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Am J Physiol Renal Physiol 293: F662-F669, 2007. First published May 16, 2007; doi:10.1152/ajprenal.00064.2007
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EDITORIAL FOCUS

ANG II provokes acute trafficking of distal tubule Na+-Cl– cotransporter to apical membrane

Monica B. Sandberg,1 Anne D. M. Riquier,1 Kaarina Pihakaski-Maunsbach,2 Alicia A. McDonough,1 and Arvid B. Maunsbach2

1Department of Physiology and Biophysics, University of Southern California Keck School of Medicine, Los Angeles, California; and 2The Water and Salt Research Center, Department of Cell Biology, Institute of Anatomy, University of Aarhus, Aarhus, Denmark

Submitted 10 February 2007 ; accepted in final form 9 May 2007


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The distal convoluted tubule (DCT) Na+-Cl– cotransporter (NCC), the target of thiazide diuretics, is responsible for the reabsorption of 5–10% of filtered NaCl. The aim of this study was to test the hypothesis that acute infusion of the angiotensin-converting enzyme (ACE) inhibitor captopril (at 12 µg/min) for 20 min provokes trafficking of NCC from apical plasma membranes (APM) to subapical cytoplasmic vesicles (SCV), which is reversed by acute ANG II infusion (ANG II at 20 ng·kg–1·min–1 along with 12 µg/min captopril) for 20 min in male Sprague-Dawley rats (250–350 g). By immuno-electron microscopy using an anti-NCC (D. Ellison) 71.5 ± SD 4.9% of the NCC gold labeling was associated with the APM in control, sham operated, and infused rats, while captopril infusion reduced NCC in APM to 54.9 ± 6.9% (P < 0.001) and markedly increased immunogold labeling of SCV. Subsequent infusion of ANG II with captopril restored NCC immunogold labeling of APM to 72.4 ± 4.2%, that is, 20% of the total NCC trafficked between APM and SCV. Likewise, on density gradients of cortex, captopril provoked redistribution of 27.3% of total NCC from low-density APM-enriched membranes to higher-density membranes and ANG II+captopril restored 20.3% of the NCC to APM-enriched fractions. Redistribution occurred independent of a change in NCC total abundance. In conclusion, this study demonstrates that ACE inhibition provokes acute trafficking of NCC out of the plasma membrane, which likely decreases DCT Na+ reabsorption, while ANG II provokes rapid trafficking of NCC from stores in subapical vesicles to the plasma membrane, which likely increases DCT Na+ reabsorption.

sodium transport; thiazide receptor; immunoelectron microscopy

THE NA+-CL– COTRANSPORTER (NCC) is expressed in the apical membrane of the distal convoluted tubule (DCT) and is the target of the thiazide diuretics, which are frequently used in the treatment of hypertension and edema (1, 29). The importance of NCC in the regulation of blood pressure and salt balance is demonstrated in the genetic disorder Gitelman's syndrome in which loss of function mutations in NCC results in salt wasting, hypokalemia, and hypotension (32). Studies in mice with NCC knocked out show that on low-sodium diets, blood pressure is significantly reduced from control (31).

There are multiple mechanisms by which NCC could be regulated to control Na+ transport in the DCT. Many studies have shown that NCC abundance is regulated by stimuli, such as dietary salt, aldosterone escape, and mineralocorticoid receptor blockade (22, 26, 33, 36), less is known about trafficking of NCC to and from the apical membrane as a way of regulating NCC. The potential importance of trafficking in the regulation of NCC has been demonstrated in vitro in Gitelman's syndrome where oocyte studies indicate that NCC is not processed properly in the endoplasmic reticulum, resulting in deficient trafficking of NCC to the plasma membrane (8, 14). A previous study from this laboratory showed that the ratio of NCC present at the apical membrane vs. intracellular pools is regulated in vivo during chronic changes in dietary salt, with less NCC present in the apical membrane during high salt intake than during low-salt intake conditions (30). Whether the trafficking of NCC to and from the plasma membrane is an important mechanism during acute conditions has yet to be determined.

ANG II is a potent vasoconstrictor and sodium-retaining hormone that plays a vital role in the regulation of blood pressure and is believed to contribute to the progression of cardiovascular and renal disease (24, 37). The role of ANG II in the regulation of NCC trafficking is not known; however, multiple studies have demonstrated the involvement of ANG II in the regulation of abundance of major renal sodium transporters such as ENaC, NKCC2, NHE3, and NCC (2, 11, 16). A role of ANG II in trafficking of the proximal tubule sodium transporter NHE3 has been demonstrated in a recent study from the McDonough laboratory: acute infusion of the angiotensin-converting enzyme (ACE) inhibitor captopril provoked the redistribution of NHE3 to the base of the proximal tubule apical microvilli accompanied by increased proximal tubule flow and diuresis (18). The presence of angiotensin II type 1 (AT1) receptors in the luminal and basolateral membranes of DCT epithelia offers a potential role for ANG II to directly regulate the DCT sodium transport (10, 25). ANG II has been shown to stimulate transport of bicarbonate, fluid, and sodium in the DCT (34), and proliferation of the DCT in an ANG II-mediated renal injury model (19, 20). Additionally, chronic treatment with the AT1 receptor antagonist candesartan together with vasopressin decreased NCC abundance compared with control or treatment with vasopressin alone (15).

The aim of this study was to test the hypothesis that acute infusion of the ACE inhibitor captopril provokes acute trafficking of NCC from the apical plasma membrane to subapical membranes and that subsequent ANG II coinfusion with captopril returns NCC to the plasma membrane.


   METHODS
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 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animal protocol. Experiments were performed using Sprague-Dawley rats (250–350 g body wt) that were kept under diurnal light conditions and had free access to food and water. Rats were anesthetized with Inactin intraperitoneally (Sigma; at 125 mg/kg), and body temperature was maintained thermostatically at 37°C. Polyethylene catheters (PE-50) were placed into the carotid artery to monitor blood pressure and into the jugular vein to infuse 4.0% BSA in 0.9% saline (at 50 µl/min) to maintain euvolemia. The left ureter was cannulated with a Surflo intravenous catheter (Terumo) for urine collection. At the completion of all surgical procedures, the animals were allowed to recover for >60 min before infusion of drugs and collection of in vivo data. Three treatment groups were compared: 1) captopril dissolved in 4% BSA and 0.9% saline was infused intravenously via the jugular vein for 20 min. 2) Following 20 min captopril infusion (12 µg/min), ANG II (20 ng·kg–1·min–1) was dissolved in 4% BSA, and 0.9% saline was coinfused with captopril (12 µg/min) for an additional 20 min. 3) Controls were infused with 4% BSA and 0.9% saline for 40 min. During the treatments, urine was collected at 10-min intervals, and volume was determined gravimetrically. Glomerular filtration rate (GFR), calculated for left kidney, was measured by infusion of FITC-inulin (5 mg/ml), as previously described by Lorenz and Gruenstein (21). Urine samples were protected from light, diluted with PBS (pH 7.4), and the fluorescence was measured (excitation = 480 nm and emission = 530 nm) with the GENios multidetection system (Phenix Research Products). After treatment, kidneys were collected for subcellular fractionation or fixed for immunoelectron microscopy. All animal experiments were approved by the University of Southern California Keck School of Medicine Institutional Animal Care and Use Committee and was conducted in accord with the National Institutes of Health Guide for the Care and Use of Laboratory Animals.

Density gradient fractionation of renal cortex. Renal cortex from each treatment group (n = 5 each) were subjected to subcellular fractionation on sorbitol density gradients. Gradients were constructed by the delivery of sorbitol to the bottom of the centrifuge tube in the following order: 1 ml of 5% sorbitol isolation buffer, a first hyperbolic gradient of 35–55% sorbitol generated by adding 18 ml of 55% sorbitol to a constant-volume mixing chamber initially loaded with 8.9 ml of 35% sorbitol, a second hyperbolic gradient of 55–70% sorbitol generated by mixing 8.2 ml of 70% sorbitol with 6.3 ml 55% sorbitol, and finally 1 ml of 80% sorbitol. The procedure for subcellular fractionation has been described in detail previously (41). In brief, the kidneys of the anesthetized rats were cooled in situ by flushing with cold PBS (to block membrane trafficking) and excised, and renal cortices were dissected and homogenized (in 5% sorbitol, 0.5 mM disodium EDTA, 0.2 mM PMSF, 9 µg/ml aprotinin, and 5 mM histidine/imidazole buffer pH 7.5) with a Tissuemizer (Tekmar Instruments), centrifuged at 2,000 g for 10 min (supernatant retained); the pellet was rehomogenized and centrifuged, and the two low-speed supernatants (S0) were pooled. A 4-ml aliquot of S0 sample was mixed with 6 ml 87.4% sorbitol, equilibrated on ice for 1 h, loaded between the two hyperbolic sorbitol gradients, centrifuged at 100,000 g for 5 h. Twelve fractions were collected, and the density of each was measured using a DA-100M Density/Specific Gravity meter (Mettler Toledo). Fractions were diluted, pelleted at 250,000 g for 75 min, resuspended in 1 ml isolation buffer and stored at –80°C pending assay.

To assess density distribution pattern of NCC, a constant volume (8 µl) of each of the 12 fractions was assayed. To assess the total pool size of NCC in crude membranes, a constant volume from each of the 12 fractions was pooled together into "total membrane" samples; protein concentration were measured, and a constant amount (8.8 µg and 4.4 µg) of total membrane protein from each sample was analyzed side by side on the same gel. Samples were denatured in SDS-PAGE sample buffer for 30 min at 37°C, resolved on 7.5% SDS polyacrylamide gels (17) and transferred to polyvinylidene diflouride membranes (Millipore Immobilon-P). In all immunoblot assays, one-half of the volume was also assayed in parallel to verify linearity of the detection system. After blocking, we incubated the blots with polyclonal antiserum against the amino terminal putative cytoplasmic domain of NCC, which was not affinity purified (4) at 1:500 dilution, then incubated with Alexa 680-labeled goat anti-rabbit (Molecular Probes), and detected and quantitated with the Odyssey Infrared Imaging System (Li-COR), and accompanying LI-COR software. Results were expressed as the percentage of the total on the gradient (signal in all 12 fractions = 100%).

Indirect immunofluorescence. At the end of the experiment, kidneys were perfusion-fixed with 4% paraformaldehyde solution, containing 0.1 M sodium cacodylate at pH 7.2, and excised. Tissue blocks were trimmed from the cortex and fixed for 2 h, rinsed in PBS, and infiltrated with 2.3 M sucrose, mounted on holders, and frozen in liquid nitrogen (23, 40). Cryosections (1.0 µm) were cut on a Reichert Ultracut S cryoultramicrotome (Leica), transferred on 2.3 M sucrose to charged glass slides, rinsed in PBS incubated with 0.05 M NH4Cl, and blocked by incubation in PBS containing 0.05 M glycine and 0.1% skim milk powder. Sections were labeled with polyclonal antiserum against NCC at 1:1,600; then they were incubated for 1 h with AlexaFluor 488-conjugated goat-anti-rabbit secondary antibodies (Molecular Probes) at 1:300 in PBS containing 0.1% skim milk powder, and were washed and mounted. Slides were viewed and recorded with a Leica TSC SP2-inverted confocal laser scanning microscope (Leica, Mannheim, Germany).

Immunoelectron microscopy. To analyze NCC subcellular distribution at the ultrastructural level, kidneys from six animals, each from control, captopril and captopril/ANG II regimes were analyzed by immuno-electron microscopy (EM). The fixation procedure is as described in the previous paragraph. Immuno-EM was performed on thin (~70 nm) cryosections prepared with random orientation from the frozen cortical tissue on a Reichert Ultracut S cryoultramicrotome (Leica), as previously described (23). The cryosections were first blocked by incubation in PBS containing 0.05 M glycine and 0.1% skim milk powder. The sections were then incubated for 1 h at room temperature or overnight in 4°C with the polyclonal anti-NCC at 1:1,600 dilution in PBS containing 0.1% skim milk powder. The primary antibody was visualized using goat anti-rabbit IgG conjugated to 10-nm colloidal gold particles (GAR EM10, BioCell Research Laboratories, Cardiff, UK), diluted 1:50 in PBS with 0.1% skim milk powder and polyethyleneglycol (5 mg/ml). The cryosections were stained with 0.3% uranyl acetate in 1.8% methyl-cellulose for 10 min before examination in a FEI Morgagni electron microscope. For quantitation, electron micrographs were recorded at x16,000 on Kodak 163 film and printed at x50,000.

Immunolabeling controls consisted of substitution of the specific primary antibody with nonimmune rabbit IgG or incubation without primary antibody.

Quantitation and statistical analysis. Data are expressed as means (±SD). Differences were regarded significant at P < 0.05.

For blood pressure, urine output, GFR, and NCC abundance paired two-tailed Student's t-test assuming equal variance was used to establish whether there was a significant effect of treatment on these parameters within the set of animals. In density gradient assays, two-way ANOVA was applied to determine whether there was a significant effect of treatment on the overall density NCC distribution pattern. After significance was established, the location of the difference in the pattern was assessed by unpaired two-tailed Student's t-test assuming equal variance with Bonferroni adjustments for multiple comparisons.

Quantitation of Immunogold labeling was performed on electron micrographs of DCT cells from six animals in each group. From each rat 6–10 cells were analyzed on sections oriented approximately at right angle to the apical cell membrane. The cells originated both from DCT1 and DCT2 (7, 27). Gold particles were ascribed to the apical cell membrane when located on its outer surface, directly above, or within 30 nm from its inner surface. Gold particles over the apical cytoplasm were counted below the analyzed stretch of cell membrane down to depth of 2 µm in two zones (Fig. 1). The first zone extended from the cell membrane down to a depth of 1 µm (except for the first 30 nm), and the second zone extended from a depth of 1 to 2 µm into the cytoplasm. The percentage of particles associated with the cell membrane, and the two cytoplasmic zones, compared with all counted particles, was determined for each cell. The mean percentage was then determined for each animal and the means for control vs. captopril and control vs. captopril/ANG II compared by two-tailed Student's t-test for unpaired samples. Background labeling was very low (~1 gold particle/µm2) and similar over all DCT nuclei, basal cytoplasm, and over surrounding proximal tubules. Counts over each DCT cell were corrected for background labeling.


Figure 1
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  		Fig. 1. Survey electron micrograph of distal convoluted tubule (DCT) wall illustrating the two apical cytoplasmic zones (Z1 and Z2) below the apical plasma membrane, where NCC immunogold labeling was quantified. The distances of the lines from the level of the apical plasma membrane are 1 and 2 µm, respectively. Apical NCC labeling of these cells was verified at higher magnification.

 

   RESULTS
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 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Effect of ACE inhibition and restoration of ANG II on mean arterial pressure and renal function. Captopril infusion at 12 µg/min or coinfusion of captopril along with ANG II at 20 ng·kg–1·min–1 had no significant effect on mean arterial pressure (MAP; baseline 99 ± 9 mmHg; captopril: 97 ± 12 mmHg; captopril + ANG II: 102 ± 13 mmHg; Fig. 2A). Urine output (volume/time), normalized to control group defined as 1, increased with captopril infusion from 1.00 ± 0.55 to 1.46 ± 0.47 and returned to baseline levels during coinfusion of captopril and ANG II to 1.13 ± 0.41. (Fig. 2C). This increase in urine output occurred in the absence of a change in GFR (baseline: 5.4 ± 0.8 ml·min–1·kg–1; with captopril: 5.2 ± 1.7 ml·min–1·kg–1; captopril + ANG II 5.8 ± 0.8 ml·min–1·kg–1; Fig. 2B).


Figure 2
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  		Fig. 2. Effects of angiotensin-converting enzyme (ACE) inhibition by captopril and subsequent captopril+ANG II infusion on mean arterial pressure (MAP; A), glomerular filtration rate (GFR; B) and urine output (C). Control, rats infused with 4.0% BSA in 0.9% saline (50 µl/min) for >30 min. Captopril, same rats subsequently infused with captopril (12 µg/min) for 20 min. Captopril+ANG II, same rats subsequently coinfused with captopril (12 µg/min) and ANG II (20 ng·kg–1·min–1) for an additional 20 min. Data presented collected at 0–10 min before (control), 10–20 min during captopril infusion, and 10–20 min during coinfusion of captopril and ANG II. A: MAP recorded from the carotid artery. B: GFR from clearance of FITC-inulin. C: Urine output measured gravimetrically and normalized to mean control. *P < 0.05 vs. control, #P < 0.05 vs. captopril assessed by paired Student's t-test.

 
Effect of ACE inhibition by captopril and ANG II restoration on subcellular distribution of NCC. On the basis of our previous study showing that NCC subcellular distribution between apical and subapical membranes is altered in response to changes in dietary salt, which also changes ANG II levels (30), we hypothesized that ANG II blockade by captopril would lead to a removal of NCC from apical membranes and that restoring ANG II would return NCC to the apical membrane. After sorbitol density gradient, centrifugation of renal cortex membranes, a constant volume of each fraction was analyzed by immunoblot to ascertain the density distribution pattern. After determining the density of the signal in each lane (in arbitrary units), the total density is calculated, defined as 100%, and the percent of the total signal in each fraction calculated for each treatment group. Thus, the results reflect the density distribution pattern of the protein probed, independent of variations in the amount of total protein present in the gradients or variation in protein concentrations across the gradients. The pattern was assessed by ANOVA, and, only if shown to be significantly altered by treatment, individual fractions were compared for statistical difference.

As shown in Fig. 3A, infusion of captopril (12 µg/min) for 20 min provoked a significant redistribution of 27.3% of total NCC detected on the gradient from low-density apical enriched membrane fractions (fractions 3 and 4) to higher-density membranes (fractions 6 and 7). Subsequent coinfusion of ANG II (20 ng·kg–1·min–1) with captopril (12 µg/min) provoked the restoration of 20.3% of NCC from higher-density to low-density apical enriched membranes (Fig. 3B). The 20-min acute treatments with captopril or captopril plus ANG II did not alter the total pool size of renal NCC (control 1.00 ± 0.45; captopril: 0.96 ± 0.31; captopril+ANG II: 1.19 ± 0.32; Fig. 3C).


Figure 3
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  		Fig. 3. Effects of ACE inhibition by captopril and subsequent captopril+ANG II infusion on density distribution and abundance of NCC. A: density distribution of NCC in control (bullet) vs. 20 min of acute captopril infusion ({circ}). B: density distribution of NaCl cotransporter (NCC) in control (bullet) vs. 20 min captopril infusion followed by coinfusion of captopril and ANG II ({triangleup}). Relative abundance, determined by immunoblot of a constant volume of each fraction, expressed as a percentage of total signal in all 12 fractions; n = 5 in each group. Values are means (±SD). *P < 0.05 compared with corresponding control. C: total abundance of NCC detected by immunoblots of a constant amount of protein (8.8 µg) from total homogenate (So) from each treatment group.

 
Effect of ACE inhibition by captopril and restoration of ANG II on NCC distribution in DCT cells. Confocal immunofluorescence and differential interference contrast analysis of NCC in control and captopril-treated animal is shown in Fig. 4. The anti-NCC antibody specifically stains the apical membrane of the DCT and, perhaps, the region just below apical membrane. This region of staining is quite narrow due to the paucity of microvilli in the DCT and was not obviously shifted to intracellular pools during acute captopril treatment. The same labeling pattern was observed both in DCT1 and DCT2 in four control and four captopril-treated animals. In DCT2, single cells or patches of cells were not stained (not shown), as has been previously observed in this region as it transitions to CNT (connecting tubule). Thus, we proceeded with immuno-EM to obtain direct and quantifiable evidence for acute trafficking of NCC by captopril and ANG II.


Figure 4
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  		Fig. 4. Indirect immunofluorescence microscopy (A and C) and overlay with differential interference contrast (B and D) of NCC in rat renal cortex of control (A and B) and captopril-treated animals (C and D). The anti-NCC antibody specifically stained the apical membrane region of the DCT in both groups. Scale bar = 40 µm.

 
Immuno-EM revealed the subcellular localization of NCC in DCT cells of control, captopril, and captopril+ANG II-infused rats. In all three groups, NCC was associated both with the apical plasma membrane and with a population of vesicles located in the apical regions of the cytoplasm, but the quantitative distributions were characteristic for each group. The overall fine structure of the apical and basolateral plasma membrane, as well as the microvilli was the same in the three groups.

In all groups, NCC labeling was observed along the entire apical plasma membrane, both over the small microvilli and between the microvilli (Fig. 5, A–C). Much of the gold label over the plasma membrane was associated with a stained inner layer of the membrane, possibly representing the cytoplasmic parts of the transporter (Fig. 6A). The cytoplasmic vesicles were 0.01–0.05 µm in diameter, and the majority appeared devoid of a clathrin-like cytoplasmic coat (Fig. 6B). Occasional multivesicular bodies with NCC-labeled vesicles were observed in the apical regions of DCT cells, notably in captopril-infused rats (Fig. 5B, inset). The frequency of labeled vesicles decreased with increasing distance from the apical plasma membrane and no specific label was observed in the basal cytoplasm. Adjacent cells in proximal and cortical collecting tubules were also devoid of specific labeling, as were intercalated cells occasionally observed adjacent to DCT cells. The same overall labeling pattern was observed both in DCT I and DCT II.


Figure 5
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  		Fig. 5. Immunoelectron microscope localization of NCC in DCT cells of control (A), captopril (B and D), and captopril+ANG II (C) infused rats. The polyclonal NCC antibody was detected with goat anti-rabbit IgG conjugated to 10-nm colloidal gold particles. In each of the three conditions, NCC is associated both with the apical plasma membrane and apical cytoplasmic vesicles. However, the relative proportions are different. Thus, in controls (A), labeling of the apical plasma membrane prevails, while after captopril infusion (B), a larger proportion of the label is associated with the apical cytoplasmic vesicles. After captopril+ANG II infusion (C), labeling of the apical plasma membrane is again dominating. Inset in B: labeling of vesicles in multivesicular body after captopril infusion. D: unspecific labeling of the basal parts of the cells is very low. Scale bars: 1 µm.

 

Figure 6
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  		Fig. 6. High-resolution immunoelectron microscope localization of NCC in apical parts of DCT cells in control (A) and captopril (B) infused rats. NCC immunogold labeling is often associated with a stained dense layer on the inner surface of the apical plasma membrane (A). The apical cytoplasmic vesicles vary slightly in size and shape. NCC labeling is observed at the vesicle membrane or within the vesicle (B). Scale bars: 100 nm.

 
Quantitative analyses (Fig. 7) showed that 71.5 ± 4.9 (SD)% of the NCC gold labeling was associated with the apical plasma membrane in controls. The 20-min captopril infusion reduced NCC in apical plasma membranes (APM) to 54.9 ± 6.9% (P < 0.001), and at the same time there was a marked increased labeling of cytoplasmic vesicles. Subsequent coinfusion of captopril and ANG II restored NCC labeling of the apical plasma membrane to 72.4 ± 4.2%, which is not significantly different from control, but significantly (P < 0.001) higher than after infusion of captopril alone.


Figure 7
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  		Fig. 7. Quantitative distribution of NCC in DCT cells analyzed by immunoelectron microscopy following control, white bar; captopril, gray bar; and captopril+ANG II infusion, hatched bar. Colloidal gold particles representing NCC antibodies were counted along the apical cell membrane, in the apical zone down to 1 µm below the membrane (Z1), and the zone between 1 and 2 µm below the membrane (Z2), as illustrated in Fig. 1. In controls, more than 70% of NCC is associated with the membrane but decreases significantly following captopril infusion. NCC traffics back to the membrane following captopril+ANG II infusion. Simultaneously, NCC increases significantly in cytoplasmic vesicles in both Z1 and Z2 following captopril infusion but traffics back to the apical plasma membrane following captopril+ANG II infusion.

 

   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
This study aimed to investigate whether abundance of NCC in the plasma membrane is acutely regulated by ANG II. Regulation by ANG II was chosen for study because there was micropuncture evidence that ANG II acutely stimulated reabsorption of bicarbonate, fluid, and sodium in the DCT (35). To isolate the primary effects of ANG II on NCC trafficking, a protocol was designed that did not alter MAP or GFR (Fig. 2). The results of our study, from both density gradient cellular fractionation and direct immunoelectron microscopy, support the hypothesis that ACE inhibition provokes internalization of NCC to subapical vesicles, along with diuresis and that ANG II provokes NCC trafficking from subapical vesicles the APM.

NCC, expressed in the apical membrane of the DCT (28), is responsible for reabsorption of just 5–10% of filtered Na+ and Cl– (28), yet the importance of NCC in sodium homeostasis and blood pressure regulation is supported by 1) the efficacy of thiazide diuretics to lower blood pressure, 2) the salt wasting and hypotension evident in Gitelman's syndrome in which NCC is inactive (8, 14), and 3) the hypertension evident in pseudohypoaldosteronism type II (PHA II) where mutations in WNK [with no lysine kinases (K)] WNK1 and WNK4 are linked to increased NCC activity (38, 39).

That trafficking of NCC could be important in the regulation of sodium transport of the DCT has been suggested by studies of genetic disorders. When the genetic variants of WNK1 or WNK4 found in PHA II are expressed in oocytes along with wild-type NCC, plasma membrane expression of NCC is substantially higher than when NCC is coexpressed with wild-type WNK1 and WNK4, leading to the speculation that this causes the increased sodium reabsorption and hypertension in PHA II (38, 39). When mutant NCC found in the salt wasting genetic condition Gitelman's syndrome is expressed in oocytes, it is not processed properly in the endoplasmic reticulum, resulting in deficient trafficking of NCC to the plasma membrane, leading to the hypothesis that this causes the decreased sodium uptake in the DCT in this disorder (8, 14).

ANG II is an important regulator of sodium homeostasis and blood pressure and is believed to contribute to the progression of cardiovascular and renal disease (24, 37). ACE inhibitors are commonly prescribed for the treatment of hypertension and are thought to exert most of their antihypertensive effects through actions on the kidney (6, 9). Whether or not ANG II acutely regulated NCC trafficking was an open question before this current study. Our previous study demonstrated that chronic changes in dietary salt, which drive changes in ANG II levels, altered the percentage of total NCC present at the apical membrane. Specifically, low-salt diet, which increases plasma ANG II, increased apical NCC, assessed by both density gradients and immuno-EM (30). Captopril treatment, in this current study provoked a redistribution of NCC out of the apical plasma membrane evident by both immuno-EM and redistribution on density gradients. There is fairly good agreement between these very distinct methods of assessment used to assess NCC redistribution: the magnitude of the captopril-induced redistribution of NCC out of the apical membrane assessed by immuno-EM was 17% while that assessed from density gradient redistribution was 27%.

In our previous study of the effects of a chronic high-salt diet, we measured, by immuno-EM, a 32% decrease in NCC in the apical membrane, as well as a 50% decrease in total cellular NCC (30). Chronic inhibition of ANG II during a high-salt diet could also decrease aldosterone levels, known to be a significant regulator of NCC abundance (25). Aldosterone is unlikely to play a role in the response to 20 min captopril treatment since it takes at least 30 min for aldosterone to induce mRNA expression of SGK1, the kinase thought to mediate the rapid effects of aldosterone (3). In a concurrent study, we have discovered that chronic ACE inhibitor treatment of spontaneously hypertensive rats (SHR) sufficient to normalize blood pressure (4 wk enalapril in drinking water) caused a persistent shift of the NCC density distribution pattern from a peak around fraction 4 to a peak around fraction 5, similar to the acute effects of ACE inhibition, which provides evidence that the SHR does not escape from the effects of ACE inhibition (L. E. Yang, P. K. Leong, and A. A. McDonough, unpublished observations).

Previously, the McDonough laboratory showed that acute infusion of the ACE inhibitor captopril, analogous to the protocol used in this study, led to redistribution of the proximal tubule sodium transporter Na+/H+ exchanger isoform 3, NHE3, from the body to the base of the microvilli associated with decreased fluid reabsorption, establishing a direct role of ANG II to regulate trafficking of sodium transporters (18). Thus, it appears that captopril treatment initiates redistribution of transporters within the proximal tubule from the body to the base of the microvilli and in the distal tubule from the apical cell membrane to cytoplasmic vesicles coincident with reduced Na+ reabsorption. Apical distribution of NCC can be decreased by either an increase in the rate of removal of NCC from APM or a decrease in the rate of delivery to APM. We cannot conclude from the studies conducted whether captopril or ANG II affects rates of delivery to or from the APM.

Is it possible that the effects of captopril on DCT NCC are secondary to the effects of captopril in the proximal tubule? The presence of AT1 receptors in the luminal and basolateral membranes of DCT epithelia offers a potential for ANG II to directly regulate the DCT (10, 25). Yet, in the study of the effects of captopril on trafficking of sodium transporters in the proximal tubule, a 20% increase in flow out of the proximal tubule was observed (18), which could impact regulation of transport beyond the proximal tubule. However, it is known that increased sodium and volume delivery to the DCT, e.g., secondary to furosemide treatment, actually increases sodium reabsorption in the DCT (13), opposite to the predicted effect of NCC internalization by captopril treatment. Thus, it appears that the reduced ANG II levels affect DCT NCC independent of effects in the proximal tubule.

Although most of the beneficial effects of ACE inhibitors have generally been attributed to the blockade of the ACE-dependent formation of ANG II, ACE inhibitors also induce the accumulation of bradykinin and activate the ACE signaling cascade (6, 9) that together would exert a complex array of effects on the kidney, including inhibition of sodium reabsorption. The issue of whether the captopril-mediated effects were mediated by accumulation of bradykinins or lack of ANG II was indirectly addressed by restoring ANG II by coinfusion with captopril. The NCC internalization was rapidly reversed by acute infusion of ANG II, suggesting that the effects stemmed from the blockade of endogenous ANG II formation rather than bradykinin accumulation.

A study by Verlander et al. (34) showed that ovariectomy led to decreased complexity of the DCT apical surface and a proportional decrease in immunogold detection of apical NCC, both restored with chronic estradiol replacement. Changes in basolateral structure have been observed in DCT after prolonged furosemide treatment (12). In our acute study, there was no apparent change in apical or basal complexity, and NCC total abundance was unaffected by either acute ACE inhibition or subsequent ANG II infusion (Fig. 3). We conclude that trafficking of NCC out of the apical membrane, rather than loss of apical complexity, is the primary mechanism of action of captopril to acutely decrease apical NCC. It also remains possible that a decrease in the rate of apical delivery of NCC, which may be sensitive to ANG II, as opposed to an increase in NCC internalization rate, could account for the decrease in apical NCC during captopril if there is rapid cycling of NCC between apical and subapical pools.

In conclusion, this study provides evidence for rapid regulated trafficking of DCT NCC between the apical plasma membrane and subapical vesicles. The results provide evidence that treatment with ACE inhibitors rapidly removes a significant fraction of NCC from the apical membrane, which likely decreases DCT sodium reabsorption and contributes to the blood pressure lowering effects of captopril. The results also provide evidence that localization of NCC to the apical plasma membrane requires baseline levels of ANG II. The acute trafficking of NCC between apical and subapical pools provides a previously unreported mechanism by which captopril and ANG II, or other regulators, are able to rapidly regulate renal sodium reabsorption.


   GRANTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by HL-085388 and DK-34316 (A. A. McDonough) and the Danish Medical Research Council and the Water and Salt Research Center established and supported by the Danish National Research Foundation (Grundforskningsfonden) (A. B. Maunsbach).


   ACKNOWLEDGMENTS
 
We thank Else-Merete Løcke and Karen Thomsen for expert technical assistance. Monica Sandberg was partially supported by predoctoral fellowship from the American Heart Association Western States Affiliate.


   FOOTNOTES
 

Address for reprint requests and other correspondence: Arvid B. Maunsbach, The Water and Salt Research Center, Dept. of Cell Biology, Institute of Anatomy, Univ. of Aarhus, DK-8000 Aarhus C, Denmark (e-mail: maunsbach{at}ana.au.dk)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 

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