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PROTEIN AND VESICLE TRAFFICKING, CYTOSKELETON

Microtubules are needed for the perinuclear positioning of aquaporin-2 after its endocytic retrieval in renal principal cells

¹Leibniz-Institut für Molekulare Pharmakologie (FMP), Campus Berlin-Buch, Berlin, Germany; and ²Institut für Pharmakologie, Charité-Universitätsmedizin Berlin, Campus Benjamin Franklin, Freie Universität Berlin, Berlin, Germany

Submitted 21 December 2006 ; accepted in final form 6 July 2007

	ABSTRACT

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Water reabsorption in the renal collecting duct is regulated by arginine vasopressin (AVP). AVP induces the insertion of the water channel aquaporin-2 (AQP2) into the plasma membrane of principal cells, thereby increasing the osmotic water permeability. The redistribution of AQP2 to the plasma membrane is a cAMP-dependent process and thus a paradigm for cAMP-controlled exocytic processes. Using primary cultured rat inner medullary collecting duct cells, we show that the redistribution of AQP2 to the plasma membrane is accompanied by the reorganization of microtubules and the redistribution of the small GTPase Rab11. In resting cells, AQP2 is colocalized with Rab11 perinuclearly. AVP induced the redistribution of AQP2 to the plasma membrane and of Rab11 to the cell periphery. The redistribution of both proteins was increased when microtubules were depolymerized by nocodazole. In addition, the depolymerization of microtubules prevented the perinuclear positioning of AQP2 and Rab11 in resting cells, which was restored if nocodazole was washed out and microtubules repolymerized. After internalization of AQP2, induced by removal of AVP, forskolin triggered the AQP2 redistribution to the plasma membrane even if microtubules were depolymerized and without the previous positioning of AQP2 in the perinuclear recycling compartment. Collectively, the data indicate that microtubule-dependent transport of AQP2 is predominantly responsible for trafficking and localization of AQP2 inside the cell after its internalization but not for the exocytic transport of the water channel. We also demonstrate that cAMP-signaling regulates the localization of Rab11-positive recycling endosomes in renal principal cells.

dynein; Rab11

The cytoskeleton plays a central role in vesicular transport (50, 37). Our group (19, 42, 43) has previously shown that depolymerization of F-actin by cytochalasin D or by inhibition of the small GTPase RhoA resulted in the redistribution of AQP2 to the plasma membrane of primary cultured renal inner medulla collecting duct (IMCD) cells in the absence of cAMP elevation. Early work suggested that microtubules also play an important role in the AVP-regulated water reabsorption (32, 33). However, Tajika et al. (41) recently showed that in polarized Madin-Darby canine kidney (MDCK) cells stably expressing human AQP2 the forskolin-induced AQP2 redistribution to the apical plasma membrane was not prevented by the disruption of microtubules.

Microtubules regulate the localization of the perinuclear Rab11-positive recycling compartment (9, 47). The small GTPase Rab11 is involved in the recycling of different membrane proteins, including the transferrin receptor (31), the thromboxane A₂ receptor (44), and the glucose transporter GLUT4 (46). Rab11 was also detected on AQP2-bearing vesicles isolated from rat kidney or primary cultured IMCD cells (1, 27). Here, it apparently serves as the vesicular receptor for the motor protein myosin Vb, which transports AQP2-bearing vesicles along the F-actin cytoskeleton to the plasma membrane through a Rab11-positive compartment (27). Constitutive recycling of AQP2 has been demonstrated by Brown and coworkers (17).

The present study focuses on the role of microtubules in AQP2 trafficking in renal principal cells. Using IMCD cells, we show that the AQP2 redistribution from intracellular vesicles to the plasma membrane is accompanied by the reorganization of microtubules and the redistribution of Rab11 to the cell periphery. Our data indicate that microtubules are mainly necessary for the positioning of AQP2 and Rab11 in the perinuclear region after endocytic retrieval of AQP2. The exocytic transport of AQP2 to the plasma membrane is apparently largely independent of microtubules.

	MATERIALS AND METHODS

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Reagents. AVP was synthesized by M. Beyermann (FMP Berlin), dibutyryl-cAMP (dbcAMP) was purchased from Biolog (Bremen, Germany), and erythro-9-[3-(2-hydroxynonyl)]adenine (EHNA) was from Calbiochem (Darmstadt, Germany). All other chemicals were obtained from Sigma (Deisenhofen, Germany).

Antibodies. AQP2 was detected with an affinity-purified rabbit antibody (H27) and used as previously described (19). Microtubule staining was achieved with a mouse MAb directed against -tubulin (1:200; Calbiochem). Rab11 staining was performed with a mouse MAb (1:100; BD Bioscience, Heidelberg, Germany). For decorating the Golgi apparatus, we used mouse MAb 58K Golgi marker (1:100; Abcam, Cambridge, UK). Clathrin was detected with a specific mouse MAb (1:100; BD Biosciences). Mouse IgG₁ and IgG_2a isotype control antibodies and rabbit serum (all purchased from Sigma) were used as negative controls for immunostaining (data not shown). F-actin was stained with Alexa fluor 647-conjugated phalloidin (Invitrogen, Karsruhe, Germany). Nuclei were counterstained with 4',6-diamidino-2-phenylindole. Secondary antibodies were Cy3 or Cy5 coupled (1:600; Dianova, Hamburg, Germany).

Cell culture. Primary cultures of renal IMCD cells were obtained from kidneys of at least 6-wk-old Wistar rats as described previously (23, 39). The cells were maintained in DMEM adjusted to 600 mosM and supplemented with 1% Ultroser, 0.45% glucose, 1% nonessential amino acids, 2 mM L-glutamine, 50 µg/ml gentamicin, 20 U/ml nystatin, and 500 µM dbcAMP. Nystatin and dbcAMP were removed 16 h before the experiment.

Immunocytochemistry and quantification of immunofluorescence intensities. IMCD cells were grown on coverslips. The cells were fixed for 15 min in buffer containing 2.5% paraformaldehyde, 100 mM cacodylate, and 100 mM sucrose and permeabilized by incubation in 0.1% Triton X-100 for 5 min. After samples were washed in PBS, unspecific binding sites were blocked with 0.014% fish skin gelatin (Sigma). Primary and secondary antibodies were applied in a humidifying chamber for 45 min each at 37°C. The cells were washed with PBS, and coverslips were mounted with Immu-Mount (Thermo Shandon, Frankfurt, Germany). In all cases, we controlled specificity of staining by using isotype control immunoglobulins. Specificity of secondary antibodies was confirmed by omitting primary antibodies.

Fluorescence signals were detected by confocal laser scanning microscopy (LSM 510 META-UV; Carl-Zeiss, Jena, Germany) using a Plan Neofluor x100/1.3 oil objective. To quantitatively evaluate the cellular distributions of AQP2, Rab11, and microtubules, fluorescence signal intensities were measured in defined regions of interest (ROIs) in close proximity around the nucleus (2 µm around the nucleus) and at the plasma membrane (0.5 µm apart from the plasma membrane; for details see supplementary Fig. S1). (The online version of this article contains supplemental data.) The ratios of perinuclear to plasma membrane fluorescence signal intensities were calculated from these values. The data are shown as means ± SE.

Determination of polymerized tubulin. To estimate the effects of the microtubule-disrupting agent nocodazole and microtubule-stabilizing taxol in IMCD cells, the cells were treated with these compounds and lysed in tubulin extraction buffer [in mM: 20 PIPES (pH 6.8), 140 NaCl, 1 Mg₂Cl, 1 EGTA, 0.5% Nonidet P-40 (NP-40), 0.5 PMSF, 4 taxol] at room temperature. Lysates were centrifuged at 13,000 g for 10 min at room temperature. The pellets were considered NP-40-insoluble fractions containing polymerized tubulin (30). Each fraction was mixed with SDS-PAGE sample buffer and boiled for 10 min. Proteins were separated by SDS-PAGE, and tubulin was detected by Western blotting.

Measurements of water permeability and cell volume. Laser scanning reflection microscopy was applied to determine the changes in water permeability and corresponding cell volume of IMCD cells as previously described (24). As a modification of the original method, we did not trypsinize the cells before the measurement to avoid possible effects of trypsinization on the cytoskeleton. In brief, IMCD cells were grown on coverslips, and unstained individualized cells that resembled principal cells were selected by the transmission mode of the Zeiss LSM 510 Meta UV microscope. The optical settings were changed to the reflection mode. The cells were initially bathed in 600 mosM culture medium, which was changed to 300 mosM medium at a defined time point during the measurement. This change in osmolarity led to an osmotic water influx into the cells and an increase of cell volume. Cells were scanned using the reflection x-z-scan mode. The increase of vertical x-z-scan section areas from single cells was determined. This increase reflects the increase of the cell volume due to osmotic water influx. In the experiments, cells were either left untreated or were pretreated with nocodazole. Redistribution of AQP2 to the plasma membrane was induced by forskolin.

Statistical analysis. Data were analyzed with GraphPad Instat software using the unpaired t-test with Welch correction. P values <0.05 were considered significant.

	RESULTS

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Elevation of cAMP leads to reorganization of microtubules and redistribution of Rab11 in IMCD cells. An increase in PKA activity induces the reorganization of microtubules in K562 leukemia cells (16). Therefore, we tested by immunofluorescence microscopy whether treatment of IMCD cells with cAMP-elevating stimuli alters the microtubule architecture. Under basal conditions, most cells showed a prominent dense network of microtubules around the nucleus with considerably less microtubule content in the cell periphery. Treatment with AVP resulted in an increased formation of microtubules in the cell periphery (Fig. 1A). To quantify the effect of AVP on the microtubule organization, ROIs around the nucleus and at the plasma membrane were defined in which fluorescence intensities were determined (for details to ROI definition, see Fig. S1 in supplemental materials). This analysis revealed that AVP decreased the ratio of perinuclear to the plasma membrane fluorescence signal intensity and thus confirmed the redistribution of microtubules (Fig. 1B). The reorganization of microtubules apparently depends on an elevated cAMP level because treatment of IMCD cells with forskolin, an activator of adenylate cyclase, caused a reorganization of microtubules similar to that induced by AVP (data not shown).

View larger version (50K): [in this window] [in a new window]   Fig. 1. Arginine vasopressin (AVP) increases reorganization of microtubules in inner medullary collecting duct (IMCD) cells. A: IMCD cells were left untreated, treated with AVP (100 nM, 30 min), or treated with AVP that was removed after 30 min by washout followed by incubation in cell culture medium for 1 h. Tubulin was detected by immunostaining. Scale bar = 20 µm. B: quantification of the cellular distribution of microtubules after AVP treatment and AVP washout. The distribution was quantified by calculating the ratio between fluorescence signal intensities in the defined regions of interest around the nucleus and underneath the plasma membrane (for details of the quantification, see legend to supplemental Fig. S1). Values are means ± SE (n > 31 cells for each condition; 3 independent experiments). *Significantly different from controls (P < 0.05).

Microtubules localize the Rab11-positive recycling compartment in cells (47, 9). Disruption of microtubules by nocodazole results in the redistribution of Rab11 to the cell periphery of MDCK cells (21). We investigated whether the reorganization of microtubules under AVP treatment is accompanied by the redistribution of Rab11 in IMCD cells. Although under basal conditions Rab11 is mainly localized perinuclearly (Fig. 2; see also Ref. 27), AVP induced the redistribution of Rab11 from the perinuclear localization to a more diffuse distribution throughout the cell (Fig. 2). As previously shown, treatment of IMCD cells with AVP caused the redistribution of AQP2 to the plasma membrane (Fig. 2) (23). Quantitative analyses carried out as described in Fig. S1 in the supplemental materials confirmed these observations (Fig. 2). To confirm that the secondary anti-rabbit antibody does not cross-react with primary mouse anti-Rab11, we stained the cells with mouse anti-Rab11 and anti-rabbit-Cy3. To rule out cross-reactivity of the secondary anti-mouse antibody with the primary rabbit anti-AQP2 antibody, cells were stained with rabbit anti-AQP2 and anti-mouse-Cy5 antibodies. No signals were observed in either case (data not shown), demonstrating no cross-reactivity of secondary antibodies.

View larger version (39K): [in this window] [in a new window]   Fig. 2. Effects of AVP and nocodazole on the distribution of aquaporin-2 (AQP2) and Rab11 in IMCD cells. A: detection of -tubulin in Nonidet P-40 (NP-40)-insoluble fractions of IMCD cells. IMCD cells were either left untreated (basal) or treated with nocodazole (30 µM) or taxol (10 µM) for the indicated times. The cells were lysed, and NP-40-insoluble, polymerized tubulin-containing fractions were prepared and analyzed by Western blotting for tubulin content. Shown is the result of 1 of 2 independent experiments that yielded similar results. The separation by SDS-PAGE causes polymerized tubulin to migrate as the monomeric form in the gel. B: IMCD cells were left untreated or incubated with AVP (100 nM, 30 min) or nocodazole (30 µM, 2 h). Where indicated, nocodazole was removed after 2 h by washout, and cells were further incubated in cell culture medium for 1 h. AQP2, Rab11, and clathrin were detected by immunostaining. Overlays of AQP2 and Rab11 or AQP2 and clathrin images are also shown (bottom, merge). Scale bar = 20 µm. C: quantification of the intracellular distribution of AQP2 and Rab11 following the treatments indicated in B. The distribution was quantified by calculating the ratios between fluorescence signal intensities in the perinuclear region and at the plasma membrane as indicated in the legend to supplemental Fig. S1 (n > 32 cells for each condition; means ± SE; 3 independent experiments). *Significantly different from controls (P < 0.05).

When IMCD cells were treated with nocodazole for 2 h, a loss of the pronounced localization of AQP2 and Rab11 around the nucleus was evident. Instead, both proteins were found throughout the cell, including the plasma membrane (Fig. 2, B and C). A washout of nocodazole led to repolymerization of microtubules (not shown; see also Fig. 4A) and rescued the perinuclear localization of AQP2 and Rab11 (Fig. 2, B and C), indicating that this process of positioning is dependent on microtubules. Thus, under basal conditions, a major part of AQP2 is kept in a Rab11-positive compartment, which is held in a perinuclear position by microtubules. Interestingly, after washout of nocodazole, AQP2- and Rab11-positive compartments appear more compact than shown in untreated cells (Fig. 2B). Treatment of renal epithelial LLC-PK₁ cells stably expressing rat AQP2 with an inhibitor of V-ATPase [bafilomycin A₁ (125 nM; 2 h)] or exposure to 20°C for 2 h caused a similar clustering of AQP2 and of clathrin in a perinuclear region, which represents the trans-Golgi network (17). In contrast, no clustering of clathrin was observed in IMCD cells when they were stained for clathrin after nocodazole washout (Fig. 2B), indicating that the AQP2-containing compartments affected by bafilomycin A₁ in LLC-PK₁ cells and by nocodazole in IMCD cells are not the same.

View larger version (42K): [in this window] [in a new window]   Fig. 3. AVP induces the redistribution of AQP2 from intracellular compartment to the plasma membrane in the absence of microtubules. A: IMCD cells were treated with AVP (100 nM, 30 min), AVP was washed out, and cells were further incubated in cell culture medium for 2 h (washout) without or with nocodazole (30 µM). In addition, cells were treated with forskolin (50 µM) during the last 15 min of washout. The cells were fixed, and AQP2 and Rab11 were detected by immunostaining. Overlays of AQP2 and Rab11 are also shown (bottom row, merge). Scale bar = 20 µm. B: quantification of the intracellular distribution of AQP2 and Rab11 in IMCD cells treated as indicated in A. The distribution was quantified by calculating the ratio between fluorescence signal intensities in the perinuclear region and at the plasma membrane as indicated in supplementary Fig. S1 (n > 22 cells for each condition; means ± SE; 3 independent experiments). *Significantly different from controls (P < 0.05). C: IMCD cells were challenged with a change from 600 to 300 mosM medium. The increase of cell volume due to an increase in osmotic water permeability was determined by laser scanning reflection microscopy. Cells were left untreated or pretreated with nocodazole (30 µM, 2 h). Where indicated, the redistribution of AQP2 to the plasma membrane was induced by forskolin (50 µM, 15 min). After this, cells were transferred into hypotonic medium and swelling was measured. The graph shows the increase of vertical x-z-scan section areas in %, which reflects increase of cell volume due to water uptake (n > 20 cells per experimental condition; means ± SE of 3 independent experiments). *Significantly different from the untreated cells (P < 0.0001); significantly different from cells that were pretreated with forskolin (FSK) alone (P < 0.05).

View larger version (62K): [in this window] [in a new window]   Fig. 4. Disruption of the Golgi apparatus does not affect the perinuclear positioning of AQP2. A: IMCD cells were left untreated or treated with nocodazole (30 µM, 2 h), 5 µg/ml brefeldin A (BFA) for 2 h, 5 µg/ml BFA for 2 h with addition of AVP (100 nM) for the last 30 min, or nocodazole (30 µM, 2 h) removed by washout with cells further incubated for 2 h in cell culture medium containing 5 µg/ml BFA. The cells were fixed, and AQP2, Golgi apparatus marker 58K, and tubulin were detected by immunostaining. Because both anti--tubulin and anti-58K antibodies were raised in mouse, cells were either stained for AQP2 and 58K (top 3 rows), or cells from the same experiments were stained for AQP2 (not shown) and -tubulin (bottom row). Shown are representative images. Scale bar = 20 µm. B: quantification of the intracellular distribution of AQP2 under the conditions indicated in A. The distribution was quantified by calculating the ratio between fluorescence signal intensities in the perinuclear region and in the plasma membrane as indicated in supplementary Fig. S1 (n > 33 cells for each condition; means ± SE; 3 independent experiments). *Significantly different from controls (P < 0.05).

AVP treatment after incubation with nocodazole resulted in an enhanced redistribution of Rab11 to the cell periphery compared with treatments with AVP or nocodazole alone (Fig. 3, A and B; compare with Fig. 2, B and C).

Removal of AVP causes a rapid decrease of water permeability of collecting ducts (40) and the internalization of AQP2 into principal cells (Fig. 3A). After washout of AVP, AQP2 and Rab11 were again mainly localized perinuclearly (Fig. 3A). However, when cells were treated with nocodazole during washout of AVP, a diffuse intracellular distribution of both AQP2 and Rab11 was observed (Fig. 3A). We addressed the question of whether perinuclear positioning of AQP2 and Rab11 after an initial AVP-induced redistribution to the plasma membrane and subsequent internalization on removal of AVP is required for redistribution of AQP2 to the plasma membrane in response to a second stimulation. For this purpose, IMCD cells were treated with AVP for 30 min followed by a washout and further incubation with or without nocodazole. During the last 15 min of the incubation, forskolin was added. As shown in Fig. 3A, forskolin induced the redistribution of AQP2 to the plasma membrane. This was not prevented by depolymerization of microtubules by nocodazole (Fig. 3, A and B). Apparently, transport of AQP2 to the perinuclear region is not a prerequisite for the redistribution of AQP2 to the plasma membrane although this seems to be the "normal route" (27). The second redistribution of AQP2 to the plasma membrane was achieved by a second treatment with forskolin. In contrast, a second treatment with AVP induced the AQP2 redistribution to the plasma membrane only in the minority of cells. This is most likely due to an increased internalization and degradation of the V2R after the first AVP treatment (7) and is consistent with a previous study (18). The explanation is supported by the observation that AVP induced the redistribution of AQP2 to the plasma membrane after a previous treatment with forskolin followed by 2 h washout, conditions where no downregulation of the V2R occurs (data not shown).

Disintegration of the Golgi apparatus does not induce the redistribution of AQP2 from the perinuclear region toward the cell periphery. The Golgi apparatus is maintained in its structure and in adjacent position to the nucleus by microtubules and microtubule-associated proteins (2). When IMCD cells were treated with nocodazole, the Golgi apparatus lost its tubular structure and dispersed into smaller vesicle-like structures that were dispersed throughout the cytoplasm (Fig. 4A). To evaluate whether the loss of the perinuclear position of AQP2 after nocodazole treatment may be due to disintegration of the Golgi apparatus, brefeldin A (BFA), a fungal metabolite that causes reversible dispersion of the Golgi, was used. BFA had no influence on the perinuclear position of AQP2 while the Golgi apparatus was completely dispersed (Fig. 4A). IMCD cells were next treated with nocodazole for 2 h, nocodazole was washed out, and cells were further incubated with BFA. Under these conditions, cells showed repolymerized microtubules and a dispersed Golgi at the same time. Importantly, a perinuclear position of AQP2 was restored even without reorganization of the Golgi apparatus (Fig. 4). Interestingly, in IMCD cells treated with BFA, no AVP-induced redistribution of AQP2 to the plasma membrane was observed (Fig. 4).

Inhibition of the microtubule-associated motor protein dynein alters the perinuclear positioning of AQP2 and Rab11. We applied the dynein inhibitor EHNA to investigate whether the perinuclear positioning of AQP2 and Rab11 involves this microtubule-associated minus-end-directed motor protein. EHNA treatment resulted in an impaired transport of both proteins to the perinuclear space under basal conditions and on internalization of AQP2 after removal of AVP (Fig. 5). These observations point to a role of dynein in this step of transport. Several authors showed a role of dynein in the stabilization of microtubules (20, 28). In IMCD cells, however, microtubules appeared unaffected by EHNA when microscopically inspected (data not shown).

View larger version (60K): [in this window] [in a new window]   Fig. 5. Erythro-9-[3-(2-hydroxynonyl)]adenine (EHNA), an inhibitor of the microtubule-dependent motor protein dynein, prevents the transport of AQP2 and Rab11 to the perinuclear region. A: IMCD cells were left untreated or treated with AVP (100 nM, 30 min) followed by a washout and further incubation in cell culture medium for 2 h, with EHNA (100 µM, 2 h), or with AVP (100 nM, 30 min) followed by a washout and subsequent incubation in cell culture medium containing EHNA (100 µM) for 2 h. Scale bar = 20 µm. B: quantification of the intracellular distribution of AQP2 and Rab11 under the conditions indicated in A. The distribution was quantified by calculating the ratio between fluorescence signal intensities in the perinuclear region and at the plasma membrane (see supplementary Fig. S1 for details; n > 17 cells for each condition; means ± SE; 3 independent experiments). *Significantly different from controls (P < 0.05).

	DISCUSSION

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The clustering of AQP2 and Rab11 in IMCD cells after washout of nocodazole showed similarity with the bafilomycin A-induced clustering of AQP2 and clathrin in AQP2-expressing LLC-PK₁ cells (17). However, we did not find clathrin clustering after washout of nocodazole in IMCD cells [note the limited colocalization of AQP2 and clathrin in Fig. 2B and the large extend of overlap of the two proteins in LLC-PK₁ cells in the work of Gustafson et al. (17)]. One possible explanation is that AQP2 (at least partially) localizes in different compartments in IMCD and LLC-PK₁ cells. This may be due to the different origins of the two cell types. Whereas LLC-PK₁ cells are from swine cortical collecting ducts and stably overexpress AQP2 at a constitutively high level under the control of the cytomegalovirus promoter, IMCD cells are derived from rat renal IMCDs and express AQP2 endogenously. For the maintenance of the AQP2 expression in IMCD cells, dbcAMP is present. However, it is removed 16 h before the experiments, and the amount of AQP2 in the endoplasmic reticulum, in cis- and medial-Golgi, and in trans-Golgi in IMCD cells appears to be lower than that in LLC-PK₁ cells. The clustering of AQP2 and Rab11 most likely results from the differences in microtubule organization after washout of nocodazole (compare organization of microtubules under basal conditions and after washout of nocodazole in Fig. 4A).

Disruption of microtubules only weakly inhibited the forskolin-induced increase in osmotic water permeability in IMCD cells (Fig. 3C). Several studies indicate that the microtubule-disrupting agents colchicine and nocodazole inhibit the vasopressin-induced increase in osmotic water permeability in renal collecting ducts (12, 32). However, nocodazole did not affect the redistribution of AQP2 from intracellular vesicles to the plasma membrane in MDCK (41) or in IMCD cells (present study) when estimated by immunofluorescence microscopy. These data suggest that the AQP2 redistribution to the plasma membrane can operate when microtubules are disrupted. Alternatively, we speculate that not the inhibition of the trafficking of AQP2 per se but rather the sorting to the apical plasma membrane might be impaired under disruption of microtubules. It has been demonstrated that transport of the polymeric immunoglobulin receptor to the apical membrane was drastically affected by nocodazole, whereas only a minor effect on the transport to the basolateral membrane was observed (8). Misrouting would explain reduced vasopressin-induced water permeability observed in renal collecting ducts (12, 32). Indeed, treatment of renal collecting ducts with colchicine resulted in displacement of AQP2-bearing vesicles from the apical compartment (35). Because the amount of apically inserted AQP2 is limiting for water permeability in the collecting duct, even partial misrouting of the water channel may result in a significant impact on this parameter. Importantly, we have observed a small but significant reduction of forskolin-induced increase in water permeability of IMCD cells under treatment with nocodazole. It remains to be determined whether this effect is due to additional sorting of AQP2 to the basolateral plasma membrane to which the majority of AQP2 in IMCD cells is sorted (24, 38).

After internalization of AQP2, its redistribution to the plasma membrane after a second cAMP-elevating challenge (forskolin) was not prevented by disruption of microtubules (Fig. 3, A and B). This suggests that transport of AQP2 to the perinuclear region is not a prerequisite for the cAMP-dependent AQP2 redistribution to the plasma membrane, although this seems to be the "normal route" (27). However, the role of microtubules in the perinuclear positioning of AQP2 and the finding that treatment of IMCD cells with cAMP-elevating agents resulted in simultaneous reorganization of microtubules and redistribution of AQP2 suggest that microtubules are involved in directing the AQP2 transport, presumably to the apical plasma membrane (see above).

Our data are in line with the recent study by Tajika et al. (41), in which the role of microtubules in perinuclear positioning of AQP2 in stably transfected MDCK cells was demonstrated. Both studies show that disruption of microtubules results in a redistribution of AQP2 to the cell periphery and that the disruption does not prevent cAMP-induced redistribution of AQP2 to the plasma membrane and the internalization of AQP2. However, in the MDCK cell model system, no cAMP-induced changes in the microtubule architecture were observed and no data on water permeability were provided (41). In addition, we demonstrate that the redistribution of the Rab11-positive recycling compartment and not disruption of the Golgi apparatus is responsible for the redistribution of AQP2 when microtubules are disrupted. Moreover, our results show that the motor protein dynein is involved in centripetal transport of AQP2 after its internalization (see below).

AVP induced the redistribution of Rab11 from the perinuclear space to the cell periphery (Fig. 2B). In addition, myosin Vb redistributes to the cell periphery in response to AVP in IMCD cells (27). The data suggest an effect of cAMP on the recycling compartment in renal principal cells. Although the redistribution of the Rab11 compartment may be the result of microtubule reorganization, we provide evidence that, even under conditions when microtubules are disrupted, AVP treatment induced additional redistribution of Rab11 (Fig. 3, A and B). The redistribution of Rab11 to a position in close proximity to the plasma membrane was rarely observed after treatment with nocodazole or AVP alone (Fig. 2B) but occurred frequently when the cells were treated with AVP after depolymerization of microtubules (Fig. 3A). Thus nocodazole and AVP have additive effects, indicating that the AVP-induced redistribution of Rab11 is at least partially independent of the AVP-induced reorganization of microtubules. To our knowledge, this is the first report showing that an elevation of cAMP affects the distribution of the Rab11-positive recycling compartment. Thus Rab11-dependent recycling may be directly influenced by stimuli modulating cellular cAMP levels.

Dynein is a microtubule-associated minus-end-directed motor protein that moves vesicles and organelles toward the microtubule-organizing center (49). Dynein was suggested to be localized on AQP2-bearing vesicles (26). However, its presence on these vesicles was not confirmed by a recent study by Knepper and coworkers (1). In our experiments, the dynein inhibitor EHNA induced a redistribution of AQP2 and Rab11 similar to that induced by microtubule disruption. EHNA is also known to inhibit phosphodiesterase 2, a cGMP-stimulated phosphodiesterase that hydrolyzes cAMP and cGMP (10). The inhibitory effect on phosphodiesterase 2 is unlikely to be relevant for the observed effect because it is not expressed in rat IMCD cells (51). Moreover, radioimmunoassays revealed that EHNA did not increase levels of cAMP in IMCD cells (data not shown). EHNA induced alterations of microfilaments in African green monkey kidney and 3T3 cells (36). However, IMCD cells treated with EHNA showed no detectable changes in the appearance of F-actin when stained with Alexa fluor 647-phalloidin (data not shown). A reason for the lack of these side effects may be the much lower concentration of EHNA used in our study (100 µM) than used in several early studies (1 mM) (25, 36). Thus we believe that the effect of EHNA on the distribution of AQP2 and Rab11 is due to impairment of the dynein-mediated centripetal transport.

Among aquaporins, AQP2, AQP1, and AQP8 redistribute to the plasma membrane in response to cAMP elevation. In untreated rat hepatocytes, AQP8 is present intracellularly but redistributes to the plasma membrane after treatment with dbcAMP (15). In contrast to the redistribution of AQP2, the redistribution of AQP8 is completely blocked when microtubules were depolymerized (15). In cholangiocytes, AQP1 redistributes from intracellular vesicles to the apical membrane in a cAMP-dependent and colchicine-sensitive manner (45). The data suggest that the molecular machineries executing cAMP-dependent trafficking of aquaporins may be different for AQP1, AQP2, and AQP8.

In summary, our study indicates that microtubules play an important role in the positioning of both AQP2 and Rab11 in perinuclear compartments and suggests that the motor protein dynein is involved in their transport toward this compartment. The effects of cAMP-elevating agents on the architecture of microtubules and the distribution of Rab11 indicate that dynamic changes of the cytoskeleton and the recycling compartment may be necessary to ensure efficient and coordinated trafficking of AQP2 in renal principal cells under these conditions. The present study provides further insights into molecular mechanisms underlying the role of the cytoskeleton in the AQP2 trafficking and underlines that the redistribution of AQP2 from intracellular vesicles to the plasma membrane mainly proceeds along F-actin, whereas microtubules are predominantly involved in trafficking of AQP2 after its internalization.

Several diseases, including liver cirrhosis and syndrome of inappropriate antidiuretic hormone secretion, are associated with AVP-dependent water retention. The most prominent example, however, is chronic heart failure. Better understanding of the molecular mechanisms directing AVP-dependent water reabsorption, i.e., of the AQP2 trafficking in renal principal cells, may lead to the discovery of novel drug targets for the treatments of such diseases (22).

	GRANTS

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This work was supported by grants from the Deutsche Forschungsgemeinschaft (Kl1415/1-1 and Kl1415/3-1), the European Union (thera-cAMP, contract no. 037189), the Deutscher Akademischer Austauschdienst (Vigoni programme), and the Fonds der Chemischen Industrie.

	ACKNOWLEDGMENTS

 
We are grateful to Andrea Geelhaar, Brunhilde Oczko, and Jenny Eichhorst for excellent technical assistance.

	FOOTNOTES

 

Address for reprint requests and other correspondence: E. Klussmann, Leibniz-Institut für Molekulare Pharmakologie (FMP), Campus Berlin-Buch, Robert-Rössle-Str. 10, 13125 Berlin, Germany (e-mail: klussmann{at}fmp-berlin.de)

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	REFERENCES

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
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