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1Laboratory of Kidney and Electrolyte Metabolism, National Heart, Lung, and Blood Institute, Bethesda, Maryland; and 2The Water and Salt Research Centre, University of Aarhus, Aarhus, Denmark
Submitted 24 July 2006 ; accepted in final form 18 September 2006
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ABSTRACT |
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IMCD; water transport; serine-256; channel; inner medullary
As part of a large-scale phosphoproteomics screen in native rat inner medullary collecting duct (IMCD) cells, we recently identified additional residues in the COOH-terminus of AQP2 that are phosphorylated, namely S261, S264, and S269 (9). In that study, mass spectrometry-based quantification of AQP2 phosphopeptides suggested that the relative abundance of phosphorylated S261 (pS261) decreases in response to short-term vasopressin treatment, although the exact cause of this decrease could not be definitively addressed using this method. Residue S261 is conserved among all identified mammalian AQP2 isoforms.
To further explore S261 phosphorylation in the context of vasopressin signaling, we have developed a new phospho-specific antibody to this site. The present study addresses initial antibody characterization as well as cellular and subcellular distributions of pS261 in various distal tubule segments from rat and mouse kidney. Finally, potential changes in S261 phosphorylation with short-term [deamino-Cys1,D-Arg8]vasopressin (dDAVP) exposure were addressed using both in vitro and in vivo methods.
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METHODS |
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Protocol 1: isolated IMCD experiment. Pathogen-free male Sprague-Dawley rats (Taconic Farm, Germantown, NY) were maintained on an autoclaved pelleted rodent chow (413110–75-56, Zeigler Bros., Gardners, PA) and ad libitum drinking water. All experiments were conducted in accord with an animal protocol approved by the Animal Care and Use Committee of the National Heart, Lung, and Blood Institute (ACUC protocol number H-0110).
Protocol 2: normal Wistar rats and CD-specific AQP2 knockout mice. The following animal protocols have been approved by the boards of the Institute of Anatomy and Institute of Clinical Medicine, University of Aarhus, according to the licenses for use of experimental animals issued by the Danish Ministry of Justice. Wistar rats were obtained from Møllegaard Breeding Center (Ejby, Denmark). CD-specific knockout mice were from an in-house stock and were originally generated by Rojek et al. (19). Rats were maintained on a standard rodent diet (Altromin no. 1320, Chr. Petersen A/S, Ringsted) and had free access to water.
Protocol 3: short-term dDAVP infusion of Brattleboro rats. Homozygous Brattleboro rats were obtained from Harlan Nederland (Horst, The Netherlands). Before the experiment, rats had free access to standard rat chow (Altromin no. 1320, Chr. Petersen A/S). Six Brattleboro rats were treated with subcutaneous injections of 50 ng dDAVP (Sigma-Aldrich, catalog no. V1005) in 500 µl saline/animal, and six vehicle-injected Brattleboro rats served as controls. After 2 h, the rats were anesthetized by halothane and were killed during perfusion fixation of the kidneys. Between injection of dDAVP and fixation of the kidney, rats had free access to water but not food. This is the same protocol as previously described by Christensen et al. (3).
Materials
Synthetic, NH2-terminally biotinylated peptides corresponding to amino acids 241–271 of the COOH-terminus of rat AQP2 (..EPDTDWEEREVRRRQSVELHSPQSLPRGSKA) were generated by Anaspec (San Jose, CA). For three of these peptides, various residues were modified by phosphorylation, including S256, S261, and doubly-phosphorylated S256 and S261. An affinity-purified rabbit polyclonal antibody (anti-pS261) was generated against a proprietary sequence from the COOH-terminus of rat AQP2 that included pS261 (PhosphoSolutions, Aurora, CA). The affinity-purified antibody anti-pS256, which recognizes AQP2 phosphorylated at Ser-256, has been described previously (17). Total AQP2 antibodies L127 (16) and chicken anti-AQP2 (1) were used for immunoblotting and immunofluorescence, respectively. Goat polyclonal antibodies to glucose-regulated protein 78 (GRP78) and cathepsin D were from Santa Cruz Biotechnology (Santa Cruz, CA). The Golgi matrix protein 130 (GM130) mouse monoclonal antibody was from BD Transduction Laboratories (San Jose, CA). FITC-conjugated Dolichos biflorus agglutinin (DBA) was from Sigma.
IMCD Isolation and Treatment with dDAVP
IMCD fractions were freshly prepared from the inner medulla of rat kidney using the method of Stokes et al. (20) with some modifications (10, 18). IMCD samples were incubated in bicarbonate buffer in the presence or absence of 1 nM dDAVP for 1, 2, 5, 15, or 30 min in a pH/temperature-controlled chamber with gentle mixing under an atmosphere of 95% air–5% CO2 at 37°C.
Immunoblotting and Dot Blotting
Enriched IMCD pellets were solubilzed in 1.5% SDS/Tris, pH 6.8. Total protein concentrations were determined by the bicinchoninic acid assay (Pierce) using BSA as the standard. Samples were then diluted in Laemmli buffer (10 mM Tris, pH 6.8, 1.5% SDS, 6% glycerol, 0.05% bromophenol blue, and 40 mM DTT). Protein samples (10 µg) were subjected to SDS-PAGE and immunoblotting as described (4), with reagent and protocol modifications based on the Odyssey Infrared Imaging System (LiCor, Lincoln, NE). Affinity-purified anti-pS261 and anti-pS256 were used at 1:2,000 and 1:500, respectively, for both dot blot and immunoblot analysis. A fluorescently labeled secondary antibody IRDye 680 goat anti-rabbit IgG (LiCor) was used at 1:5,000 for detection of all primary antibodies. Band densities for both pS256 and pS261 were initially normalized to the total AQP2 signal, followed by calculation of the normalized abundance ratio (dDAVP vs. control) for each individual trial for each time point. These ratios were then converted to log2 values followed by calculation of the mean ± SE (n = 3) for each time point. Paired t-tests were performed as appropriate.
Immunohistochemistry
Kidneys were perfusion fixed through the abdominal aorta. Blood was flushed from the kidneys with cold 0.01 M PBS (pH 7.4) for 15 s, before switching to cold 3% paraformaldehyde in 0.1 M cacodylate buffer (pH 7.4) for 3 min. The kidney was removed, and the midregion was sectioned into 2- to 3-mm transverse sections and immersion fixed for an additional 1 h, followed by 3 x 10-min washes with 0.1 M cacodylate buffer, pH 7.4. The tissue was dehydrated in graded ethanol and left overnight in xylene. After tissue embedding in paraffin, 2-µm sections were cut on a rotary microtome (Leica Microsystems A/S, Herlev, Denmark).
For immunolabeling, the sections were dewaxed with xylene and rehydrated with graded ethanol before endogenous peroxidase activity was blocked by 0.5% H2O2 in absolute methanol for 10 min. Sections were microwave boiled in a target retrieval solution (1 mM Tris, pH 9.0, with 0.5 mM EGTA) for 10 min, before nonspecific binding was blocked with 50 mM NH4Cl in PBS for 30 min, and 3 x 10 min blocking with PBS blocking buffer (1% BSA, 0.05% saponin, and 0.2% gelatin). The sections were incubated with primary antibody (diluted in PBS with 0.1% BSA and 0.3% Triton X-100) overnight at 4°C. The sections were washed 3 x 10 min with PBS wash buffer containing 0.1% BSA, 0.05% saponin, and 0.2% gelatin and incubated with horseradish peroxidase-conjugated secondary antibody (goat anti-rabbit immunoglobulin, DAKO P448, DAKO A/S, Glostrup, Denmark) for 1 h at room temperature. After 3 x 10-min rinses with PBS wash buffer, the sites of antibody-antigen reaction were visualized with a brown chromogen produced within 10 min by incubation with 0.05% 3,3'-diaminobenzidine tetrachloride (Kem-en Tek, Copenhagen, Denmark) dissolved in distilled water with 0.1% H2O2. Mayer's hematoxylin was used for counterstaining, and, after dehydration, cover slips were mounted with hydrophobic medium (Eukitt, O. Kindler, Freiburg, Germany). For sections prepared for immunofluorescence, a secondary fluorescent antibody was used (goat anti-rabbit IgG, Alexa Fluor 488 11008, and goat anti-chicken IgG Alexa Fluor, 546 A11040 [GenBank] , Molecular Probes, Eugene, OR). After a 1-h incubation at room temperature, cover slips were mounted with a hydrophilic mounting media containing anti-fading reagent (n-propyl-gallat, P-3101, Sigma Chemical, St. Louis, MO). Light microscopy was carried out on a Leica DMRE (Leica Microsystems A/S, Herlev, Denmark). Laser confocal microscopy was carried out on a Leica TCS-SP2 laser confocal microscope (Heidelberg, Germany).
Assessment of Colocalization
To quantify the degree of colocalization between phosphorylated and total forms of AQP2, images were obtained with a Zeiss 510 confocal system equipped with UV-Vis lasers (Carl Zeiss, Jena, Germany). Images were acquired sequentially using a 364-nm laser line and emission between 385 and 470 nm for 4,6-diamidino-2-phenylindole, a 488-nm laser line and emission between 505 and 550 nm for Alexa 488, and a 561-nm laser line and emission over 585 nm for Alexa 568. Z-series of high-resolution (100 nm/pixel) images were obtained throughout the cells with a x63, 1.2-numerical aperture C-Apochromat water immersion objective. The three-dimensional (3D) data sets were used for 3D reconstructions and analyzed for colocalization. Quantification of colocalization was carried out using the colocalization module of Imaris 5.0 software package (Bitplane AG, Zurich, Switzerland). Pixel codistribution was calculated for green (pS256 or pS261) and red (total AQP2) channels throughout the 3D data sets. Two-dimensional histograms (fluorograms) show the distribution of pixel intensities. The percentage of colocalized material (intensity and volume) and the Pearson correlation coefficient in the colocalized volume (Rcoloc; 1 = perfect correlation, 0 = no correlation, and –1 = perfect inverse correlation) were compared.
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RESULTS |
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TOPABSTRACTMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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Three synthetic COOH-terminal AQP2 phosphopeptides (pS256, pS261, and pS256/261), as well as an unphosphorylated AQP2 peptide (see METHODS for sequence information), were spotted onto a nitrocellulose membrane and probed with anti-pS256, anti-pS261, and anti-total AQP2 antibodies (Fig. 1A). Anti-pS256 recognized AQP2 peptide phosphorylated at S256, but did not cross-react with either peptide phosphorylated at S261 or with unphosphorylated peptide. Anti-pS261 recognized AQP2 phosphorylated at S261, but did not cross-react with either peptide phosphorylated at S256 or unphosphorylated peptide. Both phospho-specific antibodies recognized the doubly phosphorylated AQP2 peptide. As expected, total AQP2 antibody recognized unphosphorylated as well as all phosphorylated peptides.
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Immunolabeling of AQP2 pS261 in Mouse and Rat
To further characterize the specificity of the anti-pS261 antibody, immunoperoxidase labeling of kidney tissue sections from CD-specific AQP2 knockout mice (19) was performed. Strong immunolabeling was present in the IMCD of wild-type control mice (Fig. 2A), while a general absence of labeling was observed in the IMCD of knockout mice (Fig. 2B, note that a few cells were positive for pS261 in the knockout mice due to incomplete expression of the Cre-recombinase).
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To better define the subcellular distribution of AQP2-pS261 in rat collecting duct, double labeling was performed for AQP2 phosphorylated at pS261 and for various subcellular markers: DBA for plasma membrane, GRP78 for endoplasmic reticulum (ER), GM130 for Golgi, and cathepsin D for lysosomes (Fig. 6). In all instances, results were similar for both IMCD and OMCD. DBA (Fig. 6A) exhibited strong apical labeling, which did not coincide with pS261, which was predominantly intracellular. GM130 (Fig. 6C) staining was punctate but also did not colocalize with pS261. Both GRP78 (Fig. 6B) and cathepsin D (Fig. 6D) labeling were extensively cytoplasmic; however, there was minimal colocalization with pS261. These results suggest that the majority of intracellular pS261 resides in a location other than the ER, Golgi, and lysosomes.
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To determine the effects of short-term vasopressin on S261 phosphorylation, rat IMCD suspensions were enriched by differential centrifugation from freshly isolated whole inner medullas (according to METHODS) and incubated in the presence or absence of 1 nM dDAVP for 30 min. Proteins were immediately solubilized and subjected to immunoblotting using antibodies to pS256, pS261, and total AQP2 (Fig. 7A). All antibodies, including anti-pS261, recognized a discrete band at 
29 kDa, consistent with the size of the unglycosylated AQP2 monomer, as well as a broad smear centered at 37 kDa, corresponding to various glycosylated forms of AQP2. S256 phosphorylation increased in samples treated with dDAVP for 30 min (Fig. 7A, left), as previously demonstrated (9, 21). In contrast, levels of both glycosylated and unglycosylated pS261 decreased in response to short-term dDAVP (Fig. 7A, middle). Neither of these effects was due to a change in overall AQP2 protein abundance (Fig. 7A, right). To further explore these changes in phosphorylation and to obtain quantitative data, a time course study was performed. Isolated rat IMCD samples were incubated with dDAVP as described above, and proteins were harvested at five different time points (1, 2, 5, 15, and 30 min) for immunoblot analysis. To provide a clearer visual representation, these ratios were converted to log2 values. After only 1 min of dDAVP treatment, pS256 increased approximately twofold (log2 = 1.17 ± 0.4) and remained elevated out to 30 min (Fig. 7B). The level of pS261 began to decrease by 15 min and was significantly reduced 2.7-fold (log2 = –1.57 ± 0.77) by 30 min.
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To investigate whether the decrease in pS261 expression after short-term dDAVP treatment could be recapitulated in vivo, we carried out immunoperoxidase labeling of pS261 in kidney tissue sections from Brattleboro rats treated with dDAVP for 2 h. Immunolabeling of pS261 AQP2 was markedly decreased in all kidney regions. In the initial part of the inner medulla, the labeling was virtually absent compared with untreated Brattleboro rats, which showed mainly dispersed intracellular labeling (Fig. 8, A and B). In the middle of the inner medulla, the reduction in pS261 abundance with dDAVP treatment was detectable but less prominent than in the initial part of the inner medulla (Fig. 8, C and D), and a faint labeling in the apical and subapical domain was observed after 2 h of dDAVP (Fig. 8D, bottom inset). Some IMCD cells in the middle of the inner medulla also showed large granules in the subnuclear region (Fig. 8D, top inset). Initial IMCD cells from the same animals labeled with an anti-pS256 antibody showed an increase in labeling of the apical plasma membrane domain and less intracellular labeling compared with the untreated Brattleboro rats (Fig. 8, E and F).
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DISCUSSION |
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TOPABSTRACTMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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Immunohistochemical analysis revealed that AQP2 phosphorylated at S261 was present in cells of the CNT as well as in principal cells of all collecting duct segments from CCD to terminal IMCD. In normal rats with free access to water, the subcellular distribution of pS261-AQP2 varied, depending on the tubule segment, being apical and basolateral in CNT; mostly subapical in CCD, OMCD, and the initial part of the IMCD; and broadly distributed in cytoplasm in middle IMCD. In CCD, OMCD, and initial IMCD, pS261 appeared to have a distinct distribution compared with pS256. While pS261 staining was mostly subapical and punctate in appearance, pS256 staining was strongly apical with relatively little cytoplasmic labeling. Quantification of colocalization demonstrated that the pS256 signal overlaps significantly more than pS261 with total AQP2. The difference in distribution of pS261 and pS256 indicates the presence of distinct subcellular pools of AQP2 and suggests that phosphorylation at S261 may be involved in regulation of the subcellular localization of AQP2. An analysis of the subcellular distribution of AQP2-pS261 with various markers demonstrated that the majority of cytoplasmic pS261 is not located in the ER, Golgi, or lysosomes. Other candidate structures in which pS261 could reside might include early, late, or recycling endosomes or secretory vesicles.
The hypothesis that pS256 and pS261 play divergent roles in regulation of AQP2 is supported by evidence obtained in a previous study using protein mass spectrometry (9). In that study, we utilized quantitative phosphoproteomics to analyze changes in phosphoprotein abundance with short-term vasopressin exposure and found that the abundance of AQP2 monophosphorylated at S256 increased 7.2-fold, while the abundance of AQP2 monophosphorylated at S261 decreased 2.5-fold in the presence of 1 nM dDAVP for 10 min. However, this prior study could not clarify whether the decrease in monophosphorylated AQP2 at S261 was due to phosphorylation at S256 to produce the diphosphorylated form or due to an absolute reduction of AQP2 phosphorylated at S261. In the present study, immunoblotting with the pS261 antibody showed that pS261 was significantly decreased only after 30 min of dDAVP exposure, much later than the increase in pS256 (2 min), demonstrating an absolute reduction in AQP2 phosphoryated at S261. The decrease in pS261 detected within 10 min of dDAVP exposure in the previous study was likely due to formation of diphosphorylated AQP2, which effectively decreased the mono-pS261 peptide signal during analysis by mass spectrometry. Thus both mechanisms appear to be in play.
In the present study, we were also able to detect a decrease in overall pS261 abundance in tissue sections from Brattleboro rats treated with dDAVP for 2 h. While there was nearly a complete absence of pS261 labeling in the initial part of the IMCD, there appeared to be a small amount of apical labeling of pS261 in the middle IMCD, suggesting that pS261 is not always located intracellularly. These sections also contained a number of larger intracellular structures that stained positive for pS261 (Fig. 8D, inset arrows). The identity of these structures remains to be determined. To be consistent with this localization, it can be speculated that phosphorylation at S261 either inhibits exocytosis or stimulates endocytosis.
While it is evident that there is a decrease in overall pS261 abundance with short-term vasopressin exposure, the exact cause of this decrease (e.g., increased phosphatase activity, decreased kinase activity, or protein degradation) is as yet unidentified. If the mechanism involves dephosphorylation of S261, a candidate phosphatase would be protein phosphatase 2B (calcineurin), which has been found in endosomes containing AQP2 and can dephosphorylate AQP2 in vitro (11). In addition, calcineurin A
null mice were shown to have diminished apical expression of AQP2 in the presence of dDAVP (7). Although the authors suggested that this phenotype was due to a combination of defective ER to Golgi transport and decreased AQP2 S256 phosphorylation, it may be informative to explore phosphorylation of S261 in the context of short-term vasopressin in these animals. If regulation of a specific kinase is involved, we can speculate that it is a member of the so-called "proline-directed" serine/threonine kinase subfamily because of the presence of proline at position 262. This proline is mutated in a subset of patients with autosomal recessive nephrogenic diabetes insipidus (5). Members of the proline-directed protein kinase family include mitogen-activated protein kinases (ERK1, p38), as well as cyclin-dependent kinases (cdc2, Cdk5). Upon sequence comparison, there is no similarity between the proline-directed S261 consensus and the basophilic PKA consensus of S256 (..RRQS..), further evidence that phosphorylation at these two residues may be regulated by distinct signaling pathways. There is also the possibility that the decrease in pS261 abundance results from enhanced protein degradation by lysosomal and/or proteasomal pathways. Evidence suggests that both of these pathways regulate the transient decrease in AQP2 abundance following increased vasopressin exposure in a cortical collecting duct cell line (8).
In conclusion, this study presents the first direct evidence demonstrating regulation of AQP2 phosphorylation at S261 by vasopressin. It also shows that the intracellular distribution of pS261-AQP2 differs from that of total AQP2 as well as pS256-AQP2, suggesting a unique role in AQP2 trafficking or function. Although the signaling pathways regulating AQP2 phosphorylation at S261 remain unknown, the findings in this paper are tantalizing clues to guide future studies of regulation of water transport in the renal collecting duct.
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GRANTS |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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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.
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