cell _m with FCCP, high PCO, and hypoxia. Ten CBs from five rats were used for the glomus cell _m study. The initial study was carried out to confirm that the fluorescent probe JC-1 functioned in a manner consistent with that expected for a mitochondrial voltage-sensitive dye. JC-1 is taken up selectively by mitochondria, and the uptake is dependent on the mitochondrial potential (40). This was evident from the intense red J-aggregate formations of the JC-1 fluorescence (excluding the nucleus) in the glomus cell due to high _m during normoxia (~125 Torr PO₂; Fig. 5A). In the presence of FCCP, a protonophore uncoupler that abolishes the electrochemical gradient, there was a rapid disappearance of J-aggregates (red spots) with very little detected at 120 s (Fig. 5A).

View larger version (44K): [in this window] [in a new window]   Fig. 5.   JC-1 fluorescence responses of rat glomus cells. A: J-aggregates in cultured rat glomus cell in absence and presence of carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP). Images were acquired for 50 s during normoxia and 120 s with FCCP (1 frame/10 s). Time (see Fig. 1 legend) is shown in each panel, with start of FCCP at time 0. In normoxia (~125 Torr PO2), glomus cell mitochondria in red, yellow, and green pseudocolors represent different concentrations of J-aggregates, which indirectly suggest variation in mitochondrial membrane potential (m; top); there is no uptake of JC-1 fluorescence in the nucleus. In the presence of 1 µM FCCP, nearly all mitochondria depolarized, as indicated by disappearance of J-aggregates, because of loss of electrochemical gradient (bottom). Magnification ×1,000; scale bar, 7 µm. B: effect of high PCO (500 Torr) on J-aggregate formations in rat glomus cell mitochondria. Images were acquired every 10 s for 50 s during normoxia and 180 s for high PCO. Time (see Fig. 1 legend) is shown in each panel, with start of high PCO at time 0. Top: J-aggregate formations of JC-1 fluorescence under normoxia (~130 Torr PO2). Note variations of mitochondrial density within the cell and large nucleus devoid of stain. Bottom left: 3-dimensional reconstruction during normoxia clearly shows very high concentration of J-aggregates (red), indicating very high m; few yellow and green pseudocolors correspond to high and low concentrations of J-aggregates, which indirectly suggest high and low m. Middle: decrease in J-aggregates with high PCO, indicating mitochondrial depolarization. Bottom (middle and right panels): 3-dimensional images show that few deenergized mitochondria (green) at 60 s are energized (red) at 180 s (arrows). Magnification ×1,000; scale bar, 7 µm. C: J-aggregate formations in glomus cell mitochondria during normoxia and hypoxia. Images were acquired for 50 s during normoxia and for 180 s during hypoxia. Time (see Fig. 1 legend) is shown in each panel, with start of hypoxia at time 0. Top: in normoxia (~125 Torr PO2), large and small clusters of J-aggregates are visible, depending on density of mitochondria. Bottom left: 3-dimensional reconstruction shows distribution of J-aggregate formations within mitochondria, as indicated by red, yellow, and green pseudocolors. Middle: effect of hypoxia (~10 Torr PO2) on glomus cell m. Disappearance of J-aggregates is not evident in all mitochondria (at 120 s). Bottom middle: major reorientation of mitochondria during hypoxia compared with normoxia (arrow). Bottom right: mitochondrial potential regained from one time frame (60 s) to another (120 s; arrow). Magnification ×1,000; scale bar, 6 µm. D: time course of percent change in JC-1 fluorescence during FCCP (10 cells), high PCO (15 cells), and hypoxia (30 cells). Arrow, switch to FCCP, PCO, and hypoxia. *P < 0.05.

View larger version (26K): [in this window] [in a new window]   Fig. 6.   Fluorescence localization of J-aggregates of JC-1 in mitochondria of a rat PC-12 cell. A: energized mitochondria exhibiting high m during normoxia (~130 Torr PO2). B and C: hypoxia (~10 Torr PO2) and FCCP (1 µM) showing loss of J-aggregates, indicative of mitochondrial depolarization. D: recovery of J-aggregates with normoxia after 2 min. E: time course of average change in JC-1 fluorescence in PC-12 cells during hypoxia (20 cells) and FCCP (10 cells). Images were acquired every 20 s for 120 s. Magnification ×600; scale bar, 6 µm. Arrow, switch to FCCP and hypoxia.

View this table: [in this window] [in a new window]   Table 1.   Summary of the Em and m responses

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

In the present study, we have reported for the first time the use of voltage-sensitive fluorescent dyes, bis-oxonol and JC-1, for indirect assessments of glomus cell E_m and _m, respectively. Glomus cell membrane depolarization was confirmed by an increase in bis-oxonol fluorescence using high [K⁺]_e. Glomus cell membrane hyperpolarization was demonstrated by high PCO, which decreased the fluorescence level. Loss of glomus cell mitochondrial J-aggregates by the protonophore uncoupler FCCP confirmed mitochondrial depolarization and also suggested that J-aggregate formations are dependent on the mitochondrial electrochemical gradient, as reported by others (40). As expected, high PCO and hypoxia also resulted in loss of J-aggregates of JC-1 fluorescence, indicative of mitochondrial depolarization. Thus bis-oxonol and JC-1 have proven to be effective in expressing the E_m and _m changes in the glomus cells in response to the above stimuli as expected. However, fluorescence responses of glomus cell E_m to hypoxia were not uniform.

According to the plasma membrane hypothesis, hypoxia must depolarize glomus cell E_m, as borne out in some patch-clamp recordings (11, 27, 37, 46). However, there are indications that glomus cell depolarization may not be a prerequisite for an increase in [Ca²⁺]_i, neurotransmitter release, and neural discharge. Cyanide is reported to hyperpolarize rabbit glomus cells (5), yet it increases glomus cell Ca²⁺ (16) and always stimulates neural discharge (33). Furthermore, E_m recorded in isolated rat glomus cells using sharp electrode impalement showed depolarization in 29% and hyperpolarization in 64% of the cells tested with hypoxia (36). Interestingly, glomus cells are also reported to be unresponsive to hypoxia-induced catecholamine secretion and increase in [Ca²⁺]_i (7, 39). A recent report in hippocampal neurons has suggested that, even at hyperpolarized E_m, an increase in [Ca²⁺]_i could be achieved (34). Our imaging data showed that glomus cells that depolarized with high K⁺ concentration were insensitive (45%), hyperpolarized (35%), or depolarized (20%) in response to acute hypoxia and did not comply with the uniform depolarization reported in patch-clamp studies (11, 27, 37, 46). Moreover, the bis-oxonol response with high PCO showing apparent hyperpolarization of glomus cell E_m indicates that the increase in [Ca²⁺]_i and neural excitation due to high PCO (25) may not be due to membrane depolarization.

We cannot reject some methodological issues that may be related to unresponsiveness/hyperpolarization of glomus cells in response to hypoxia. Cellular damage due to dissociation could be a reason, because PC-12 cells, which were not dissociated, did not display variation of the E_m response with hypoxia. Moreover, because subpopulations of glomus cells are identified on the basis of size and synapses to nerve endings (28), many glomus cells may not respond or may respond differently to hypoxia. Isolated glomus cells most commonly hyperpolarize because of the lack of type II cells (36), which resemble morphologically the glial cells and can exert significant regulatory effects on excitable cells (9). Finally, it is possible that the magnitude of hypoxia (~10 Torr PO₂) used in the present study was not severe enough to affect the O₂-sensing molecule in all the cells. Previous studies correlating the PO₂ with the [Ca²⁺]_i response (11) and catecholamine release (38) indicate that 3 Torr PO₂ is necessary to affect most of the glomus cells. Nevertheless, our result showed a tendency for E_m depolarization of the glomus cell (at 10 Torr PO₂), but the response was not consistent, because the hypoxia PO₂ was not severe enough.

Glomus cell mitochondrial depolarization was qualitatively assessed by the loss of J-aggregates (not J-monomers) of the JC-1 fluorescence, which was imaged under green excitation, and the concentrations of the aggregates were expressed in pseudocolors. Accordingly, we detected a very high concentration of J-aggregates (red pseudocolor) in the glomus cell mitochondria during normoxia, indicating very high resting _m (Fig. 5, A-C) Also, less aggregation of JC-1 fluorescence in the mitochondria was evident because of variation in resting _m (yellow and green pseudocolors). A previous study using rhodamine 123 showed mitochondrial depolarization with hypoxia but was unable to demonstrate heterogeneity of _m within a glomus cell (16). It is quite likely that all glomus cell mitochondria may not adopt the same potential or that there may be regional heterogeneity in _m due to 1) the localized proton circuit of mitochondria leading to an uneven distribution of membrane potential across inner membrane, 2) unequal rate of respiration and/or ATP synthesis, and 3) uneven distribution of Ca²⁺ along the surface of the mitochondria. Although we did not study the effect of anoxia, the results demonstrated that _m of glomus cells were sensitive to a hypoxic PO₂ of 10 Torr. Using a photometric technique, Duchen and Biscoe (16) found graded depolarization of _m with graded changes in PO₂ (between 40 and 0 Torr) and proposed a mitochondrial population sensitive to hypoxia, while all mitochondria would respond to anoxia. In the present study, a 60% decrease in JC-1 fluorescence (or disappearance of J-aggregates) at the end of hypoxic exposure (Fig. 5D) indicates that certain mitochondria within the glomus cells may remain in the energized state and need not depolarize in response to hypoxia (~10 Torr PO₂). This could be a mere biological variation as manifested by the mitochondria. A greater decrease in JC-1 fluorescence (~90%) during high PCO could be due to a stimulus that is greater than hypoxia (~10 Torr PO₂). Our imaging data showing glomus cell mitochondrial depolarization with hypoxia and high PCO are not enough to support the notion of different populations of mitochondria in the glomus cell.

Perspective

We cannot predict that mitochondria in glomus cells are unique. However, the imaging technique has the potential to address this issue if one could show in comparative studies that CB mitochondria behave differently. For this reason, additional studies are needed to examine whether mitochondria from other nonexcitable cells, e.g., erythropoietin-secreting cells of the kidney, depolarize to high PCO and low O₂. It is worth mentioning that in the CB the [Ca²⁺]_i (43), dopamine (32), and CSN (22) response curves against decreasing PO₂ are shifted to the right compared with the leftward shift of, for example, the hypoxia-inducible factor-1 response in HeLa cells (20). A recent study indicates that oxidative phosphorylation in liver mitochondria is more efficient in hypoxia than in normoxia (18).

If the glomus cells do not depolarize, then how does intracellular Ca²⁺ rise? Despite E_m hyperpolarization (as shown here), CO can readily diffuse into glomus cells and possibly stimulates Ca²⁺ release from the intracellular stores (31). Duchen and Biscoe (16) proposed that Ca²⁺ might simply follow _m, with the release of Ca²⁺ from the internal stores. Similarly, Rizzuto et al. (42) reported close apposition between mitochondria and endoplasmic reticulum membranes and depolarization of the mitochondria likely to release Ca²⁺ from the endoplasmic reticulum. Alternatively, Nowicky and Duchen (34) suggested that, with altered mitochondrial metabolism during hypoxia, there might be a shift in the activation curve of Ca²⁺ channels in the hyperpolarizing direction, so that significant numbers of channels open at resting E_m and could provide increased influx of Ca²⁺. Our explanation is that, with _m depolarization, there could be intracellular release of Ca²⁺ from the stores, but it may not be sufficient to increase neurotransmitter release and CSN discharge, in agreement with the capacitative Ca²⁺ entry (4). In the sequence of capacitative Ca²⁺ entry, the stores are continuously emptied by inositol trisphosphate-stimulated release of Ca²⁺ and, in the process, are replenished by influx of Ca²⁺ from an extracellular source. If that is the case, then glomus cell E_m has to depolarize, and, at some point downstream in the chemotransduction pathway, influx of Ca²⁺ must be necessary for neural discharge (44).

Overall, if the glomus cell has to be depolarized with a contribution from mitochondria, then the metabolic and membrane hypothesis cannot exist independently. It has been reported that the onset of glomus cell mitochondrial depolarization precedes cell membrane depolarization, indicating a very rapid means of communication between mitochondria and cell membrane (10). Therefore, we name our hypothesis the metabomembrane hypothesis. The summary for the steps in the hypoxia chemotransduction model could be as follows. Hypoxia, which affects oxidative phosphorylation, causes _m depolarization and Ca²⁺ release from the stores, followed by E_m depolarization and influx of Ca²⁺. This would lead to a rapid rise in cytoplasmic Ca²⁺ concentration, thus stimulating neurosecretion and, finally, sensory excitation. The fact that an increase in the CB sensory response is not always accounted for by the cellular response (43) raises the possibility that the nerve terminals are also important at some point later in the chemotransduction pathway, considering that the glomus cells are important in initiating the response.

We conclude that the voltage-sensitive fluorescent probe bis-oxonol apparently can determine the glomus cell E_m depolarization in response to high K⁺ and hyperpolarization induced by high CO, but we are unable to confirm depolarization as the critical event during hypoxia. Nevertheless, the depolarizing trend was there, but because hypoxia PO₂ was not severe enough, it was not consistent. The apparent _m determined by the probe JC-1 confirmed depolarization induced by hypoxia and high CO. So, in relative terms, glomus cell _m depolarization could be a reliable phenomenon compared with E_m and could support the mitochondrial hypothesis of hypoxic chemoreception in the CB. An appropriate name could be the metabomembrane hypothesis.