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The renal thiazide-sensitive Na-Cl cotransporter as mediator of the aldosterone-escape phenomenon
Xiao-Yan Wang,1 Shyama Masilamani,1 Jakob Nielsen,1 Tae-Hwan Kwon,2 Heddwen L. Brooks,1 Søren Nielsen,2 and Mark A. Knepper1
1Laboratory of Kidney and Electrolyte Metabolism, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, Maryland, USA2 Department of Cell Biology, Institute of Anatomy, University of Aarhus, Aarhus, Denmark
Address correspondence to: Mark A. Knepper, National Institutes of Health, Building 10, Room 6N260, 10 Center Drive MSC 1603, Bethesda, Maryland 20892-1603, USA. Phone: (301) 496-3064; Fax: (301) 402-1443; E-mail: knep@helix.nih.gov.
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1Laboratory of Kidney and Electrolyte Metabolism, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, Maryland, USA2 Department of Cell Biology, Institute of Anatomy, University of Aarhus, Aarhus, Denmark
Address correspondence to: Mark A. Knepper, National Institutes of Health, Building 10, Room 6N260, 10 Center Drive MSC 1603, Bethesda, Maryland 20892-1603, USA. Phone: (301) 496-3064; Fax: (301) 402-1443; E-mail: knep@helix.nih.gov.
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1Laboratory of Kidney and Electrolyte Metabolism, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, Maryland, USA2 Department of Cell Biology, Institute of Anatomy, University of Aarhus, Aarhus, Denmark
Address correspondence to: Mark A. Knepper, National Institutes of Health, Building 10, Room 6N260, 10 Center Drive MSC 1603, Bethesda, Maryland 20892-1603, USA. Phone: (301) 496-3064; Fax: (301) 402-1443; E-mail: knep@helix.nih.gov.
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1Laboratory of Kidney and Electrolyte Metabolism, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, Maryland, USA2 Department of Cell Biology, Institute of Anatomy, University of Aarhus, Aarhus, Denmark
Address correspondence to: Mark A. Knepper, National Institutes of Health, Building 10, Room 6N260, 10 Center Drive MSC 1603, Bethesda, Maryland 20892-1603, USA. Phone: (301) 496-3064; Fax: (301) 402-1443; E-mail: knep@helix.nih.gov.
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1Laboratory of Kidney and Electrolyte Metabolism, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, Maryland, USA2 Department of Cell Biology, Institute of Anatomy, University of Aarhus, Aarhus, Denmark
Address correspondence to: Mark A. Knepper, National Institutes of Health, Building 10, Room 6N260, 10 Center Drive MSC 1603, Bethesda, Maryland 20892-1603, USA. Phone: (301) 496-3064; Fax: (301) 402-1443; E-mail: knep@helix.nih.gov.
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1Laboratory of Kidney and Electrolyte Metabolism, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, Maryland, USA2 Department of Cell Biology, Institute of Anatomy, University of Aarhus, Aarhus, Denmark
Address correspondence to: Mark A. Knepper, National Institutes of Health, Building 10, Room 6N260, 10 Center Drive MSC 1603, Bethesda, Maryland 20892-1603, USA. Phone: (301) 496-3064; Fax: (301) 402-1443; E-mail: knep@helix.nih.gov.
Find articles by Nielsen, S. in: JCI | PubMed | Google Scholar
1Laboratory of Kidney and Electrolyte Metabolism, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, Maryland, USA2 Department of Cell Biology, Institute of Anatomy, University of Aarhus, Aarhus, Denmark
Address correspondence to: Mark A. Knepper, National Institutes of Health, Building 10, Room 6N260, 10 Center Drive MSC 1603, Bethesda, Maryland 20892-1603, USA. Phone: (301) 496-3064; Fax: (301) 402-1443; E-mail: knep@helix.nih.gov.
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Published July 15, 2001 - More info
J Clin Invest. 2001;108(2):215–222. https://doi.org/10.1172/JCI10366.
© 2001 The American Society for Clinical Investigation
Received: May 18, 2000; Accepted: June 4, 2001
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Methods
Aldosterone-escape model. Experiments were conducted in male Sprague-Dawley rats (180–220 g) (Taconic Farms, Germantown, New York, USA). The aldosterone-escape protocol is diagrammed in Figure 1. All rats were maintained in metabolism cages to allow quantitative urine collections. On day –3, all rats were anesthetized with methoxyflurane (Metofane; Pitman-Moore, Mundelein, Illinois, USA) and implanted subcutaneously with osmotic minipumps (model 2ML2; Alzet, Palo Alto, California, USA) delivering 200 μg per day of aldosterone (Sigma Chemical Co., St. Louis, Missouri, USA) (6). The minipump infusion was sustained throughout the entire time course of each experiment in all rats. The intake of Na was initially maintained at a very low level (0.02 mEq/d) by ration feeding of measured amounts of a gelled mixture of low-Na food and water (see below). On day 0 (see Figure 1), half the rats were switched to a higher Na intake (2.0 mEq/d), whereas the remaining rats were continued on the 0.02 mEq/d Na intake. The rats given the higher Na intake must escape from the Na-retaining effect of aldosterone to re-establish Na balance.
Figure 1Diagram of aldosterone-escape protocol. All rats received aldosterone infusions by osmotic minipump (top). Both control and escape rats were started at day –3 on a low-Na diet providing approximately 0.02 mEq/d of Na. On day 0, the Na intake was increased to 2.0 mEq/d for the escape rats, whereas control rats continued to receive 0.02 mEq/d.
Kidneys were analyzed at various time points after the switch from 0.02 to 2.0 mEq/d Na to determine Na transporter and Na channel abundance as described below. The low-Na control rats were handled in a manner identical to the experimental rats throughout the time course, including matched caloric and water intake achieved by ration feeding (see below). Rats were euthanized by decapitation. Serum was collected at the time of decapitation for the measurement of aldosterone concentration by RIA (Coat-a-Count; Diagnostic Products Corp., Los Angeles, California, USA), Na concentration, and creatinine concentration (Monarch 2000 autoanalyzer; Instrumentation Laboratories, Lexington, Massachusetts, USA). Urinary samples were analyzed for Na and creatinine (Monarch 2000 autoanalyzer).
Ration feeding protocol. The intakes of sodium, calories, and water were carefully controlled by ration feeding of a fixed daily amount of a gelled diet that contains all of the nutrients, NaCl, and water that the rat receives in a day (6). The baseline low Na intake was achieved by feeding the rats a gelled mixture of a synthetic low-Na diet (Formula 53140000; Ziegler Brothers, Gardner, Pennsylvania, USA), deionized water (25 ml per 15 g of food), and agar (0.125 g per 25 ml of water). The Na-replete diet was the same except for addition of 2 mEq Na per 15 g of food before gelation. All animals received the equivalent of 15 g per 200 g of body weight (BW) per day of rat chow determined by weighing the gelled mixture. Analysis of the diet demonstrated that this protocol provides approximately 0.02 mEq per 200 g of BW per day of Na for the low-Na mixture and 2.0 mEq per 200 of g BW per day for the Na-replete diet.
Antibodies. Rabbit polyclonal antibodies to the following renal sodium transporters were utilized: the type 2 Na-phosphate cotransporter (NaPi-2) of the proximal tubule (14), the type 3 Na-H exchanger (NHE3) of the proximal tubule (15), the Na-K-2Cl cotransporter (NKCC2) of the thick ascending limb (16), the thiazide-sensitive Na-Cl cotransporter (NCC) of the DCT (6), and all three subunits of the epithelial Na channel (ENaC) of the collecting duct (4). The antisera were affinity purified against the immunizing peptides as described previously (6, 16). Specificity of the antibodies has been demonstrated by showing unique peptide-ablatable bands on immunoblots and a unique distribution of labeling by immunocytochemistry. In addition, we utilized a mouse mAb against the Na-K-ATPase α1 subunit (product number 05-369; Upstate Biotechnology Inc., Lake Placid, New York, USA).
Semiquantitative immunoblotting. Semiquantitative immunoblotting was used to compare sodium transporter or channel abundance between groups of rats as described in detail previously (13, 16, 17). The left kidneys were homogenized intact. The right kidneys were dissected to obtain cortex, inner stripe of outer medulla, and inner medulla. For each set of samples (experimental and control), after solubilization in Laemmli sample buffer, an initial gel was stained with Coomassie Blue as described previously (18) to confirm equal loading among samples. SDS-PAGE was performed on 7.5%, 10%, or 12% polyacrylamide gels (Ready Gels; Bio-Rad Laboratories Inc., Hercules, California, USA), and the proteins were transferred from the gel electrophoretically to nitrocellulose membranes. Membranes were probed overnight at 4°C with the respective primary antibodies and then exposed to secondary antibody (goat anti-rabbit IgG conjugated with horseradish peroxidase, no. 31463, diluted to 1:5,000; or rabbit anti-mouse IgG conjugated with horseradish peroxidase, no. 31450, diluted to 1:5,000; both from Pierce Chemical Co., Rockford, Illinois, USA) for 1 hour at room temperature. Sites of antibody-antigen reaction were visualized using a luminol-based enhanced chemiluminescence substrate (LumiGLO; Kirkegaard and Perry Laboratories, Gaithersburg, Maryland, USA) before exposure to X-ray film (Kodak 165-1579; Eastman Kodak Co., Rochester, New York, USA). The band densities were quantitated by laser densitometry (Model PDS1-P90; Molecular Dynamics Inc., Sunnyvale, California, USA). The densitometry values were normalized to facilitate comparisons.
Immunocytochemistry. Control (n = 3) and aldosterone-escape (n = 3) rats were prepared as described above. The kidneys were fixed by perfusion with cold PBS (pH 7.4) for 15 seconds via the abdominal aorta followed by cold 4% paraformaldehyde in 0.1M cacodylate buffer (pH 7.4) for 3 minutes. The kidneys were removed and postfixed for 1 hour, followed by three 10-minute washes with 0.1M cacodylate buffer (pH 7.4). The tissue was dehydrated in graded ethanol and left overnight in xylene. The tissue was embedded in paraffin, and 2-μm sections were cut on a rotary microtome (Leica,Herlev, Denmark). Localization of NCC was carried out using indirect immunofluorescence or immunoperoxidase labeling as described elsewhere (19). The primary antibody was rabbit anti-rat NCC L573 (6). For confocal microscopy, the secondary antibody was Alexa 488 conjugated goat anti-rabbit IgG antibody (A-11008; Molecular Probes Inc., Eugene, Oregon, USA). For immunoperoxidase labeling, the secondary antibody was horseradish peroxidase conjugated to goat anti-rabbit IgG (P448; DAKO A/S, Glostrup, Denmark). For immunoperoxidase labeling, counterstaining was done using Mayer’s hematoxylin. Microscopy was carried out with Leica DMRE light microscope (Leica, Herlev, Denmark) and a Zeiss LSM510 laser confocal microscope (Brock & Michelsen, Birherød, Denmark).
In some of the immunofluorescence studies, double labeling was carried out using the mAb to the Na-K-ATPase α-1 subunit (see above) with a goat anti-mouse secondary antibody conjugated to Alexa 546 (A11003; Molecular Probes Inc.).
RNA preparation. Total kidney RNA was extracted from the right kidneys of control and aldosterone-escape rats treated as described previously (20) using RNAzol B (Tel-Test Inc., Friendswood, Texas, USA). RNA purity and concentration were assessed spectrophotometrically. RNA integrity was confirmed by inspection of ribosomal RNA bands on ethidium bromide–stained agarose gels.
RNA probe synthesis. The DNA template for the synthesis of the NCC RPA probe was generated by PCR from rat kidney cDNA using the following primers: NCC Forward primer 5′-CCT GCT GTG TCA TCA CAT C-3′, and NCC T7 Reverse primer: 5′-GCG CGT AAT ACG ACT CAC TAT AGG GAG AGG AGT TAA GGG AAC AGG AGG AGA AAT G-3′. The reverse primer (antisense primer) was made with the T7 sequence incorporated at the 5′-end of the PCR primer. The PCR product was used as the template without cloning it. The DNA template for the β-actin probe was provided by the manufacturer (7794; Ambion Inc., Austin, Texas, USA). Biotin-labeled antisense RNA probes for NCC and β-actin were synthesized from the templates with the MEGAscript kit (Ambion Inc.). The product was excised from 10% acrylamide SDS-PAGE gel following visualization with thin layer chromotography ultraviolet shadowing plates, and was eluted using the manufacturer’s buffer (414; Ambion Inc.).
RPA. The RPA was carried out using an RPA assay kit (RPA III, 1414; Ambion Inc.) according to the protocol recommended by the manufacturer. Solution hybridization was performed with 10 μg total RNA samples from control and aldosterone-escape kidneys using the biotin-labeled antisense RNA probes for NCC (1 ng per tube) and β-actin (10 ng per tube). The hybridization was carried out at 56°C. After ribonuclease digestion, the products were denatured by heating to 94°C for 4 minutes and run on a urea-TBE polyacrylamide gel (5% polyacrylamide) and transferred to nylon membranes. The products were visualized by incubation of the membranes with an alkaline phosphatase/streptavidin probe and incubation with a chemiluminescence reagent (Bright-Star; Ambion Inc.) followed by exposure to light-sensitive film (BioMax Light, double coated; Kodak). Band densities were assessed by densitometry (PDS1-P90; Molecular Dynamics Inc.). Positive and negative controls were run with the probe alone (plus yeast RNA), with and without digestion.
Presentation of data and statistical analyses. Statistical comparisons were accomplished by unpaired t test (when variances were the same) or by Mann-Whitney rank-sum test (when variances were significantly different between groups). P values less than 0.05 were considered statistically significant.
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Results
Aldosterone escape model. Figure 1 summarizes the aldosterone escape protocol. The escape phenomenon was elicited in aldosterone-infused rats by switching from a low to a high NaCl intake. Control rats received the same aldosterone infusion but were maintained on a low-NaCl intake and thus did not undergo escape. Table 1 shows data on BW, serum aldosterone concentration, creatinine clearance, water excretion rate, Na excretion rate, and urinary Na concentration in the initial set of rats studied. In the experimental group, the Na excretion rate rose virtually to the level of the intake, indicating that these rats successfully escaped from the Na-retaining action of aldosterone. The rise in Na excretion in the aldosterone-escape group occurred without a change in creatinine clearance, indicating that the increase in excretion was due to decreased renal tubule absorption.
Na transporter abundances in kidney. To determine whether the increase in Na excretion in the aldosterone-escape rats was associated with a decrease in the abundance of any of the major Na transporters expressed along the renal tubule, we screened renal homogenates using an ensemble of affinity-purified rabbit polyclonal antibodies. Figure 2 shows the initial results for renal cortex and outer medulla. Quantitative analysis of the data is shown in Table 2. In renal cortex, the major finding was a marked decrease in the abundance of NCC (the thiazide-sensitive cotransporter of the DCT) in aldosterone-escape rats. The NCC abundance fell to 17% ± 5% of the mean for the control rats (Table 2). The band densities corresponding to NHE3, NaPi-2, NKCC2, the α subunit of ENaC, and the β subunit of ENaC were not significantly changed in the aldosterone escape group compared with the aldosterone/low Na control rats. Although there was no change in the abundance of the 85-kDa form of γ-ENaC, there was a significant decrease in the abundance of the aldosterone-induced 70-kDa form, suggesting a partial reversal of the effect of aldosterone on γ-ENaC (see Introduction). Interestingly, there was a significant increase in the abundance of the α-1 subunit of the Na-K-ATPase, a change that is unlikely to be related to the escape phenomenon because it would predict an increase in renal tubule Na absorption (see Discussion).
Figure 2Immunoblots assessing Na transporter and Na channel abundances in homogenates from renal cortex (left) and renal outer medulla (right) in control (aldosterone-low Na) and escape (aldosterone-high Na) rats. Each lane was loaded with a sample from a different rat. Table 2 summarizes densitometric analysis.
In outer medulla, the pattern of responses for NHE3, NKCC2, and the three subunits of ENaC was the same as in the cortex (Figure 2; Table 2). (Note that NCC and NaPi-2 are not expressed in inner stripe of outer medulla and therefore were not examined by immunoblotting in outer medullary samples.) In contrast to the cortex, Na-K-ATPase α-1 subunit abundance was not increased in outer medulla.
Figure 3 shows immunoblots for NCC and α-ENaC in whole-kidney samples from the same rats as shown in Figure 2. Once more, there was a marked suppression of the abundance of NCC (as seen in the cortical samples) to 25% ± 7% of the mean for the control rats, whereas α-ENaC abundance was not decreased (Table 2).
Figure 3Immunoblots assessing NCC and α-ENaC abundances in homogenates from whole kidney in control (aldosterone/low Na) and escape (aldosterone/high Na) rats. Densitometric analysis revealed that only NCC abundance was decreased.
Immunocytochemistry. Figure 4 shows immunofluorescence labeling of NCC in the renal cortex of control and aldosterone-escape rats. Figure 4a shows double labeling using antibodies to the thiazide-sensitive cotransporter NCC (green) and the Na-K-ATPase (red). Labeling conditions and exposure settings on the confocal microscope were identical for the two images. As can be seen, the anti-NCC antibody labeled exclusively the apical membrane region of a subset of tubules recognized to be DCTs (21). The NCC labeling was decreased in the DCT cells from aldosterone-escape rats. The Na-K-ATPase labeling marks the basolateral plasma membrane domains in DCTs and other segments. The Na-K-ATPase labeling dominates the basal two thirds of the cells owing to the extensive basal infoldings of the DCT cells. Figure 4b shows a pair of low-power NCC immunofluorescence micrographs from control and aldosterone-escape rats. Labeling conditions and exposure settings on the confocal microscope were again identical for the two images. A relatively uniform decrease in NCC labeling in all DCTs can be appreciated. Similar observations were made in two additional pairs of rats.
Figure 4Confocal images showing immunofluorescence labeling of NCC in rat renal cortex. (a) Double labeling for NCC (green) and Na-K-ATPase α-1 subunit (red). ×250. (b) Low-power images of NCC labeling. ×40.
Figure 5 shows similar sections labeled for NCC using immunoperoxidase techniques. The upper two panels show high-power images demonstrating a marked decrease in NCC labeling in aldosterone escape versus control. There was no obvious redistribution of NCC labeling in DCT cells. Lower-power images (bottom panels) show that the decrease in NCC occurs generally throughout the cortex. Similar observations were made in three pairs of rats. Thus, we conclude that the aldosterone-escape process is associated with a marked decrease in the abundance of NCC in the DCT cells, but that there is no marked change in the cellular distribution of NCC in these cells.
Figure 5Immunoperoxidase NCC labeling in rat renal cortex of control and aldosterone-escape rats. Upper panels, ×625; lower panels, ×75.
Time course of aldosterone escape. To determine whether the increase in Na excretion in aldosterone escape correlates with the demonstrated decreases in the abundance of NCC, we carried out time course studies using the protocol shown in Figure 1, and sampled rats at 1, 2, and 4 days after switching from the low-Na to the high-Na diet. Creatinine clearances (in milliliters per minute per 200 grams of BW) for the three time points were: day 1, control 0.83 ± 0.09, escape 0.73 ± 0.04 (NS); day 2, control 0.84 ± 0.07, escape 0.72 ± 0.04 (NS); day 4, control 1.10 ± 0.06, escape 1.08 ± 0.14 (NS). The time course of changes in NCC in renal cortex after increasing Na intake in the aldosterone escape protocol is shown in Figure 6. NCC abundance fell significantly in the first day after increasing Na intake, and the increase in Na excretion correlated with the decrease in NCC abundance.
Figure 6Time course of changes in NCC abundance and Na excretion after increase in NaCl intake in aldosterone-escape experiments. Experiments were conducted following the protocol shown in Figure 1. NCC abundance was assessed a relative band density in immunoblots as described above. *Significant difference relative to 0 day time point (before the increase in Na intake).
NCC mRNA abundance. To assess whether the decrease in NCC protein abundance in the kidneys of aldosterone-escape rats is owing to a decrease in NCC mRNA, we assessed mRNA abundance by RPAs. As shown in the left panel of Figure 7, addition of increasing amounts of renal cortical mRNA results in a progressive increase in band density for both NCC and β-actin, illustrating that the assay is capable of detecting changes in mRNA abundance when loaded with 2–18 μg per tube. The right panel shows an absence of a difference in NCC mRNA levels between control and aldosterone-escape rats. In three such pairs of rats, there was no change in band densities for NCC mRNA (normalized to β-actin): control, 0.27 ± 0.02; aldosterone-escape, 0.27 ± 0.05 (no significant difference). Immunoblots using homogenates from the opposite kidneys of these rats confirmed that NCC protein abundance was markedly decreased (data not shown). Thus, the decrease in NCC abundance in aldosterone escape does not depend on a decrease in NCC mRNA levels.
Figure 7Assessment of NCC mRNA abundance in aldosterone-escape by RPA. Left: A titration series with 2–18 μg of renal cortical total RNA. NCC probe, 389 bp; β-actin probe, 127 bp. Right: Comparison of whole-kidney NCC mRNA abundance for a control rat versus an aldosterone-escape rat using 10 μg total RNA. Densitometric analysis of three pairs showed no significant difference in NCC mRNA between control and aldosterone-escape (see the text).
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