Human Molecular Genetics, 2002, Vol. 11, No. 1 59-67
© 2002 Oxford University Press
Distinct subcellular expression of endogenous polycystin-2 in the plasma membrane and Golgi apparatus of MDCK cells
Department of Human and Clinical Genetics, Sylvius Laboratory, Leiden University Medical Center, 2333AL Leiden, The Netherlands and 1Department of Pathology, Leiden University Medical Center, 2333ZA Leiden, The Netherlands
Received September 7, 2001; Revised and Accepted November 5, 2001.
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ABSTRACT |
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Polycystin-2 is a predicted integral membrane protein with non-selective cation channel activity. The protein is encoded by the PKD2 gene, which is mutated in ~15% of patients with autosomal dominant polycystic kidney disease (ADPKD). Polycystin-2 can interact with the transmembrane protein polycystin-1, the product of the PKD1 gene. However, endoplasmic reticulum (ER) localization was reported for (heterologously expressed) polycystin-2 in cultured cells and baso-lateral localization has been reported in renal tissues. Using two polyclonal antisera raised against polycystin-2 we demonstrated distinct expression of the endogenous protein in the Golgi apparatus and the plasma membrane of MDCK cells. In contrast, most of the heterologously expressed polycystin-2 (PC2–EGFP) remained in the ER, substantially overlapping with the staining pattern of protein-disulfide isomerase (PDI), a marker for the ER. Only in a small subset of these cells weak plasma membrane signals were observed. Membrane staining was also suggested by immunoelectron microscopy and was confirmed by subcellular fractionation on sucrose density gradients. The plasma membrane staining disappeared following extraction with a buffer containing Triton X-100, whereas signals for polycystin-1 and E-cadherin remained visible, suggesting that polycystin-2 is neither tightly bound to the Triton X-100 insoluble cytoskeleton, nor to these proteins. We conclude that endogenous polycystin-2 is transported via the Golgi apparatus to the plasma membrane and has a broader membrane localization than polycystin-1. These data suggest that polycystin-2 can move freely in certain regions of the membrane where it probably functions as a channel, activated by, or in complex with, polycystin-1.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Autosomal dominant polycystic kidney disease (ADPKD) is one of the most commonly inherited disorders, with an incidence of 1 in 1000. The disease is characterized by the formation of a large number of fluid-filled cysts in both kidneys that together with interstitial fibrosis, cause chronic renal failure in 50% of patients by the age of 60 years (1). Frequent extra-renal manifestations are hypertension, cardio-valvular abnormalities, hepatic and pancreatic cysts, and the occurrence of cerebral aneurysms (2).
In 85% of patients the causative mutation is located in the PKD1 gene on chromosome band 16p13.3 (3,4). Of the remaining 15%, most mutations are located in the PKD2 gene on chromosome band 4q21 (5–7). The proteins encoded by these genes, polycystin-1 and polycystin-2, respectively, are members of a protein family with at least three additional homologues that are not mutated in patients with ADPKD (8–12). Computer analyses predict that these proteins are integral membrane glycoproteins. Polycystin-1 is the largest of these proteins and contains a subset of structural motifs that may be involved in protein–protein interactions (13–15). Immunostaining and precipitation experiments suggest that polycystin-1 indeed functions in cell–cell and/or cell–matrix interactions and could be a receptor, binding to a yet unknown ligand (16–21). The C-terminus of polycystin-1 has been shown to interact through an 
-helical coiled-coil domain with a region in the C-terminus of polycystin-2 (22,23). This protein shows homology to the pore forming units of cation channels (7,24). Functional biochemical studies have revealed measurable non-selective cation channel activity of both polycystin-2 alone (25,26) and co-assembled with polycystin-1 (27). Polycystin-1 affects cell proliferation and apoptosis and the C-terminus of polycystin-1 can also modify several signalling pathways such as G-protein signalling, AP-1 transcription and Wnt-signalling (28–33).
As ADPKD patients with either a PKD1 or a PKD2 mutation essentially show a similar phenotype and because polycystin-1 and polycystin-2 seem to biochemically interact (22,23), a similar subcellular localization of polycystin-2 is expected as to that observed for polycystin-1. In renal tissues, expression of both proteins is developmentally regulated, with the highest expression of polycystin-1 in fetal kidneys and of polycystin-2 in mature kidneys (34–36). Antibodies against both proteins stain the same nephronic segments, but have a reciprocal gradient of expression with the highest expression of polycystin-1 in the distal part of the nephron in fetal tissues and in collecting ducts in adult tissues. For polycystin-2 the expression levels start to increase in primitive tubules and are highest in the distal convoluted tubules and the medullary thick ascending limbs of the loop of Henle’s duct in adult tissues (34). The extra-renal polycystin-2 distribution in mice, rat and human is widespread and shows many similarities to the reported polycystin-1 expression (17,34–38). These data indicate that polycystin-1 and -2 may function both together and independently (34,36,37). At the subcellular level, polycystin-1 is predominantly localized near junction complexes in the plasma membrane (16,18,19,21,34) as well as in the cytoplasm. More conflicting data have been reported for polycystin-2 expression, varying from too low to detect in cultured cells (39) to granular cytoplasmic/endoplasmic reticulum (ER)-like signals or basolateral membrane localization in renal tissues (34).
In cultured cells, ER/Golgi localization has been reported (26,39) for heterologously expressed polycystin-2. Upon co-transfection with polycystin-1 in CHO cells (27), or after treatment with chemical chaperones in oocytes (26), the localization of polycystin-2 changed to the plasma membrane. It has been hypothesized that polycystin-2 may interact with polycystin-1 in the ER-to-Golgi transition on its way to the cell surface, or that the two proteins may interact where ER and plasma membrane are in close proximity (39,40).
Our data show that most of the heterologously expressed polycystin-2–EGFP (PC2–EGFP) fusion protein is located in the ER, while endogenous polycystin-2 is distributed in at least two subcellular pools, in the cytoplasm (Golgi apparatus) and in the plasma membrane.
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RESULTS |
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Characterization of antibodies
Polyclonal antibodies were raised in rabbits against gluthatione-S-transferase (GST) fusion proteins encoding a predicted extracellular fragment, fpEX4–6, or an intracellular fragment, fpEX11–15, of polycystin-2 (Fig. 1). Western blot analysis showed that both antibodies recognized a 110 kDa fragment in crude MDCK cell extracts, consistent with the size of polycystin-2, which was not detected with preimmune sera (Fig. 2B). For further antibody characterizations and for expression studies we generated a reporter construct, in which PKD2 was cloned in-frame to the 5' end of a gene encoding an enhanced green fluorescent protein (EGFP) resulting in a PC2–EGFP fusion protein (Fig. 2A). Western blots of MDCK cells with stable overexpression of PC2–EGFP and transiently transfected COS-1 cells (not shown) showed that our polycystin-2 antibodies recognized both the expected fragment of
140 kDa (110 kDa for polycystin-2 plus 27 kDa for EGFP) and the predicted 110 kDa endogenous polycystin-2, and that antibodies against EGFP recognized only the 140 kDa protein (Fig. 2C). Differential centrifugation of MDCK, EpH4 (data not shown), LLC-PK1 (data not shown), COS-1, HCT8/E8 and m-IMCD3 cells treated with a detergent containing lysis buffer revealed polycystin-2 in the soluble S100 fraction (Fig. 2D), as has been previously reported by Cai et al. (39).
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Immunofluorescence studies on stably or transiently transfected MDCK cells overexpressing PC2–EGFP and stained with polycystin-2 antibodies showed overlapping signals at clusters in the perinuclear region (Fig. 2E–M) and in a small subset of cells at the plasma membrane (not shown). Membrane signals were too weak to assess double staining for polycystin-1. In these cells polycystin-2 co-localized with protein-disulfide isomerase (PDI), a marker for the ER (Fig. 2K–M), which is similar to previous reports (39).
Subcellular localization of polycystin-2
To study the subcellular localization, endogenous polycystin-2 was analysed in MDCK cells using affinity-purified antibodies against fpEx11–15 and antiserum against fpEX4–6. MDCK cells are from a canine renal epithelial cell line that we have previously used to study the subcellular localization of polycystin-1 (21). Strong homogenous membrane staining was observed in the lateral membrane at different focal planes, as well as cytoplasmic staining that, depending on the focal plane, varied in intensity (Fig. 3a, c, e, g, h, k–n). This pattern was observed with both antibodies although affinity-purified fpEX11–15 antibodies gave weaker cytoplasmic signals. No signal was detected with preimmune serum for fpEX4–6 (Fig. 3d). In contrast to polycystin-1 (21) and E-cadherin, polycystin-2 was easily extracted from the membrane upon treatment with a Triton X-100 containing buffer (Fig. 3i and j) suggesting that polycystin-2 is not tightly associated to the cytoskeleton.
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Subcellular localization of endogenous polycystin-2 was also studied in a variety of other cell lines. We studied: renal epithelial cells, MDCK, m-MICD-3 and LLC-PK-1; epithelial tumour cells, EpH4 and HCT8/E8; parenchyma cells, COS-1. In all cell lines cytoplasmic staining was detected (Fig. 3o–r). In addition, EpH4 and HCT8/E8 showed homogenous plasma membrane staining (Fig. 3o and p) and in a minority of m-IMCD3 cells (Fig. 3q) and occasionally also in LLC-PK1 cells (data not shown) weak plasma membrane staining was detected.
Double-label immunofluorescence (Fig. 3e–g) and confocal laser scanning microscopy (Fig. 3h) with polycystin-2 antibodies and monoclonal antibodies against E-cadherin, an adhesion molecule localizing in the plasma membrane at adherens junctions (Fig. 3e–g), showed partial overlap. A similar staining pattern was also observed for other components of the adherens junctions (-catenin, ß-catenin, 
-catenin; data not shown). Polycystin-2 also occasionally co-localized with ZO-1 (tight junctions; Fig. 3l) and desmoplakin (desmosomes; Fig. 3k). No overlap was seen with paxillin, a marker for focal contacts (data not shown). Z-scans performed by confocal laser scanning microscopy did not reveal apical membrane staining of polycystin-2. In the cytoplasm, the endogenous polycystin-2 co-localized with the Golgi 58K protein, a marker for the Golgi apparatus (Fig. 3m). In untransfected cells we did not see overlap with PDI (Fig. 3n), a marker for the ER, while in cells expressing PC2–EGFP co-localization with PDI was observed (Fig. 2k–m).
To study the membrane localization of polycystin-2 in more detail, we performed pre-embedding immunoelectron microscopy. In MDCK cells gold-labelled polycystin-2 was seen along the plasma membrane (Fig. 3b) and in the cytoplasm near a subset of cytoskeletal filaments (data not shown). The plasma membranes of adjacent cells are in close proximity to each other at sites where adhesion complexes are localized. Between these adhesion complexes the membranes diverge, increasing the intercellular space (41). In sections with desmosomes, electron dense structures that could easily be recognized by electron microscopy, some label was present close to these structures but the majority of label was detected along the membrane between the desmosomes (Fig. 3b). Preimmune serum was negative.
Thus, we observe endogenous polycystin-2 in at least two subcellular pools, in the cytoplasm (Golgi apparatus) and in the plasma membrane and these data suggest a homogenous membrane localization for polycystin-2 and no tight association with a specific set of junction complexes.
Subcellular fractionation
To further evaluate the localization of polycystin-2 in different cellular compartments and in particular to confirm and support the plasma membrane localization, we performed sucrose density gradient fractionation experiments to separate cellular organelles. Interestingly, polycystin-2 was recovered in at least two pools with peaks at fraction 5 (Fig. 4, peak I) and fraction 15 (Fig. 4, peak II). Pool I of polycystin-2 corresponded to the peak fraction of ß1-integrin, a marker for the plasma membrane, supporting the membrane localization demonstrated by immunomicroscopy. Pool II of polycystin-2 co-migrated with markers for several cytoplasmic compartments: the Golgi apparatus (Golgi 58K protein), the ER (PDI) and actin-filaments (anti-F-actin).
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These data are in accordance with our microscopic findings that endogenous polycystin-2 is present in different pools (membrane, Golgi, presumably cytoskeleton).
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DISCUSSION |
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In 1997, two reports described possible heterodimeric interactions between the gene products of the PKD1 and PKD2 gene through predicted coiled-coil domains (22,23). This is a plausible finding, given the similar phenotype of ADPKD patients with either a PKD1 or a PKD2 mutation. Previous studies by our and other groups have established polycystin-1 as an integral membrane protein localizing at multiple adhesion complexes. We have previously demonstrated that polycystin-1 is located closely to the desmosomes in MDCK cells (16,18,19,21).
In line with the previous data, we studied the expression of polycystin-2 in MDCK cells using two polyclonal antisera raised against predicted intra- and extracellular epitopes of the protein. Polycystin-2 is located in the plasma membrane in MDCK cells as well as in HCT/E8, EpH4, mIMCD-3 and occasionally in LLC-PK1 cells. We demonstrate that polycystin-2 is not restricted to the plasma membrane but is also present in the Golgi apparatus and in the vicinity of a subset of filaments. Concordantly, upon subcellular fractionation on sucrose density gradients, polycystin-2 co-migrated in a first pool with ß1-integrin, a plasma membrane marker, and in a second pool with the Golgi apparatus, ER and the cytoskeleton (actin).
Recent reports indicated that endogenous polycystin-2 functions as a plasma membrane non-selective Ca2+-channel in human synsytiotrophoblasts (25). However, in cultured cells direct localization of polycystin-2 at the plasma membrane has not yet been reported. Heterologously expressed polycystin-2 was directed to the plasma membrane only when a putative ER retention signal of polycystin-2 was mutated, when cells were treated with chemical chaperones or when cells were co-transfected with polycystin-1 (26,27,39). We also observed localization of overexpressed full-length polycystin-2 in the ER of MDCK cells and, occasionally, in the plasma membrane, while untransfected cells did not show co-distribution with the ER. From these data we conclude that heterologously expressed polycystin-2 (PC2–EGFP) does not reflect endogenous polycystin-2 distribution in MDCK cells. Overexpression may influence protein folding and intracellular trafficking.
Analysis of a variety of cell lines demonstrates cytoplasmic expression of polycystin-2 in all cell lines and differences in plasma membrane localization. In a few cell lines, polycystin-2 seemed to be limited to the cytoplasm. This phenomenon might be explained by reduced transport of polycystin-2, by short-lived residence of polycystin-2 in the plasma membrane or because polycystin-2 in not expressed in the plasma membrane. Apparently, the subcellular localization of polycystin-2 depends on cell type and stage of differentiation. Culture conditions could be contributing factors which may explain the contrasting results from previous reports (34,39). In tissues, a punctate cytoplasmic staining was only observed in proximal tubules during early development (36) and at several extra-renal locations. In other nephronic segments, predominant basal-lateral staining has been reported, concordant with our cellular data (34,36,37).
We have previously described clear co-localization of polycystin-1 with desmoplakin (I and II), a major component of the desmosomal plaque, in MDCK cells (21). To correlate these data with the expression pattern of polycystin-2 we performed double-label immunofluorescence experiments with markers for different junction complexes, as previously reported by Scheffers et al. (21). In contrast to polycystin-1, polycystin-2 only partially co-localized with adhesion complexes. As polycystin-2 could easily be extracted from the membrane and the cytoplasm with a buffer containing Triton X-100, while E-cadherin and polycystin-1 (38) remained, we conclude that polycystin-2 is neither tightly bound to the cytoskeleton, nor anchored to cell adhesion complexes, nor to polycystin-1. Polycystin-2 apparently has a broader membrane localization compared to polycystin-1 and can move freely in certain regions of the plasma membrane.
Polycystin-2 was found to homodimerize and also be able to connect to a variety of other proteins (TRPC1, CD2AP, Hax-1, polycystin-1) (22–24,42,43). Two of these proteins, Hax-1 and CD2AP, have previously been reported to indirectly bind to or organize the cytoskeleton. HAX-1 interacts with the F-actin binding protein cortactin (43). The CD2-associated protein (CD2AP) is an adapter protein that associates with a variety of membrane proteins, organizing the cytoskeleton around a polarized site (42). We observed polycystin-2 in the vicinity of cytoskeletal filaments. Two colour immunofluorescence experiments with phalloidin to stain the actin filaments, revealed that these fibres are not the actin stress fibres (data not shown). Whether polycystin-2 co-localizes with other actin filaments cannot be excluded. Polycystin-2 along filaments could reflect transport towards the plasma membrane.
Recently, several groups reported measurable non-selective cation channel activity for polycystin-2 alone (25,26) or co-assembled with polycystin-1 (27). According to the endogenous expression patterns of polycystin-2 presented in this study we propose that polycystin-2, transported via ER and Golgi apparatus, forms a plasma membrane channel as a homodimeric/multimeric complex, or perhaps together with TRPC (24) or polycystin-1 (27). Upon binding of an as-yet-unknown ligand by polycystin-1, which is anchored to an adhesion complex, the signal transduction machinery becomes activated. This results in a temporary recruitment of polycystin-2 to form a channel together with polycystin-1, or to become activated to transport Ca2+ across the plasma membrane affecting intracellular Ca2+ homeostasis (26), and thus modulating subcellular processes (Fig. 5). Additionally, polycystin-2 may function as a chaperone to transport polycystin-1 to the plasma membrane. In ADPKD cells, impaired baso-lateral trafficking of proteins and lipids as a result of a defect in cargo exit from the Golgi apparatus, has been reported by Charron et al. (44). Disturbed balances in levels of polycystins may impair protein complex formation and thus may impair baso-lateral protein trafficking in the cystic epithelium. In short, loss of function of the polycystins may lead to the disruption of protein transport, junction complex assembly and signalling, ultimately resulting in a cystic phenotype.
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MATERIALS AND METHODS |
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Cloning of PC2–EGFP
Full-length PKD2 cDNA cloned into pBluescript KSII+ (cTM4-FL) was kindly provided by S.Somlo (Yale University, New Haven, CT). A two-step strategy was used to generate the PC2–EGFP expression construct. First, a 2.22 kb SpeI–HindIII cTM4-FL fragment was cloned into pEGFP-N1 (Clontech Laboratories, Palo Alto, CA) resulting in cTM4-N1. Secondly, a PCR was performed to generate the 3' end of PKD2 using the primer combination PKD2-ex9-F (5'-TTCCGTATCATTTTGGGCGA-3') and PKD2-2971-RHindIII (5'-CCCAAGCTTTACGTGGACATTAGAACTCCCAT-3') with cTM4-FL as a template. Digestion of the amplification product with HindIII and sub cloning into the HindIII site of cTM4-N1 resulted in PC2–EGFP, with EGFP ligated in frame to the 3' end of full-length PKD2. The reading frame was confirmed by sequencing with the sequence primer (5'-CGTCGCCGTCCAGCTCGACCAG-3'; Clontech Laboratories).
Cell culture
MDCK cells [a dog renal epithelial cell line; American Tissue Culture Collection (ATCC), Rockville, MD], COS-1 cells (green monkey kidney cell line; ATCC CRL-1650), LLC-PK1 cells (porcine epithelial cell line; ATCC CL-101) and m-IMCD3 cells (mouse inner medullar collecting duct; ATCC CRL 2123) were cultured in DMEM/F12 medium supplemented with 100 U penicillin/streptomycin, Glutamax, 10 mM HEPES, 1 mM sodium-pyruvate (all from Gibco Laboratories, Grand Island, NY) and 10% (v/v) fetal bovine serum (Pan Systems, Aidenbach, Germany). EpH4 (mouse mammary epithelial cell line) and HCT8/E8 [human colon tumour cell line; ATCC CCL24 (45)] were both provided by S.Litvinov (Leiden University Medical Center, Leiden, The Netherlands).
EpH4 was cultured in the medium described above supplemented with 5 µg/ml insulin (Sigma, Zwijndrecht, The Netherlands). HCT8/E8 cells were maintained on RPMI medium (Gibco Laboratories).
Antibodies
Anti-polycystin-2 polyclonal antibodies were raised in rabbits against GST fusion proteins: fpEX4–6 (amino acids 307–422) and fpEX11–15 [amino acids 766–990 (C-terminal end)]. Fusion proteins were generated and purified as previously described by Peters et al. (46). Ex11–15 was additionally cloned in a pET28 expression vector (Novagen, Breda, The Netherlands). The His-tagged fusion protein was affinity purified via Ni2+ immobilized columns (Novagen), coupled to Sepharose 4B (Amersham Pharmacia Biotech, Roosendaal, The Netherlands), and used to purify the polyclonal anti-fpEX11–15 serum according to standard procedures. Antiserum against fpEX4–6 was used at 1:5000 dilutions for immunofluorescence and 1:1000 for western blotting, affinity-purified antibodies against fpEX11–15 at 1:5 for immunofluorescence and 1:10 for western blotting and anti-GFP serum at 1:7500 for western blotting (a gift from Dr E.Cuppen and W.Hendriks, Nijmegen University, The Netherlands). Antibodies against ZO-1, E-cadherin and desmoplakin were used for immunofluorescence techniques as previously described by Scheffers et al. (21). Mouse monoclonal antibodies against paxillin (1:500 immunofluorescence; Transduction Laboratories, Lexington, KY) and actin (1:500 western blotting; Boehringer Mannheim, Mannheim, Germany) PDI clone 1D3 (1:50 immunofluorescence, 1:250 western blotting; Sanbio BV, Uden, The Netherlands), Golgi 58K protein clone 58K-9 (1:50 immunofluorescence, 1:2000 western blotting; Sigma), -catenin clone -CAT-7A4 (1:100; Zymed Laboratorie, San Francisco, CA), ß-catenin clone 14 (1:500; Transduction Laboratories), -catenin (1:100; Transduction Laboratories) and ß1-integrin (1:2000 western blotting; Transduction Laboratories) were used. F-actin was detected with an Alexa 568-phalloidin conjugate (1:1000; Molecular Probes, Eugene, OR).
The following secondary antibodies were used: sheep-anti-mouse IgG-fluorescein isothiocyanate (FITC) (1:50; Sigma), Alexa 594-conjugated goat-anti-rabbit (1:2000; Molecular Probes), goat-anti-rat IgG–FITC (1:50; Sigma), 10 nm gold-conjugated protein A (kindly provided by J.Onderwater, Department of Molecular Cell Biology, Leiden University Medical Center, Leiden, The Netherlands) and for western blotting donkey-anti-rabbit Ig-horse radish peroxidase (HRP) (1:10 000; Amersham Pharmacia Biotech) or rabbit-anti-mouse Ig–HRP (1:10 000; DAKO, Glostrup, Denmark).
Immunofluorescence, confocal laser scanning and pre-embedding immunoelectron microscopy
Microscopy techniques were performed essentially as previously described by Scheffers et al. (21). Cells were grown on coverslips, fixed with cold acetone/methanol (2:1) or 2% (w/v) paraformaldehyde/0.2% (v/v) Triton X-100 at 4°C for 10 min, and washed with PBS (150 mM NaCl, 8 mM Na2HPO4, 1.5 mM KH2PO4, pH 7.4). Non-fat dry milk [5% (w/v)] was used to block non-specific binding. Cells were washed with PBS and incubated with specific primary antibodies in PBS/1% BSA overnight at 4°C. Upon washing, secondary antibodies were incubated for 1 h, and washed with PBS and rinsed with distilled water. Preparations were embedded in Gelvatol. To analyse the Triton X-100 insoluble fraction, cells were treated with extraction buffer [10 mM Tris–HCl, 150 mM NaCl, 2 mM CaCl2, 1% (v/v) Triton X-100, 1% (v/v) NP-40, 1 µg/ml aprotinin, 1 mM PMSF, 10 µg/ml trypsin, 1 µg/ml leupeptin, 2 mM Benzamidin, 50 mM 
aminoCa] for 15 min prior to fixation, as previously described by Peters et al. (17). Fluorescence was analysed with a Leica Aristoplan fluorescence microscope and photographed using a Cytovision (Applied Imaging) digital system. Confocal laser scanning microscopy was performed with a Zeiss LSM 510 inverted laser scanning microscope with argon 488 nm and helium–neon 543 nm lasers (Zeiss, Weesp, The Netherlands).
For pre-embedding immunoelectron microscopy cells were grown on slides of a Lab-Tek® chamber slideTM System. Prior to fixation, cells were blocked and incubated with specific and secondary antibodies and then embedded using Epon-filled gelatine capsules at 60°C for 24 h, as previously described by Scheffers et al. (21). Ultra-thin sections were cut on a Reichert ultramicrotome FCS (Vienna, Austria) parallel to the surface of the capsule. Sections were contrasted with uranyl acetate and lead citrate, and examined using a Philips CM-10 electron microscope at 60 kV.
Transient and stable transfection
Cells were cultured to 30% confluency in six- or 24-well plates (Greiner Labortechnik, Alphen a/d Rijn, The Netherlands) and transfected with TransfastTM transfection reagent at a Transfast:DNA (charge) ratio of 1:2 according to the manufacturer’s protocol (Promega, Leiden, The Netherlands). Fluorescence of the PC2–EGFP protein was determined directly after fixation or after antibody staining as described above.
Stable cell lines were generated by linearizing the plasmid prior to transfection. Cells were cultured for 2 days, transferred to 35 mm culture dishes and selected with 0.5 mg/ml G418 for positive clones.
Cell extraction and subcellular fractionation
Cell lysates were prepared by incubating the cells for 15 min at 4°C with a buffer containing 50 mM Tris–HCl pH 7.5, 150 mM NaCl, 1% (v/v) NP-40, 0.5% (w/v) NaDOC, 1% (v/v) SDS, and protease inhibitors as described above. Homogenates were scraped, centrifuged for 10 min at 10 000 g and the supernatant was boiled in 1x Laemmli buffer.
For differential centrifugation cells were scraped, homogenized and differentially centrifuged according to the protocol described by Geng et al. (47). Briefly, cell pellets were resuspended by pottering in L-buffer (15 mM Tris–HCl pH 7.4, 120 mM NaCl, 25 mM KCl, 2 mM EDTA, 2 mM EGTA, 0.5% Triton X-100, protease inhibitors) and subjected to centrifugation for 30 min at 8000 g. The supernatant was then centrifuged at 100 000 g for 2 h. P8, S100 and P100 fractions were collected, solubilized and boiled in 1x Laemmli loading buffer containing 100 mM DTT for 2 min, loaded on an 8 or 10% SDS–polyacrylamide gel and electrotransferred to nitrocellulose membranes (HybondTM ECLTM; Amersham Pharmacia Biotech). Membranes were incubated with primary and secondary antibodies and proteins were detected using the enhanced chemiluminescence system (Amersham Pharmacia Biotech).
For subcellular fractionation, cells grown to 70% confluency in a 175 cm2 tissue culture flask (Greiner Labortechnik), were washed in buffer A (0.25 M sucrose, 10 mM Tris–HCl pH 7.4, 1 mM EDTA, protease inhibitors), carefully scraped into buffer B (buffer A containing 0.179 M sucrose supplemented with 0.75 mM KCl and 19.2 mM NaCl) and centrifuged at 1000 g to collect cells. The pellet was pottered and resuspended in buffer B prior to another centrifugation step at 1000 g. Cell debris was removed and the supernatant was layered on top of a 10 ml continuous density gradient spanning the range of 0.2–1.43 M sucrose in 10 mM Tris–HCl and protease inhibitors pH 7.4. Sedimentation of subcellular organelles occurred during overnight centrifugation at 40 000 g at 4°C using a SW40 rotor and an Optima L70k Beckman coulter ultracentrifuge (Beckman, Fullerton, CA). 500 µl fractions were collected and subjected to SDS–PAGE and western blotting as described above.
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ACKNOWLEDGEMENTS |
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The authors would like to thank S.Somlo (Yale University of Medicine, New Haven, CT) for the PKD2 cDNA construct, S.Litvinov (Leiden University Medical Center, Leiden, The Netherlands) for discussion, and S.White (Leiden University Medical Center) and R.Giles (UMC, Utrecht, The Netherlands) for critical reading of the manuscript. This research was funded by the Dutch Kidney Foundation (grant C96.1578).
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FOOTNOTES |
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+ To whom correspondence should be addressed. Tel: +31 71 5276048; Fax: +31 71 5276075; Email: d.j.m.peters@lumc.nl
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REFERENCES |
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