Intracellular pH regulates superoxide production by the macula densa

Abstract

We hypothesized that elevated macula densa intracellular pH (pH_i) during tubuloglomerular feedback enhances O₂⁻ production from NAD(P)H oxidase. Microdissected thick ascending limbs from rabbits with intact macula densa were cannulated and perfused with physiological saline. When luminal NaCl was switched from 10 to 80 mM, O₂⁻ production increased from 0.53 ± 0.09 to 2.62 ± 0.54 U/min (P < 0.01). To determine whether inhibiting the Na/H exchanger blocks O₂⁻ production, we used dimethyl amiloride (DMA) to block Na/H exchange. In the presence of DMA, O₂⁻ production induced by NaCl was blunted by 40%. To study the effect of pH_i on O₂⁻ in intact macula densa cells, we measured O₂⁻ while pH_i was changed by adjusting luminal pH. When the macula densa was perfused with 80 mM NaCl and the pH of the perfusate was switched to 6.8, 7.4, and 8.0, O₂⁻ production was significantly enhanced, but not at 10 mM NaCl. To ascertain the source of O₂⁻, we used the NAD(P)H oxidase inhibitor apocynin. In the presence of apocynin (10⁻⁵ M), O₂⁻ production induced by elevating pH_i was blocked. Finally, we measured the optimum pH for O₂⁻ production by the macula densa and found optimum extracellular pH is at 7.7 and optimum pH_i is ∼8 for O₂⁻ production. We found that elevated pH_i enhances O₂⁻ production from NAD(P)H oxidase induced by increasing luminal NaCl when the lumen is perfused with 80 mM NaCl, not 10 mM, and O₂⁻ production is pH sensitive, with an optimum pH_i of 8.

tubuloglomerular feedback (TGF) is a process in which increases in luminal NaCl at the macula densa cause constriction of the afferent arteriole (Af-Art). TGF limits urinary volume and Na⁺ excretion and decreases the glomerular filtration rate by increasing Af-Art tone in response to increased NaCl at the macula densa (25, 33, 40). TGF is inhibited by a number of factors, including atrial natriuretic factor and nitric oxide (10, 15). In contrast, angiotensin and superoxide (O₂⁻) stimulate TGF (30, 35). O₂⁻ production in the macula densa stimulates TGF primarily by scavenging nitric oxide (18, 35). The source of O₂⁻ in the macula densa is primarily NAD(P)H oxidase (16).

O₂⁻ production by the macula densa is stimulated by increasing luminal NaCl, the same stimulus that initiates TGF. When luminal NaCl is elevated, the activity of luminal membrane Na-K-2Cl cotransporters is enhanced, leading to depolarization of the macula densa. This appears to be the signal that activates enhances O₂⁻ production (16). In addition to causing depolarization, elevating luminal NaCl at the macula densa also causes intracellular pH (pH_i) to increase due to activation of luminal Na/H exchanger (NHE) (15, 27). However, the effect of this change in pH_i and the role of luminal NHE on O₂⁻ production has not been studied.

NAD(P)H oxidase produces protons in addition to O₂⁻. NADPH oxidase has also been reported to be a proton pump (1, 22). The activity of the enzyme is enhanced by alkaline pHs (34, 37). It is unclear whether this is due to removal of end-product inhibition or a direct effect of pH on enzyme structure and function. We hypothesized that the increase in pH_i caused by elevating luminal NaCl, and consequent activation of NHE, enhances O₂⁻ production by NAD(P)H oxidase in the macula densa.

MATERIALS AND METHODS

Isolation and microperfusion of rabbit macula densa.

We used methods similar to those described previously to isolate and microperfuse the thick ascending limb (TAL) of the loop of Henle and attached macula densa (15). This protocol was approved by the Institutional Animal Care and Use Committee at Henry Ford Hospital. Briefly, young male New Zealand White rabbits (1.5–2.0 kg) were anesthetized with ketamine (50 mg/kg im) and given an injection of heparin (500 U iv). The kidneys were removed and sliced along the corticomedullary axis. Single superficial intact glomeruli were microdissected together with adherent tubular segments consisting of portions of the TAL, macula densa, and early distal tubule. Using a micropipette, samples were transferred to a temperature-regulated chamber mounted on an inverted microscope (TE2000-S, Nikon, Yuko, Japan). The glomeruli were positioned so the macula densa could be visualized using a holding pipette. TALs were cannulated with an array of glass pipettes (14, 15). Macula densas were perfused with physiological saline containing (in mM) 10 HEPES, 1.0 CaCO₃, 0.5 K₂HPO₄, 4.0 KHCO₃, 1.2 MgSO₄, 5.5 glucose, 0.5 Na acetate, 0.5 Na lactate, and 10 or 80 NaCl, pH 7.4. The luminal perfusion rate was 40 nl/min and controlled with a pump (11 plus, Harvard Apparatus, Holliston, MS). The bath consisted of MEM containing 0.15% bovine serum albumin and was exchanged continuously at a rate of 1 ml/min. Microdissection and cannulation were completed within 60 min at 8°C, after which the bath was gradually warmed to 37°C for the rest of the experiment. Once the temperature was stable, a 30-min equilibration period was allowed before any measurements were taken. The imaging system consisted of a microscope (TE2000-S, Nikon), digital CCD camera (IEEE 1394, Hamatsu Photonics, Hamatsu, Japan), and optical filter changer (Lambda 10–2, Sutter Instruments, Novato, CA). Images were displayed and analyzed with SimplePCI imaging software (Compix, Tualatin, OR).

Measurement of O₂⁻.

As we described (16), an O₂⁻-sensitive fluorescent dye, dihydroethidium, was used to detect O₂⁻ production by the macula densa. The cells were loaded with 10 μM dihydroethidium in 0.1% DMSO plus 0.1% pluronic acid from the lumen for 30 min and then washed for 20 min. Dihydroethidium is irreversibly converted to oxyethidium in the presence of O₂⁻ (38, 39, 42). Therefore, the rate of oxyethidium accumulation reflects O₂⁻ production. An increase or decrease in this rate indicates enhanced or blunted O₂⁻ production. O₂⁻ production is expressed as the rate of increase in the ratio of oxyethidium to dihydroethidium fluorescence intensity (units/min). Macula densa cells were exposed to 380- and 490-nm light to excite dihydroethidium and oxyethidium, respectively. Emitted fluorescence from dihydroethidium was recorded using a 420-nm dichroic mirror with a 460/50-nm band-pass filter; for oxyethidium, we used a 565-nm dichroic mirror with a 605/55-nm band-pass filter (Chroma). Square-shaped regions of interest were set inside the cytoplasm of macula densa cells, and mean intensity within them was recorded every 5 s. Changes in O₂⁻ concentration were expressed as the increase in oxyethidium/dihydroethidium ratio (units/s).

All studies were performed in the presence of the nitric oxide synthase (NOS) inhibitor N-nitro-l-arginine methyl ester (l-NAME; 10⁻⁴ M) while O₂⁻ was measured to eliminate its reaction with nitric oxide.

Calibration and measurement of pH_i.

Once TALs were perfused, macula densa cells were loaded with BCECF-acetoxymethyl ester (AM) from the lumen at 37 ± 1°C for 15 min and then washed for 10 min. Intracellular dye was excited alternately at 440 and 490 nm, and emitted fluorescence was recorded at 540 nm. Ratiometric measurements of the images were analyzed with the imaging system. The ratio was calibrated as described previously (7, 15). The pH calibration solutions contained (in mM) 95 KCl, 5 NaCl, 30 N-methyl-d-glucamine, 2.5 NaH₂PO₄, 1.5 MgSO₄, 5 glucose, 2 CaCl₂, 25 HEPES, and 0.01 nigericin and were titrated to selected pH values between 6.4 and 8.0. Nigericin was prepared as a 10 mM stock in methanol and diluted in calibration solution just before use. All solutions had an osmolality of 290 mosmol/kgH₂O adjusted with mannitol.

Chemicals.

Dihydroethidium, BCECF-AM, DMSO, and pluronic acid were obtained from Molecular Probes (Eugene, OR). All other chemicals were purchased from Sigma.

Statistics.

Data were collected as repeated measures over time under different conditions. We tested only the effects of interest, using a paired t-test or ANOVA for repeated measures. When ANOVA was used, the three pairwise comparisons were then evaluated using three paired t-tests. Significance was judged as P < 0.05 or an adjusted value using Hochberg's method for multiple testing.

RESULTS

First, we demonstrated that increasing luminal NaCl increases O₂⁻ production, as we have reported (16). When luminal NaCl was 10 mM, the rate of change in the OE/DHE ratio was 0.53 ± 0.09 U/min. After a switch to 80 mM NaCl, it was 2.62 ± 0.54 U/min (P < 0.01 vs. 10 mM NaCl; n = 5). To show that changes in the OE/DHE ratio are due to changes in O₂⁻, we used tempol (10⁻⁴ M), a stable membrane-permeant superoxide dismutase (SOD) mimetic. In the presence of tempol, when the lumen was perfused with 10 mM NaCl, the rate of increase in the OE/DHE ratio was 0.47 ± 0.09 U/min. When NaCl was switched to 80 mM NaCl, it was 0.58 ± 0.23 U/min (n = 5; Fig. 1). Thus increases in the OE/DHE ratio are due to increases in O₂⁻ concentration, and increasing luminal NaCl stimulates O₂⁻ production by the macula densa.

Fig. 1.

Tempol blocks the increase in superoxide induced by NaCl. When NaCl was switched from 10 to 80 mM, superoxide concentration increased significantly. In the presence of tempol, a stable membrane-permeant SOD mimetic, the NaCl-induced increase in superoxide was blocked (n = 5).

Increasing luminal NaCl at the macula densa elevates pH_i due to activation of NHE. To begin to study whether macula densa pH_i regulates O₂⁻ production and the role of NHE, we measured the effect of dimethyl amiloride (DMA), a NHE inhibitor, on O₂⁻ production and pH_i. During the control period, when luminal NaCl was switched from 10 to 80 mM, O₂⁻ production increased from 0.34 ± 0.15 to 2.56 ± 0.31 U/min (P < 0.01; n = 7; Fig. 2). When we repeated the switch in the presence of DMA (10⁻⁴ M), O₂⁻ production only increased from 0.29 ± 0.14 to 1.65 ± 0.36 U/min, 40% less (P < 0.05 vs. without DMA; n = 7; Fig. 2). Time controls showed no significant changes (O₂⁻ production increased from 0.42 ± 0.17 to 2.83 ± 0.22 U/min and from 0.47 ± 0.21 to 2.74 ± 0.35 U/min in the first and second luminal NaCl switches, respectively). When we repeated these experiments but measured pH_i instead of O₂⁻, we found that in the absence of DMA increasing luminal NaCl elevated pH_i from 7.1 ± 0.1 to 7.7 ± 0.2 (P < 0.01; n = 4; Fig. 3) and in the presence of DMA this increase was completely blocked (Fig. 3). Control experiments showed no significant effects of time on NaCl-induced changes in pH_i (pH_i increased from 7.0 ± 0.1 to 7.7 ± 0.1 and 7.1 ± 0.2 to 7.8 ± 0.2 in the first and second luminal NaCl switches, respectively). Taken together, these data suggest that the changes in macula densa pH_i induced by activating NHE enhance O₂⁻ production.

Fig. 2.

Blocking the Na/H exchanger blunts the superoxide production induced by increasing luminal NaCl. In a control experiment, increasing luminal NaCl stimulated superoxide production. When the Na/H exchanger was blocked with dimethylamiloride, NaCl-induced reduction in superoxide production was inhibited by 40% (n = 7).

Fig. 3.

Blocking the Na/H exchanger inhibits the increase in intracellular pH (pH_i) induced by increasing luminal NaCl. When macula densa NaCl was switched from 10 to 80 mM, pH_i increased from 7.1 ± 0.1 to 7.7 ± 0.2 (P < 0.01; n = 4). In the presence of dimethylamiloride, a Na/H exchanger inhibitor, NaCl-induced changes in pH were blocked.

The inhibition of NaCl-induced O₂⁻ production caused by DMA could be either due to blocking changes in pH_i or intracellular Na, because NHE transports both protons and Na (19). To directly study whether the effect of DMA is due to changes in macula densa pH_i, we clamped pH_i with a K⁺/H⁺ ionophore, nigericin, to equilibrate luminal and pH_i without switching luminal NaCl. The macula densa was perfused with 80 mM NaCl, and nigericin (10⁻⁵ M) was added to the perfusate for 20 min. When pH_i was adjusted to 7.0, O₂⁻ production was 1.54 ± 0.24 U/min. When we raised pH_i to 7.8, O₂⁻ production by the macula densa increased to 2.11 ± 0.28 U/min (P < 0.01; n = 5; Fig. 4). These data indicate that elevated macula densa pH_i enhances O₂⁻ production in 80 mM NaCl rather than changes in intracellular Na.

Fig. 4.

Effect of pH_i on superoxide production by the macular densa. pH_i was clamped with a K⁺/H⁺ ionophore, nigericin, to equilibrate luminal pH and pH_i. When pH_i was raised from 7.0 to 7.8 without a change in NaCl concentration, superoxide production by the macula densa was increased significantly (P < 0.01; n = 5).

Clamping pH_i with nigericin has limitations and potential unintended consequences. Thus we next tested whether we could effectively set pH_i by changing luminal pH in intact macula densa cells in both 10 and 80 mM NaCl. When the macula densa was perfused with 10 mM NaCl and luminal pH was raised from 6.8 to 7.4, 8.0, and 8.6, pH_i was 6.69 ± 0.24, 7.20 ± 0.16, 7.82 ± 0.17, and 8.23 ± 0.19, respectively. When the experiment was repeated with 80 mM NaCl, pH_i increased from 7.13 ± 0.26 to 7.57 ± 0.22, 8.16 ± 0.26, and 8.65 ± 0.28, respectively (Fig. 5; n = 6). These data indicate that changing luminal pH can be used as a mechanism to alter pH_i and to further study the role of pH in regulating O₂⁻ production.

Fig. 5.

Effect of changes in luminal pH on macula densa pH_i. When luminal pH was increased from 6.8 to 8.6, there was a linear correlation between changes in pH_i and extracellular pH of the macula densa. At any given extracellular pH, pH_i of the macula densa at 80 mM NaCl was 0.5, higher than at 10 mM (n = 6).

Our data with DMA indicate that pH may have less of an effect on O₂⁻ production when the macula densa is perfused with 10 mM than 80 mM NaCl. To examine this issue in more detail, we measured O₂⁻ production while the luminal perfusate, and thus pH_i, was adjusted to different pHs, in the presence of either 10 or 80 mM NaCl. When the macula densa was perfused with 10 mM NaCl and the pH of the perfusate was switched from 6.8 to 7.4, and 8.0, there was no significant increase in O₂⁻ (0.37 ± 0.16, 0.44 ± 0.09, and 0.57 ± 0.20 U/min, respectively) (Fig. 6; n = 5). In contrast, when the macula densa was perfused with 80 mM NaCl and the pH of the perfusate was switched from 6.8 to 7.4 and 8.0, O₂⁻ production was increased as pH increased (0.69 ± 0.16, 1.81 ± 0.32, and 2.27 ± 0.32 U/min, respectively) (P < 0.01 vs. pH 6.8; Fig. 6; n = 5). These data suggest that increasing macula densa pH_i enhances O₂⁻ generation only when the lumen is perfused with 80 mM NaCl, not 10 mM. To study whether the effect of pH was on NAD(P)H oxidase, we used the NAD(P)H oxidase inhibitor apocynin. In the presence of apocynin when the pH of the perfusate was switched from 6.8 to 7.4 and 8.0 with 80 mM NaCl in the lumen, O₂⁻ production it was 0.33 ± 0.15, 0.39 ± 0.08, and 0.54 ± 0.19 U/min, respectively (Fig. 6: n = 5). These data suggest that NAD(P)H oxidase is the primary source of O₂⁻.

Fig. 6.

Effect of luminal pH on superoxide production. When the macula densa was perfused with 10 mM NaCl and luminal pH was switched to 6.8, 7.4, and 8.0, there was no significant increase in superoxide production. When the macula densa was perfused with 80 mM NaCl and luminal pH was switched to 6.8, 7.4, and 8.0, superoxide production was significantly enhanced. In the presence of apocynin (10⁻⁵ M), NaCl-induced superoxide production was blocked (n = 5; *P < 0.01 vs. pH 6.8).

Finally, we measured the optimum pH for O₂⁻ production by the macula densa. When the pH of the perfusate was switched from 6.8 to 7.1, 7.4, 7.7, 8.0, 8.3, and 8.6 with 80 mM NaCl in the lumen, O₂⁻ was 0.69 ± 0.16, 0.89 ± 0.21, 1.81 ± 0.32; 2.46 ± 0.43, 2.26 ± 0.31, 1.61 ± 0.37, and 1.24 ± 0.29 U/min, respectively (Fig. 7; n = 5). These data suggest that the optimum extracellular pH is at 7.7, corresponding to a pH_i of ∼8, for O₂⁻ production.

Fig. 7.

Optimum pH for superoxide production at the macula densa. The macula densa was perfused with 80 mM NaCl, and luminal pH was switched from 6.8 to 8.6. Superoxide production was inhibited significantly at both low and high pH values, with a optimum luminal pH at 7.7 (pH_i ≈ 8.); n = 5.

DISCUSSION

When the luminal NaCl concentration increases at the macula densa, the cells depolarize (2) and pH_i increases due to activation of NHE (15, 27). We reported that depolarization activates NAD(P)H oxidase and induces O₂⁻ production (16). However, the role of pH_i and NHE in O₂⁻ production is unknown. We hypothesized that the elevation in macula densa pH_i caused by increasing luminal NaCl, and consequent activation of NHE, enhances O₂⁻ production from NAD(P)H oxidase.

To begin to test our hypothesis, we first studied the effect of inhibiting luminal NHE on the increase in O₂⁻ production caused by raising luminal NaCl at the macula densa. Luminal NHEs are important regulators of pH_i in the macula densa (5, 15, 26, 27), and an increase in luminal NaCl stimulates their activity (5). We found that the NHE-selective inhibitor DMA reduced the increase in O₂⁻ production caused by raising luminal NaCl from 10 to 80 mM by 40%. These data directly indicate that activation of luminal NHE that occurs when luminal NaCl is elevated stimulates O₂⁻ production. However, they do not indicate whether the increase in pH_i or the increase in intracellular Na resulting from NHE activation stimulates O₂⁻ production.

Our results showing the inhibition of NHE in the macula densa are similar to results found in other nephron segments. In the thick ascending limb, blocking NHE with methylisobutyl-amiloride or dimethylamiloride reduced O₂⁻ production (13, 21). However, unlike the macula densa, inhibiting NHE in thick limbs completely prevented the stimulation of O₂⁻ production caused by pH, high glucose, or NaCl (13, 21).

The Na/H exchanger not only regulates pH_i but also contributes to intracellular Na⁺. When the Na/H exchanger is inhibited by DMA, intracellular Na⁺ concentration is decreased and pH_i elevation is blocked (12, 15). To test whether the effect of DMA on O₂⁻ production is due to pH_i or Na⁺ inhibition, we used a K⁺/H⁺ ionophore, nigericin, to equilibrate luminal and pH_i and measured O₂⁻ production. We found that when we increased pH_i from 7.0 to 7.8 with nigericin while keeping luminal NaCl at 80 mM, there was a significant increase in O₂⁻ production. These data suggest that 1) the effects of DMA are due to its ability to prevent changes in pH_i rather than a reduction in intracellular Na; and 2) elevated macula densa pH_i enhances O₂⁻ production.

Our earlier findings indicated that depolarization of the macula densa activates NAD(P)H oxidase and stimulates O₂⁻ production (16). Nigericin may affect membrane potential, which in turn might modulate O₂⁻ production by the macula densa. To make sure that pH_i influences O₂⁻ production in the macula densa, we sought the least invasive method possible to alter pH_i. Thus we studied the effect changing pH_i by adjusting luminal pH. We found that increasing luminal pH proportionately increased pH_i. Similar to our results, Fowler et al. (5) reported that a significant elevation of pH_i at the macula densa was induced by increasing luminal NaCl. Similar findings have also been reported for other cell types (8, 37). This effect is likely mediated by altering NHE activity. Because we had already shown that changes in intracellular Na that may accompany changes in NHE activity do not affect O₂⁻ production, increasing luminal pH was chosen as the method to study the effect of pH_i on O₂⁻ production in intact macula densa cells. We found that when the macula densa was perfused with 10 mM NaCl, increasing extracellular pH did not enhance O₂⁻ production. However, at 80 mM NaCl, increasing extracellular pH significantly enhanced O₂⁻ production. The former data seem to indicate that pH_i does not modify O₂⁻ production, while the latter data seem to indicate that it does. Our explanation for these results is that pH_i only modulates O₂⁻ production only when the macula densa is depolarized and NAD(P)H oxidase activated, as occurs when 80 mM NaCl is in the lumen. Under these circumstances, the macula densa is depolarized and depolarization activates O₂⁻ production as we have previously reported (16). In contrast, when 10 mM NaCl is in the lumen, the macula densa is hyperpolarized and O₂⁻ production is not activated. Thus pH_i has no effect. Our data indicate that pH_i is an important modulator for O₂⁻ production induced by NaCl, accounting for ∼30% of O₂⁻. We believe that depolarization of the macula densa is a major factor in control O₂⁻ production as we reported previously (16).

The above data imply that NAD(P)H oxidase is the source of O₂⁻ because we have shown that depolarization stimulates O₂⁻ production by this enzyme. To study whether this was in fact the case, we tested the ability apocynin to block the effects of pH_i on O₂⁻ production in the presence of 80 mM NaCl. Similar to our previous report (16), we found that apocynin blocked the stimulation of O₂⁻ production caused by increasing pH_i. These data suggest that NAD(P)H oxidase is the primary source of O₂⁻ induced by NaCl and affected by pH_i.

Our findings that O₂⁻ production is pH dependent only after NAD(P)H oxidase is activated are in agreement with published reports. After NAD(P)H oxidase is activated by agonists to induce O₂⁻ production, elevating pH_i pharmacologically with nigericin (11, 24), raising the external sodium concentration (34), or increasing extracellular pH (8, 23, 37) raises O₂⁻ concentrations in macrophages, leukocyte, neutrophils, and eosinophils.

To further understand the pH dependence of O₂⁻ production in the macula densa, we varied pH_i over a greater range. We found that O₂⁻ production first increased as pH_i was raised, reaching a maximum ∼pH 8.0, and then declined. Thus the correlation of NAD(P)H oxidase with pH was not a linear response. A number of studies have reported similar effects of pH on NAD(P)H oxidase activity and shown profound inhibition by either a low or high pH, with the optimum of ∼6.0–7.5 (20, 36). In phorbol myristate acetate-activated normal human neutrophils, a single enzymatic entity that was dormant in unstimulated cells, optimally active at pH 7.0, above or below the optimal pH significantly inhibits O₂⁻ production (36). Morgan et al. (20) studied the pH dependence of NAD(P)H oxidase by patch-clamping human eosinophils. They found that in intact eosinophils, NAD(P)H oxidase is highly sensitive to pH and has an optimum ∼7.5, decreasing drastically at higher or lower values. However, some reports found a linear response between pH and O₂⁻ production in zymosan-treated neutrophils and in chemotactic factor-stimulated human neutrophils. In each instance, the amount of O₂⁻ release correlated directly with pH_i and was enhanced by intracellular alkalinization from 7.0 to 8.1 (6, 34). Two other studies reported that lowering pH to 6.0 had no effect on O₂⁻ generation (28, 31). The explanation for these apparent discrepancies is unclear.

Exactly how pH_i regulates NAD(P)H oxidase is not fully understood. Since protons are generated together with O₂⁻ by NAD(P)H oxidase (1, 22), elevation pH may just relieve product induced inhibition of the enzyme. It has long been found that addition or removal of proton ions changes the charges on the protein structure of the enzyme, disrupting the ability to couple with its substrate (4). Experiments with a cell-free O₂⁻-forming system suggested that this decline in respiratory burst activity at low pH was due to inefficient activation of the O₂⁻-forming enzyme under acidic conditions (6). Bormann et al. (3) reported that activation of PLA₂ in rabbit neutrophils is pH sensitive. The optimum pH was 8.0–8.5, with activity rapidly decreasing at pH values below 7.0. PLA₂ is important for regulation of NAD(P)H oxidase activity. It controls the arachidonate-sensitive assembly of the complete oxidase complex by modulating the translocation of both p47^phox and p67^phox (32, 41). It is tempting to speculate that pH_i might be acting through this route.

We are aware of the recent publication (9) in which the authors conclude that apocynin may act as a scavenger of reactive oxygen species rather than a specific inhibitor of NAD(P)H oxidase in some cells. However, this conclusion is primarily based on Fig. 3 of the paper in which apocynin at a concentration of 1,000 μM reduces the amount of superoxide detected by either oxyethidium or lucigenin when produced by cell-free systems. Importantly, at concentrations of 100 and 10 μM, apocynin showed no scavenging activity of superoxide in these two assays. In addition, 1 mM apocynin did not have any effect on superoxide levels in HEK293 cells transfected with Nox 2 and p47 measured with lucigenin. These data clearly show that apocynin is not simply a superoxide scavenger. In our preparation we used apocynin at a concentration of 10 μM. Given that this concentration is 100 times less than that needed to scavenge superoxide, it is unlikely that apocynin is acting via this mechanism in our preparation. In conclusion, we found that elevated pH_i enhances the O₂⁻ production induced by increasing luminal NaCl. O₂⁻ production is very sensitive to pH, and optimum pH_i is ∼8. We have recently reported that increasing luminal NaCl at the macula densa activates nNOS by elevating pH_i (15, 17). In physiological situations, both nNOS and NAD(P)H oxidase activate during TGF. NO blunts tubuloglomerular feedback via the cGMP pathway (29). Superoxide at the macula densa augments tubuloglomerular feedback by scavenging nitric oxide (18), since after blocking nNOS with 7-NI, superoxide has no effect on tubuloglomerular feedback. The interaction and resulting balance between O₂⁻ and NO in the macula densa will modify the TGF response. Enhanced O₂⁻ generation and/or impaired production of NO or its metabolites (such as peroxynitrate) by the macula densa might contribute to pathophysiological changes in hypertension.

GRANTS

This work was supported by Grants SDG-0630288N from the American Heart Association (R. Liu) and RO1-HL-86767 from the National Heart, Lung, and Blood Institute (R. Liu).