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PKD1 Haploinsufficiency Causes a Syndrome of Inappropriate Antidiuresis in Mice

Ali K. Ahrabi^*, Sara Terryn, Giovanna Valenti, Nathalie Caron, Claudine Serradeil-Le Gal^||, Danielle Raufaste^||, Soren Nielsen^¶, Shigeo Horie^**, Jean-Marc Verbavatz and Olivier Devuyst^*

^* Division of Nephrology, Université catholique de Louvain Medical School, Brussels, Belgium; Laboratory of Cell Physiology, Center for Environmental Sciences, Hasselt University, Diepenbeek, Belgium; Department of Physiology, University of Bari, Bari, Italy; Department of Physiology and Pharmacology, University of Mons-Hainaut, Mons, Belgium; ^|| Sanofi-Aventis, Toulouse, France; ^¶ The Water and Salt Research Center, University of Aarhus, Aarhus, Denmark; ^** Department of Urology, Teikyo University, Tokyo, Japan; and Cell and Molecular Imaging, CEA/Saclay, Gif-sur-Yvette, France

Address correspondence to: Dr. Olivier Devuyst, Division of Nephrology, UCL Medical School, 10 Avenue Hippocrate, B-1200 Brussels, Belgium. Phone: +32-2-764-5450; Fax: +32-2-764-5455; E-mail: devuyst{at}nefr.ucl.ac.be

Received for publication January 18, 2006. Accepted for publication March 9, 2007.

	Abstract

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Mutations in PKD1 are associated with autosomal dominant polycystic kidney disease. Studies in mouse models suggest that the vasopressin (AVP) V2 receptor (V2R) pathway is involved in renal cyst progression, but potential changes before cystogenesis are unknown. This study used a noncystic mouse model to investigate the effect of Pkd1 haploinsufficiency on water handling and AVP signaling in the collecting duct (CD). In comparison with wild-type littermates, Pkd1^+/– mice showed inappropriate antidiuresis with higher urine osmolality and lower plasma osmolality at baseline, despite similar renal function and water intake. The Pkd1^+/– mice had a decreased aquaretic response to both a water load and a selective V2R antagonist, despite similar V2R distribution and affinity. They showed an inappropriate expression of AVP in brain, irrespective of the hypo-osmolality. The cAMP levels in kidney and urine were unchanged, as were the mRNA levels of aquaporin-2 (AQP2), V2R, and cAMP-dependent mediators in kidney. However, the (Ser256) phosphorylated AQP2 was upregulated in Pkd1^+/– kidneys, with AQP2 recruitment to the apical plasma membrane of CD principal cells. The basal intracellular Ca²⁺ concentration was significantly lower in isolated Pkd1^+/– CD, with downregulated phosphorylated extracellular signal–regulated kinase 1/2 and decreased RhoA activity. Thus, in absence of cystic changes, reduced Pkd1 gene dosage is associated with a syndrome of inappropriate antidiuresis (positive water balance) reflecting decreased intracellular Ca²⁺ concentration, decreased activity of RhoA, recruitment of AQP2 in the CD, and inappropriate expression of AVP in the brain. These data give new insights in the potential roles of polycystin-1 in the AVP and Ca²⁺ signaling and the trafficking of AQP2 in the CD.

	Introduction

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Autosomal dominant polycystic kidney disease (ADPKD) is the most frequent inherited nephropathy and an important cause of ESRD (1). Mutations in two genes, PKD1 and PKD2, have been associated with ADPKD. Mutations in PKD1 account for approximately 85% of the affected families, and they are associated with a renal disease that progresses more rapidly than in families with PKD2 (2). PKD1 and PKD2 encode integral membrane proteins, polycystin-1 and polycystin-2, that interact in renal primary cilia and regulate the proliferation and differentiation of renal tubular cells via different signaling pathways (3). Mutations in PKD1/PKD2 disrupt these pathways, leading to cystogenesis by a combination of increased cellular proliferation, abnormal fluid secretion, and dedifferentiation (1,3). All nephron segments may be involved in cyst formation in ADPKD, but an important fraction of the cysts is derived from the collecting ducts (CD) (4,5).

In vitro studies have shown that cAMP plays a major role in cystogenesis. Exposure to cAMP agonists stimulates fluid secretion across monolayers of ADPKD cyst-lining epithelial cells (6), as well as the proliferation of these cells (7). Furthermore, increased levels of cAMP resulting from the activation of vasopressin (AVP) V2 receptor (V2R) pathway in CD cells may contribute to the progression of cystogenesis. In two cystic models that are orthologous to human autosomal recessive PKD (PCK rat) and nephronophthisis (pcy mouse) and one cystic model that is orthologous to human ADPKD type 2 (Pkd2^–/tm1Som mouse), increased renal cAMP levels compared with normal mice, paralleled with higher expression of aquaporin-2 (AQP2) and V2R, have been reported (8–10). The administration of V2R antagonists to these models lowered renal cAMP and inhibited the development and progression of established renal cystic disease (8–10), motivating trials to test the efficacy of V2R antagonists in patients with ADPKD (11). It is important to note that all rodent models tested so far develop renal cysts (and subsequent renal failure) within a few weeks of age.

In normal CD cells, the stimulation of V2R by AVP leads to the phosphorylation of AQP2 on the Ser256 residue and its subsequent insertion in the apical plasma membrane, an essential step to mediate final urine concentration (12,13). A mild impairment in urinary concentrating ability, with increased circulating AVP levels, has been described in patients with ADPKD and cystic kidneys (11,14,15). However, this urinary concentrating abnormality is probably not specific, because any modification of the medullary architecture (e.g., cystic changes) impairs the constitution of the corticomedullary osmotic gradient, resulting in nephrogenic diabetes insipidus (16). Considering that an activation of the V2R pathway has been involved in PKD mouse models with cysts originating from the CD (8–10), we hypothesized that the complex chain of events that mediates urinary concentration in the CD could be modified early, before cystogenesis.

In this study, we used a well-established mouse model with a targeted deletion of Pkd1 (Pkd1^+/–) (17) to test whether Pkd1 haploinsufficiency causes abnormal water handling and AVP signaling in the CD before cystogenesis and renal failure. Like other Pkd1-null mutants, the homozygous Pkd1^–/– mice die in utero with massive cystic kidneys, hydrops fetalis, and cardiovascular defects (17–20). By contrast, there is no consistent phenotype in heterozygous Pkd1^+/– mice that do not develop renal cysts until (in a few individuals) a very old age (21,22). Our investigations reveal for the first time that reduced Pkd1 gene dosage results in inappropriate antidiuresis and positive water balance, reflecting decreased intracellular calcium ([Ca²⁺]_i) levels with lowered RhoA activity and recruitment of AQP2 in the apical membrane of CD principal cells and inappropriate expression of AVP in the brain.

	Materials and Methods

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Pkd1 Mice and Sampling
Experiments were conducted on age- and gender-matched adult mice (aged 20 to 35 wk) with a targeted deletion of the exons 2 to 5 and part of exon 6 of Pkd1, resulting in a null allele (17). The mice were maintained on a mixed 129/sv/C57BL/6J background. They were housed in light- and temperature-controlled room with ad libitum access to tap water and standard chow (Pavan, Oud-Turnhout, Belgium). Previous experiments showed that the Pkd1 mice had similar heart rate and BP (N. Morel, et al., unpublished data, 2007). Water handling at baseline and during various protocols was assessed in individual metabolic cages, after appropriate training. Blood and tissue samples were obtained at time of killing, after anesthesia with Sevoflurane (Abbott, Ottignies, Belgium) and exsanguination. Blood was collected by venous puncture, and plasma samples were kept at –20°C. The sampling procedures were exactly similar in both groups. Tissue samples were immediately processed for fixation and mRNA/protein extraction. The experiments were conducted in accordance with the National Research Council Guide for the Care and Use of Laboratory Animals and were approved by the local Animal Ethics Committee.

Water Handling Protocols
Plasma samples and 24-h urine collections were obtained at baseline, and the urinary concentrating ability was tested after 24-h water deprivation. The capacity to excrete a water load was tested after intraperitoneal injection of 2 ml of sterile water; urine was collected under a plastic-wrapped container on an hourly basis for the next 6 h. The aquaretic effect of the V2R antagonist SR121463B (Sanofi-Aventis, Chilly-Mazarin, France), which has a high affinity for renal V2R from several species, including rat, mouse, and human (K_i = 0.26 ± 0.04 nM) (23), was tested after intraperitoneal administration of dosages that ranged from 0.1 to 30 mg/kg and hourly determination of diuresis for the next 6 h as described above.

V2R Binding Assays and Autoradiography
Renomedullary preparations from Pkd1^+/+ and Pkd1^+/– mice (or CHO membranes that expressed human V2R used as positive controls) were incubated in a 50-mM Tris-HCl buffer (pH 8.1) that contained 2 mM MgCl₂, 1 mM EDTA, 0.1% BSA, 0.1% bacitracin, and [³H]SR121463 (0.8 to 28 nM for saturation experiments or 2 nM for binding studies). The reaction was started by the addition of membranes (7.5 µg/assay for CHO and 100 to 130 µg/assay for mouse renal tissue) and incubation for 45 min at 25°C, stopped by filtration through Whatman GF/B filters as described previously (24). Nonspecific binding was determined in the presence of 1 µM SR121463B. Data for equilibrium binding (apparent equilibrium dissociation constant [K_d] and maximum binding density [B_max]) were calculated using an interactive nonlinear regression program (25).

For performance of autoradiography, kidneys from Pkd1 mice were frozen at –40°C in isopentane and further stored at –80°C. Serial sections (15 µm) were mounted onto gelatin chrome-alum slides, rinsed to eliminate endogenous AVP, and incubated with 1.5 nM [³H]SR121463 alone (total binding) and in the presence of 1 µM unlabeled SR121463B or AVP (nonspecific binding) as described previously (24). After incubation, the sections were washed three times for 10 min each in ice-cold binding buffer, dipped in distilled water, and dried under a stream of cold air. Rinsed labeled sections were placed on a phosphor-imaging plate for 4 d and further analyzed with a BAS5000 Bio-Image Analyser (Fuji, Tokyo, Japan). SR121463B, monophosphate salt, and [³H]SR121463 (47.5 Ci/mmol) were synthesized at Sanofi-Aventis, whereas AVP was obtained from Sigma Chemical Co. (L'Isle d'Abeau, France).

Plasma and Urine Analyses
Sodium, urea, creatinine, and calcium were measured using a Kodak Ektachem DT60II Analyzer (Johnson & Johnson, New Brunswick, NJ), and osmolality was measured using a Fiske Osmometer (Needham Heights, MA). The nitrite/nitrate (NOx) concentrations were measured in urine and plasma using a colorimetric assay (Cayman Chemical, Ann Arbor, MI). Because sevoflurane may induce the release of AVP, thereby increasing plasma values in our protocols, we measured urine AVP levels, which were not obtained under sevoflurane anesthesia, using RIA (Peninsula Laboratories, San Carlos, CA). For cAMP determinations, whole kidneys were ground under liquid nitrogen and homogenized in 10 volumes of 0.1 M HCl. The homogenate was centrifuged at 600 x g for 10 min, and the supernatant was collected, diluted (1:10) in 0.1 M HCl, and processed with acetylation using an enzyme immunoassay kit (Sigma-Aldrich, St. Louis, MO). The urine samples were diluted (1:5000) in 0.1 M HCl and were processed without acetylation. The urinary prostaglandin E₂ (PGE₂) was measured by EIA (Amersham Biosciences, Piscataway, NJ).

Reverse Transcription–PCR and Real-Time Reverse Transcription–PCR
Total RNA from mouse kidney and brain (26) was extracted with Trizol (Invitrogen, Merelbeke, Belgium), treated with DNase I, and reverse-transcribed into cDNA. The primers (Supplementary Table 1) were designed using Beacon Designer 2.0 (Premier Biosoft International, Palo Alto, CA). Changes in target gene mRNA levels were determined by semiquantitative real-time reverse transcription–PCR (RT-PCR) with an iCycler IQ System (Bio-Rad Laboratories, Hercules, CA) using SYBR Green I. Real-time semiquantitative PCR analyses were performed in duplicate as described previously (27). The PCR conditions were 94°C for 3 min followed by 31 cycles of 30 s at 95°C, 30 s at 61°C and 1 min at 72°C. Negative controls excluded amplification from genomic DNA. For each assay, standard curves were prepared by serial four-fold dilutions of cDNA samples. The efficiency of the reactions was calculated from the slope of the standard curve [efficiency = (10^–1/slope) – 1] (27).

Antibodies
Rabbit polyclonal antibodies against AQP2 (Sigma-Aldrich), Ser256 phosphorylated AQP2 (p-AQP2) (12), AQP1 (Chemicon, Temecula, CA), AQP3 (a gift from J.-M. Verbavatz, CEA Saclay), extracellular signal–regulated kinase 1/2 (ERK1/2; C16) and Tyr204 p-ERK1/2 (Santa Cruz Biotechnologies, Santa Cruz, CA), mouse monoclonal RhoA and Ser188 p-RhoA (Santa Cruz Biotechnologies), and mouse mAb against -actin (Sigma-Aldrich) were used.

Immunoblotting
Kidneys were ground under liquid nitrogen and homogenized as described previously (27). The homogenate was centrifuged at 1000 x g for 15 min at 4°C. The resulting supernatant was either kept at –80°C (as the "total extract" fraction) or centrifuged at 100,000 x g for 120 min at 4°C. The pellet ("membrane" fraction) was suspended in homogenization buffer before determination of protein concentration and storage at –80°C. SDS-PAGE was performed under reduced (kidney) or nonreduced (urine) conditions. After blotting on nitrocellulose, the membranes were incubated overnight at 4°C with primary antibodies, washed, incubated for 1 h at room temperature with peroxidase-labeled antibodies (Dako, Glostrup, Denmark), and visualized with enhanced chemiluminescence. Normalization for -actin was obtained after stripping and reprobing. Densitometry analysis was performed with a StudioStar Scanner (Agfa-Gevaert, Mortsel, Belgium) using the NIH-Image V1–57 software.

Immunohistochemistry
Kidney samples were fixed in 4% paraformaldehyde (Boehringer Ingelheim, Heidelberg, Germany) in 0.1 mol/L phosphate buffer (pH 7.4) before embedding in paraffin. The 6-µm sections were stained with hemalum-eosin or incubated for 30 min with 0.3% H₂O₂, followed by 20 min with 10% normal serum, and 45 min with the primary antibodies diluted in PBS that contained 2% BSA. After washing, sections were successively incubated with biotinylated secondary anti-IgG antibodies, avidin-biotin peroxidase, and aminoethylcarbazole (Vectastain Elite; Vector Laboratories, Burlingame, CA). Sections were viewed under a Leica DMR coupled to a Leica DC300 digital camera (Leica, Heerbrugg, Switzerland).

Immunoelectron Microscopy
For electron microscopy, kidney samples from Pkd1 mice were fixed overnight in 4% paraformaldehyde and 0.1% glutaraldehyde in PBS and washed in PBS. Small samples, including the outer medulla and the top of inner medulla, were embedded in unicryl, and 80-nm-thick sections were cut. Sections were preincubated in 20 mM Tris buffer (pH 7.5) that contained 0.1% BSA, 0.1% fish gelatin, and 0.05% Tween 20 (buffer-T), followed by a 90-min incubation in the same buffer-T that contained a 1:100 dilution of anti-AQP2 polyclonal antibodies. Sections were washed three times in buffer-T, then incubated in a 1:25 dilution of 10 nm of gold-conjugated secondary antibodies for 45 min. After washing in Tris, sections were stained with uranyl-acetate and lead citrate and photographed on a Philips EM 400 microscope (FEI, Eindhoven, Netherlands). Three samples from three pairs of mice were processed, and at least 10 pictures of outer medullary CD principal cells were randomly taken for each sample. The data are expressed as number of gold particles per micron of apical membrane length. Ultrastructural examination of the vasa recta was performed on three pairs of kidney slices and fixed overnight in 2% glutaraldehyde before washing in PBS.

Measurement of RhoA Activity
Quantification of active RhoA (GTP-bound) was measured by selective affinity precipitation of GTP-Rho (Upstate, Temecula, CA), following the procedure described in detail previously (28,29). The kidneys were ground under liquid nitrogen and homogenized in ice-cold 1x Mg²⁺ lysis/wash buffer according to the manufacturer's instructions (Upstate, Temecula, CA). The homogenates were centrifuged at 15,000 x g for 30 min, and the supernatant of each sample was collected. A total of 30 µg of GST-tagged fusion protein, corresponding to residues 7 to 89 of mouse Rhotekin Rho Binding Domain, bound to glutathione-agarose beads, was added to the supernatant of each sample (500-µl exact) and were rotated overnight at 4°C. Beads were washed three times with Mg²⁺ lysis/wash buffer, and bound proteins were separated by SDS-PAGE and detected by Western blotting using a monoclonal RhoA antibody (28,29).

Measurement of Intracellular Ca²⁺ Concentration
Inner medullary collecting ducts (IMCD) were isolated from collagenase-digested medulla from three pairs of Pkd1^+/+ and Pkd1^+/– mouse kidneys. The tubule segments were seeded onto glass coverslips and examined using an inverted Nikon TMD35 epifluorescence microscope (Analis, Namur, Belgium) in a thermostated chamber at 37°C. The intracellular Ca²⁺ concentration ([Ca²⁺]_i) was measured as described previously (30). Briefly, after measurement of the background signal, isolated tubules were loaded with Fura-2 by incubation with the membrane-permeant acetoxymethyl (AM) ester form of the dye (10 µM) for 1 h at 37°C. Tubules were excited at 340 and 380 nm, and the fluorescence emission was recorded at 510 nm. Data collection time for an image was 2 s. Fura-2 was calibrated in vivo at the end of each experiment, according to the equation derived by [Ca²⁺]_i = K_d R_bf [(r – r_min)/(r_max – r)], where K_d is the dissociation constant of Fura-2 for Ca²⁺ (135 nM), R_bf is the maximum fluorescence intensity as a result of excitation at 380 nm (in the absence of Ca²⁺) divided by the minimum fluorescence intensity at 380 nm (in the presence of saturating Ca²⁺), r is the F₃₄₀/F₃₈₀ fluorescence ratio, r_max and r_min are the F₃₄₀/F₃₈₀ fluorescence ratios in the presence of saturating Ca²⁺ and in the absence of Ca²⁺, respectively. r_max was obtained by permeabilization of the tubules with Ca²⁺ ionophore ionomycin (10 µM), in the presence of 5 mM extracellular Ca²⁺. For obtaining subsequently the minimum ratio r_min, the tubules were exposed to a Ca²⁺ free solution (containing 10 mM EGTA) with 10 µM ionomycin and 1,2-bis(o-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA)-acetoxymethyl (10 µM) to buffer intracellular Ca²⁺.

Statistical Analyses
Data are means ± SEM. LogED₅₀ (the dosage of agonist that provokes 50% of the maximum response) values were calculated by nonlinear curve fitting of the individual concentration-effect curve data (GraphPad, San Diego, CA). Comparisons between groups were performed using two-tailed unpaired t test. Significance level was P < 0.05.

	Results

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Kidney Structure and Baseline Parameters
Macroscopic and histology analyses (Supplementary Figure 1) confirmed the normal kidney structure in heterozygous Pkd1^+/– versus Pkd1^+/+ mice (20 to 35 wk old). In particular, no cysts or tubular dilations were observed in any segment of the Pkd1^+/– kidneys. The baseline clinical and biologic parameters are shown in Table 1. The Pkd1^+/– mice had similar body and kidney weight as wild-type (WT) littermates and similar plasma urea and creatinine values. The Pkd1^+/– mice showed a three-fold decrease in urinary output, associated with a higher urine sodium/osmolality and a lower plasma sodium/osmolality in comparison with WT littermates, despite similar water intake. These observations were confirmed in other sets of adult mice, irrespective of gender. The Pkd1^+/– mice were also characterized by significantly lower cumulative urinary excretion of AVP, calcium, and NOx and a trend for lower urinary PGE₂, whereas the cAMP levels in kidney and urine and the plasma NOx were unchanged. The urinary concentrations of calcium and AVP both were higher in Pkd1^+/– mice, reflecting the net water reabsorption. These data show that, in the absence of cystic changes and renal failure, the Pkd1^+/– mice are in positive water balance, with similar water intake but lower plasma sodium and osmolality.

View larger version (19K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 1. Response to water deprivation and water loading in Pkd1 mice. (A) Urine output was measured during 24-h baseline (BL) and 24-h water deprivation (WD) in four pairs of Pkd1+/+ and Pkd1+/– mice. The Pkd1+/– mice had a significantly lower urine output at BL and after 24-h WD. The WD resulted in a similar weight loss in both groups. (B) Urine osmolality (Uosm) was higher at BL and after WD in the Pkd1+/– group versus Pkd1+/+ group. *P < 0.05, Pkd1+/– versus Pkd1+/+. (C) Nine pairs of mice were administered an intraperitoneal injection of 2 ml of sterile water. In comparison with wild-type (WT) littermates, Pkd1+/– mice showed a significantly delayed ability to excrete water up to 3 h after water load. *P < 0.035, #P < 0.005, Pkd1+/– versus Pkd1+/+.

Characterization of Renal V2R and Response to V2R Antagonist
To characterize the distribution and affinity of V2R, we performed autoradiography of Pkd1^+/+ and Pkd1^+/– kidneys that were incubated with a highly selective V2R ligand, alone or with an excess of unlabeled ligand or AVP (Figure 2A). We observed a dense specific labeling confined in the inner/outer medulla and papilla area, corresponding to the main localization of the V2R in rodent CD. The binding pattern, as well as the binding parameters (K_d and B_max) were similar in both groups. We next tested the aquaretic response of Pkd1 mice to the V2R antagonist SR121463B. Using incremental dosages, we showed a dosage-dependent increase in the aquaretic effect in WT mice, whereas Pkd1^+/– mice showed a decreased sensitivity to the V2R antagonist (lower diuresis during the 6-h period) at dosages that ranged from 0.1 to 10 mg/kg (Figure 2B). The decreased response to SR121463B was confirmed by a right shift of the dosage-response curve (Figure 2C), with Pkd1^+/– mice showing a significantly higher ED50 in comparison with Pkd1^+/+ mice (Log ED50: 0.723 ± 0.03 in Pkd1^+/– versus 0.507 ± 0.07 in Pkd1^+/+; n = 5 pairs; P = 0.02). These data demonstrate that, despite similar distribution and binding parameters of V2R, the Pkd1^+/– mice have a decreased sensitivity to a V2R antagonist.

View larger version (34K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 2. Autoradiography and binding properties of renal V2 receptors and efficacy of the vasopressin (AVP) V2 receptor (V2R) antagonist SR121463B in Pkd1 mice. (A) Autoradiography of Pkd1+/+ and Pkd1+/– kidneys that were incubated with the highly selective V2R ligand [3H]SR121463 alone (1.5 nM, total binding; a and b) and in the presence of 1 µM unlabeled SR121463B (c and d) or AVP (e and f; nonspecific binding). The obtained autoradiograms show a dense specific labeling confined in the inner/outer medulla and papilla area, corresponding to the main localization of the V2R in rodent collecting ducts (CD). Binding studies showed that [3H]SR121463 binds with high affinity to renal V2R. The distribution pattern is similar in Pkd1+/+ and Pkd1+/– kidneys. The saturation binding experiments revealed similar binding parameters of [3H]SR121463 for renal V2R in Pkd1 mice (NS; i.e., apparent equilibrium dissociation constant [Kd] and maximum binding density [Bmax]). (B) Hourly urine output after injection of various dosages (0.1 to 30 mg/kg) of SR121463B in five pairs of mice. In comparison with WT littermates (solid trait), Pkd1+/– mice (dashed trait) showed a systematically lower urine excretion during the 6-h period for dosages that ranged from 0.1 to 10 mg/kg. Each point is the mean of five mice in each group. (C) Dosage-response curves and cumulative ED50 determination. A significant higher ED50 is observed in Pkd1+/– mice versus WT littermates (Log ED50 mean 0.507 ± 0.07 in Pkd1+/+, 0.723 ± 0.03 in Pkd1+/–; n = 5 pairs). The Log ED50 value corresponds to dosage concentrations of 3.2 mg/kg in Pkd1+/+ and 5.3 mg/kg in Pkd1+/– mice. Diuresis values (µl per 6-h period) were significantly lower in Pkd1+/– versus Pkd1+/+ at dosage concentrations of 1, 3, and 10 mg/kg. *P < 0.05.

View larger version (27K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 3. Real-time reverse transcription–PCR quantification of the mRNA expression of mediators that are involved in AVP signaling in the kidney and brain of Pkd1 mice. The mRNA levels were first adjusted to glyceraldehyde-3-phosphate dehydrogenase (GAPDH), then normalized to the WT level set at 100% using the following formula: Ratio = 2 [Ct(GAPDH – Target+/–) – Ct(GAPDH – Target+/+).These normalized values (mean ± SEM) are shown in the right column. In addition to the haploinsufficiency in Pkd1, there was a significant decrease in the expression of endothelin-1 (ET1) in Pkd1+/– kidneys. Nine pairs of kidneys and 11 pairs of brains were analyzed.

View larger version (60K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 4. Expression of aquaporin 1 (AQP1), AQP2, and phosphorylated AQP2 (p-AQP2): Immunoblotting. (A) Representative immunoblots for AQP1, AQP2, and p-AQP2 in homogenate (H) and membrane (M) fractions that were prepared from Pkd1 mouse kidneys. Equal loads (20 µg) were compared, as verified by similar -actin expression. Although there is no difference in AQP2 expression in the H fractions, there is an upregulation of AQP2 and p-AQP2 in the M fractions from Pkd1+/– kidneys. There is no difference in AQP1 expression in these very M fractions. (B) Densitometry analysis (core and glycosylated bands) confirms that there is a significant increase of AQP2 (relative OD 161 ± 2%; P = 0.01) and p-AQP2 (relative OD 214 ± 6%; P = 0.0002) in the M fractions of Pkd1+/– kidneys. (C) Representative immunoblot for AQP2 and p-AQP2 in Pkd1 mouse urine. Samples (15 µl) were loaded and analyzed by Western blot under nonreducing condition. *P < 0.05, Pkd1+/– versus Pkd1+/+.

View larger version (89K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 5. Immunostaining and electron microscopy immunogold labeling for AQP2 in the kidneys of Pkd1 mice. (A) In comparison with Pkd1+/+, there is a strong increase in the apical signal for AQP2 (a and b) and p-AQP2 (c and d) in the kidneys of Pkd1+/– mice. The typical staining for AQP3 (e and f) in the basolateral membrane of the principal cells (PC) is similar in both groups. Several unlabeled CD cells correspond to intercalated cells. Bar = 50 µm. (B) Representative electron microscopy gold labeling of AQP2 in the outer medullary CD of Pkd1+/+ (a) and Pkd1+/– (b) mouse kidney. The labeling was restricted to PC and particularly abundant at the plasma membrane. However, AQP2 labeling was remarkably more intense in Pkd1+/– than in Pkd1+/+ kidneys. IC, intercalated cell; N, nucleus; J, tight junction; L, collecting duct lumen. Bar = 0.5 µm. (C) Morphometric analysis of the density of gold particles at the apical plasma membrane of Pkd1+/+ and Pkd1+/– PC confirmed an approximately two-fold increase (*P < 1.10–6) in the apical AQP2 in the Pkd1+/– versus Pkd1+/+ kidney (number of gold particles per millimeter of membrane length: 11.44 ± 0.87 versus 5.18 ± 0.58). The total number of particles and total membrane length were 725 particles over 140 mm (29 cells from 3 Pkd1+/+ mice) and 1633 particles over 142 mm (27 cells from 3 Pkd1+/– mice).

View larger version (36K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 6. Activity of RhoA and extracellular signal–regulated kinase (ERK) signaling in Pkd1 kidneys: Immunoblotting and affinity precipitation. (A) Representative immunoblots for p-RhoA and RhoA in homogenates from Pkd1 mouse kidneys. Equal loads (20 µg per lane) were compared, as verified by similar -actin expression. After probing for p-RhoA, the membrane was stripped and reprobed for RhoA. The upregulation of p-RhoA in the Pkd1+/– group is reflected by the downregulation of RhoA. Densitometry analysis confirms the significant increase of the p-RhoA over RhoA ratio in Pkd1+/– kidneys versus 100% of the Pkd1+/+ group (749 ± 165%; P < 0.001). (B) Representative immunoblot for quantification of active RhoA (GTP-bound) in Pkd1 mouse kidneys. Equal loads (20 µl per lane) were compared. Affinity precipitation followed by Western blotting confirmed that the amount of active RhoA was decreased in the Pkd1+/– kidneys. Densitometry analysis confirms that there is a significantly decreased amount of active RhoA in Pkd1+/– kidneys (relative OD 35 ± 13%; P = 0.01). (C) Representative immunoblot for phosphorylated and total ERK1/2 in homogenates from Pkd1 mouse kidneys. Equal loads (20 µg per lane) were compared and verified by similar -actin expression. There is a significant decrease of the p-ERK1/2 over ERK1/2 ratio in Pkd1+/– kidneys, as confirmed by densitometry (46 ± 7% versus Pkd1+/+ taken as 100%; P = 0.02).

View larger version (10K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 7. Baseline intracellular Ca2+ concentrations ([Ca2+]i) in inner medullary CD (IMCD) tubules of Pkd1 mice. Different regions (n = 24 per group) from six Pkd1+/+ (filled symbols) and six Pkd1+/– (open symbols) IMCD tubules that originated from three pairs of mice were analyzed. [Ca2+]i were lower in the Pkd1+/– group (open symbols) compared with WT group (filled symbols; 146 ± 3.0 versus 186 ± 3.5 nM, respectively; P < 0.0001).

	Discussion

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In contrast with the Pkd1-null mouse models, which are embryonically lethal, heterozygous Pkd1 mice have a normal growth and no detectable abnormalities at birth and during adulthood (17–21). With the exception of a limited number of renal and liver cysts in a minority of old mice (17,21,22), no detailed functional phenotype has been associated with Pkd1 haploinsufficiency. Recent studies in cystic mouse (Pkd2^–/tm1Somand pcy) and rat (PCK) models with various degree of renal failure pointed out that increased cAMP levels, secondary to abnormal V2R signaling in CD cells, could play a role in cyst progression (8–10). However, the effects of such an abnormal signaling could be masked by nonspecific structural changes that alter the osmotic water handling by the CD (16). Thus, adult Pkd1^+/– mice offer the opportunity to test the functional consequences of a Pkd1 haploinsufficient state on AVP signaling and water handling in the absence of intercurrent mechanisms.

Several lines of evidence show that Pkd1^+/– mice have a SIAD. At baseline, the Pkd1^+/– mice have a decreased urinary output with higher urine sodium/osmolality and lower plasma sodium/osmolality. After water deprivation, the Pkd1^+/– mice are able to concentrate urine to a greater extent than WT littermates. Conversely, they have an impaired ability to excrete a water load. All of these elements indicate that haploinsufficiency in Pkd1 is associated with abnormal osmoregulation and a positive water balance. That both the water intake and the expression of AVP in the brain are unchanged, irrespective of the chronic hypo-osmolality, does suggest a central defect in Pkd1^+/– mice. Such a central defect could reflect high expression levels of polycystins in the brain (20) and a potential role in the pathways that regulate AVP secretion. Of note, two potential mechanisms may explain the significantly lower values of urine AVP excretion in the Pkd1^+/– mice: (1) The water retention, causing a decrease in the urinary output, and (2) that urinary AVP excretion is influenced by the osmolar clearance (Cosm), as a result of interference with the reabsorption/degradation of filtered AVP in the proximal tubule (31). Accordingly, the lower urinary AVP excretion could reflect the significant decrease in Cosm in Pkd1^+/– versus WT mice (3.9 ± 0.2 versus 6.2 ± 0.3 µl/min; n = 15 pairs; P < 0.0001), leading to accelerated tubular degradation of AVP. Recent studies have shown an increased reactivity of the aortic and renal vasculature in Pkd1^+/– mice (32), and a reduced renal blood flow could explain such a reduced Cosm. At any rate, the positive water balance of noncystic Pkd1^+/– mice contrasts with the mild concentrating defect reported in patients with ADPKD (14). The existence of nephrogenic diabetes insipidus in conditions that are associated with structural changes in the medulla (16) and the correlation between the number of renal cysts and the extent of the concentrating defect (33) suggest that the latter primarily reflects cystic changes in the medulla of patients with ADPKD (11).

Kidney-specific mechanisms are also involved in the inappropriate antidiuresis phenotype of the Pkd1^+/– mice (Figure 8). In normal conditions, the binding of AVP to V2R at the basolateral pole of CD principal cells triggers a heterotrimeric G-protein–coupled cascade, activating AC6 and increasing cAMP levels, which leads to the phosphorylation of AQP2 at Ser256 by protein kinase A (PKA), followed by the trafficking of p-AQP2 to the apical plasma membrane and the increase in the osmotic water permeability of the cells. The cAMP-induced translocation of AQP2 is facilitated by PKA-mediated phosphorylation (Ser188) of the small GTP-binding protein RhoA, causing Rho inactivation and depolymerization of F-actin (28). In some conditions, the V2R-mediated antidiuretic actions of AVP may be balanced by the apical V1aR and CaR in the principal cells (31,34,35). Binding of luminal AVP to V1aR stimulates phospholipase C (PLC), leading to inositol trisphosphate receptor (IP3R)-mediated release of Ca²⁺ from the endoplasmic reticulum. In turn, increased [Ca²⁺]_i activates phosphodiesterase-1 (PDE1) and inhibits AC6, leading to decreased cAMP. However, activation of CaR by high luminal calcium concentrations leads to (1) increased [Ca²⁺]_i via PLC and IP3R and (2) activation of protein kinase C (PKC) and phosphorylation of ERK1/2, followed by activation of phospholipase A2 (PLA₂), release of PGE₂, and prostaglandin EP₃ receptor (EP₃R)-mediated activation of RhoA, resulting in F-actin formation and reduced insertion of AQP2 into the apical plasma membrane (28,36,37). The stimulation of CaR may also activate PKC isoforms that mediate AQP2 endocytosis (34,38). Several abnormalities in these signaling pathways could potentially lead to the inappropriate water retention in the Pkd1^+/– mice, as discussed next.

View larger version (32K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 8. Model for the effects of PKD1 haploinsufficiency on AVP signaling and AQP2 trafficking in the PC of CD. Representation of a typical PC of the CD showing the influence of [Ca2+]i levels and various signaling pathways on the trafficking of AQP2 and the modifications in Pkd1+/– mice. In the normal state, the binding of AVP to the basolateral V2R activates adenylyl cyclase 6 (AC6), resulting in cAMP-dependent activation of protein kinase A (PKA) and phosphorylation of AQP2 (Ser256) and its insertion into the apical membrane. The process is facilitated by PKA-mediated phosphorylation of RhoA (Ser188), which inactivates RhoA and causes the depolymerization of F-actin. The V2R-mediated effects of AVP could be balanced by the apical vasopressin-1a (V1aR) and calcium-sensing (CaR) receptors. Luminal AVP activates V1aR, which increases [Ca2+]i via phospholipase C (PLC) and inositol trisphosphate receptor (IP3R). In turn, the increased Ca2+ activates phosphodiesterase-1 (PDE1) and inhibits AC6, leading to decreased cAMP. High extracellular, urinary Ca2+ levels can activate CaR, leading to (1) increased [Ca2+]i via PLC and IP3R and (2) activation of PKC and dual phosphorylation of ERK1/2 via PLC and diacylglycerol (DAG), leading to activation of phospholipase A2 (PLA2) and release of PGE2, activation of prostaglandin EP3 receptor, and downstream activation of RhoA with subsequent F-actin formation, which reduces the insertion of AQP2 into the apical plasma membrane. The polycystin-1 and -2 interact in the primary cilium located in the apical plasma membrane of CD cells. In response to luminal flow, the polycystin-1/2 complex regulates [Ca2+]i by mediating a Ca2+ entry into the cell, which releases Ca2+ stores via the ryanodine receptors (RyR) on the endoplasmic reticulum (ER). The haploinsufficient Pkd1 state is characterized by the increased recruitment of AQP2 into the apical plasma membrane, reflecting decreased [Ca2+]i, decreased ERK-PLA2 activity, increased PKA-mediated phosphorylation of AQP2 and RhoA, and decreased activity of RhoA. The decreased [Ca2+]i may also result in increased efficiency of the cAMP-mediated signaling in microdomains of the cell. These events increase the efficiency of V2R-mediated signaling, leading to the recruitment of AQP2 in the apical plasma membrane and inappropriate reabsorption of water by the PC. The large arrows indicate the changes that were documented in Pkd1+/– mice. The increased urinary concentrations of calcium and AVP are between parentheses because there is no evidence of stimulated apical receptors. Adapted from references (31,34–36).

Second, we documented an increased p-AQP2 and recruitment of AQP2 in the apical plasma membrane of CD cells and increased p-RhoA coupled to decreased activity of RhoA in the Pkd1^+/– kidneys. These data suggest that inactivation of RhoA, causing the depolymerization of F-actin, facilitates the AVP-elicited insertion of AQP2 into the apical plasma membrane of the Pkd1^+/– mice. Several factors could contribute to these modifications, including the decreased [Ca²⁺]_i; the increased PKA-mediated phosphorylation of AQP2 and RhoA (28); and the less active ERK-PLA2 pathway, as indicated by the lower p-ERK1/2 over total ERK1/2 ratio and a trend for lower urinary PGE₂ excretion (28,36). These modifications of the ERK pathway are different from observations in cultured renal cells. Yamaguchi et al. (44) showed that the cAMP-dependent proliferation of cultured human ADPKD cyst-lining cells is mediated through phosphorylation/activation of ERK. By contrast, cAMP inhibits ERK activity and slows proliferation in normal epithelial cells from human kidney cortex. However, when immortalized mouse M1 cortical CD cells were treated with calcium channel blockers or EGTA to lower [Ca²⁺]_i, the cells converted to a cystic-like phenotype, with cAMP-dependent activation of ERK and proliferation. Of note, lowering [Ca²⁺]_i alone (a situation that is similar to that observed in Pkd1^+/– kidneys) was not sufficient to activate ERK and proliferation in these cells (45). Many elements contribute to the differences between the native, noncystic Pkd1^+/– kidneys and cultured cells, including time course and magnitude of [Ca²⁺]_i modifications, residual levels of polycystin-1, and adaptation mechanisms, yet the lower [Ca²⁺]_i levels that were observed in the Pkd1^+/– CD cells may represent an intermediate state, in which a further loss of polycystin-1 or changes in cAMP levels may lead to a proliferative or cystic phenotype. Conversely, a low [Ca²⁺]_i could decrease the activity of Ca²⁺-dependent protein phosphatases and calcineurin-A and , resulting in higher levels of phosphorylated AQP2 and a reduced recycling from the apical plasma membrane (46).

Third, the apical V1aR and CaR pathways seem to be inactive in Pkd1^+/– mice, as evidenced by lower [Ca²⁺]_i in isolated CD, decreased activity of the ERK-PLA2 pathway, and decreased RhoA activity. As discussed, the apical CaR may sense urinary calcium levels and influence AQP2 targeting to adjust water homeostasis. The experiments of Sands et al. (38), performed on isolated rat IMCD, showed that increasing luminal calcium from 1 to 5 mM causes a 30% decrease in the AVP-elicited osmotic water permeability but no change in the basal permeability. Similar calcium concentrations were used to show the link between CaR activation and AQP2 trafficking in cultured cells (34). These calcium concentrations are 100-fold higher than those observed in Pkd1^+/– mice (50 µM, only 1.5-fold higher than in WT mice). A significant polyuria has also been observed in hypercalciuric mouse models in vivo. For instance, the diuresis is increased two-fold in the Trpv5-null mice, characterized by very high calciuresis (averaging 250 µmol/24 h, six-fold higher than in WT littermates) (47). Again, these conditions are very distinct from the Pkd1^+/– mice, which have no real hypercalciuria (average 20 µmol/24 h). Regarding the luminal V1a receptors, their antagonistic action has been demonstrated only in rabbit CD, with no evidence in rat or mouse kidney (31). Therefore, the increased luminal concentrations of calcium and AVP in Pkd1^+/– mice versus WT probably have limited biologic relevance.

Fourth, the unchanged cAMP levels in kidney and urine and the stable expression of AQP2 and V2R mRNA expression in the kidney of Pkd1^+/– mice argue against a direct role of chronically increased cAMP levels in this model. Furthermore, the mRNA level of targets of the V2R signaling pathway (including urea transporter 1 and the subunit of epithelial sodium channel) were similar in Pkd1^+/+ and Pkd1^+/– mice. These data contrast with the elevation of cAMP, paralleled by the upregulation of AQP2 and V2R mRNA that was observed in other PKD mouse models (8,9). However, all of these models show renal cysts, with a positive correlation between cAMP level and the magnitude of cystic changes, suggesting a causal link between the two (10). Although we did not detect an increase in cAMP in whole tissue, we cannot exclude that a lower [Ca²⁺]_i could favor a local increase in cAMP by a dual effect on AC6 and phosphodiesterase E1 in the principal cells of the Pkd1^+/– mice. Indeed, recent studies suggested that compartmentalization of cAMP signaling in microdomains may participate in the regulation of AQP2 trafficking (48).

Fifth, in addition to the aforementioned mechanisms, the trafficking of AQP2 in CD cells can be stimulated by cGMP via activation of NOS (49). The Pkd1^+/– mice showed no difference in the renal expression of eNOS and nNOS isoforms but a significant decrease in the urinary excretion of NO metabolites. These data, which confirm previous reports of impaired NO synthesis in this mouse model (17) and patients with ADPKD (50,51), suggest that the cGMP-mediated cascade is probably not involved in the recruitment of AQP2. Furthermore, the decreased [Ca²⁺]_i in Pkd1^+/– CD could participate in the decreased renal NOS activity, because eNOS and nNOS both are Ca²⁺ dependent. Therefore, a reduced generation of NO could decrease the medullary blood flow, sensitizing the Pkd1^+/– mice to the sympathetic tone and contributing to the antidiuresis phenotype by a positive effect on medullary hypertonicity. Another factor is medullary ET1, which antagonizes the AVP-induced cAMP accumulation in CD cells and increases medullary blood flow in vivo (52). The ET1 action is mediated by the ETB receptor, and it involves PLC, PLA2, and PKC, as well as [Ca²⁺]_i, NO, and PGE2. Mice that lack ET1 in the CD show no abnormalities at baseline but a reduced ability to excrete urine during acute water loading (53). Together with the reduced [Ca²⁺]_i and decreased NO production, the mild but significant decrease in the mRNA expression of ET1 evidenced in Pkd1^+/– kidneys could thus contribute both to increased AVP signaling and to decreased medullary blood flow, leading to water retention.

The Pkd1^+/– mice show a significant resistance to V2R antagonism, despite similar distribution and affinity of the V2R and unchanged mRNA expression of AVP. This observation may be relevant for the use of V2R antagonists in patients with ADPKD, with the aim to interfere with the cystogenic effect of cAMP (11). In the sole orthologous model of ADPKD that has been investigated thus far (Pkd2^–/tm1Som mouse), treatment with the V2R antagonist OPC31260 has not been associated with significant changes in urine output and osmolality, and a comparison of its efficacy in WT and Pkd2 mice has not been reported (9). Of interest, it has been shown recently that a partial loss of Pkd2 attenuated the polyuria and increased the urine osmolality in a mutant V2R mouse model of nephrogenic diabetes insipidus (54), suggesting that an alteration of the polycystin pathway may indeed cause water retention by CD principal cells. Another interesting observation is that hyponatremia is frequently observed in neonates with autosomal recessive polycystic kidney disease (55).

Finally, it should be pointed out that the Pkd1^+/– mice that were studied here represent a potentially interesting model to investigate water handling in the CD. A new disease entity, the nephrogenic SIAD, has recently been attributed to activating (gain-of-function) missense mutations in AVPR2, the gene that encodes V2R in humans (56). To the best of our knowledge, there are no genetically modified mice with such an antidiuresis phenotype at baseline. An impaired ability to lower urine osmolality and increase urinary water excretion was recently reported in mice that lack the taurine transporter gene Taut (57), which could play a role in primary cilia (58). Thus, in addition to the interactions between polycystins and calcium signaling pathways, the Pkd1^+/– mice could give insights into the mechanisms that govern osmoregulation, AVP signaling, and trafficking of AQP2 in the CD.

	Disclosures

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None.

	Acknowledgments

 
These studies were supported in part by the Fonds National de la Recherche Scientifique, the Fonds de la Recherche Scientifique Médicale, an Action de Recherche Concertée (ARC 05/10-328), an IAP VI, and the EuReGene integrated project (FP6).

Some of these data were presented during the 39th annual meeting of the American Society of Nephrology; November 16 through 19, 2006; San Diego, CA; and published in abstract form (J Am Soc Nephrol 17: 513A, 2006).

We are grateful to V. Beaujean, Y. Cnops, H. Debaix, F. Jouret, K. Parreira, and L. Wenderickx for excellent assistance and Profs. L. Bankir, D. Bichet, L. Guay-Woodford, J.-C. Henquin, N. Morel, Y. Pirson, A. Robert, R. Sandford, P. Steels, V. Torres, E. Van Kerkhove, and Dr. I. Smets for helpful discussions. We thank Sanofi-Aventis for providing SR121463B.

	Footnotes

 
Published online ahead of print. Publication date available at www.jasn.org.

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