Human Molecular Genetics, 2000, Vol. 9, No. 18 2743-2750
© 2000 Oxford University Press
Polycystin-1, the product of the polycystic kidney disease 1 gene, co-localizes with desmosomes in MDCK cells
Department of Human and Clinical Genetics and 1Department of Pathology, Leiden University Medical Center, Wassenaarseweg 72, 2333 AL Leiden, The Netherlands
Received 3 August 2000; Revised and Accepted 4 September 2000.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Polycystin-1 is a novel protein predicted to be a large membrane-spanning glycoprotein with an extracellular N-terminus and an intracellular C-terminus, harboring several structural motifs. To study the subcellular localization, antibodies raised against various domains of polycystin-1 and against specific adhesion complex proteins were used for two-color immunofluorescence staining. In Madine Darby canine kidney (MDCK) cells, polycystin-1 was detected in the cytoplasm as well as co-localizing with desmosomes, but not with tight or adherens junctions. Using confocal laser scanning and immunoelectron microscopy we confirmed the desmosomal localization. By performing a calcium switch experiment, we demonstrated the sequential reassembly of tight junctions, subsequently adherens junctions and finally desmosomes. Polycystin-1 only stained the membrane after incorporation of desmoplakin into the desmosomes, suggesting that membrane-bound polycystin-1 may be important for cellular signaling or cell adhesion, but not for the assembly of adhesion complexes.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Polycystin-1 is the product of the polycystic kidney disease 1 (PKD1) gene. Mutations in this gene can be found in patients with autosomal dominant polycystic kidney disease (ADPKD) (1). In a minority of patients with ADPKD mutations in a second gene, PKD2, encoding polycystin-2, were found to be responsible for the disease (2). ADPKD is a systemic, rather than organ-specific, disorder. Organs such as the pancreas, ovaries and most notably the liver, can also be affected by cysts. Other manifestations can appear as well, including hypertension, cardiac valve abnormalities, intracranial aneurysms and aortic dissections (3,4). In Caucasians, 1 in 1000 individuals suffer from this disease, usually requiring treatment for end stage renal failure between the ages of 50 and 70 years.
Despite the ongoing investigations to elucidate the mechanisms behind the observed symptoms, many aspects of the disease remain poorly understood. For example, in the cystic kidney, a wide variety of abnormalities can be seen with respect to epithelial cell growth and differentiation, such as thickening of the basal membrane and abnormal polarization by altered localization of membrane proteins (e.g. Na+-K+-ATPase, ankyrin, E-cadherin and epidermal growth factor receptor). Additionally, the expression patterns of many intra- and extracellular proteins change, including adhesion molecules, growth factors, transcription factors and matrix proteins (5–8). These observations indicate that significant changes occur in the differentiation process of epithelial cells in cystic kidneys.
Cytoplasmic signaling pathways underlying cellular differentiation are generally controlled by transmembrane proteins that can bind extracellular ligands to stimulate specific intracellular processes. On the basis of computer analysis, polycystin-1 appeared to be a large (
450 kDa) membrane-spanning glycoprotein containing an extracellular N-terminus and an intracellular C-terminus. The structural motifs of the extracellular part of polycystin-1 include leucine-rich repeats, lectin-binding domains, immunoglobin-like domains and a sea urchin receptor for Egg Jelly (REJ) domain involved in sperm–egg interactions (9–12). The cytoplasmic terminus of polycystin-1 has been shown to interact through an 
-helical coiled-coil domain with a region of the C-terminus of polycystin-2 (13,14). This end of polycystin-1 is also responsible for binding heterotrimeric Gi/Go proteins, binding the regulator of G protein signaling RGS7, activating the AP-1 transcription factor and modulating Wnt signaling (15–18). These data suggest that polycystin-1 could be involved in cell–cell interactions and/or signal transduction.
To study the subcellular localization and expression of polycystin-1 in order to gain insight into its function, we analyzed Madine Darby canine kidney (MDCK) cells, a well characterized renal epithelial cell line. MDCK cells maintain their polarized architecture in confluent monolayers by distinct cellular junctions. The tight junctions, involved in barrier and fence functions, are placed at the most apical side of the epithelial cell membrane, whereas the gap junctions, involved in intracellular communication by transport of chemical compounds, are positioned at the most basal side of the membrane (19,20). The adherens junctions and desmosomes are located in between. These two types of junction consist of homophilic calcium-dependent interacting proteins, so-called cadherins, to form either cell–cell contacts or to facilitate an indirect connection to the cytoskeletal network, by binding through actin microfilaments or through intermediate filament (IF) networks (21,22).
Our results show that a fraction of polycystin-1 co-localizes with desmoplakin in the desmosomal plaque region. Furthermore, we show that de novo assembly of desmosomes starts before the re-location of polycystin-1 to desmosomes.
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RESULTS |
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Subcellular localization of polycystin-1
MDCK cells stained with polyclonal antibodies raised against fusion proteins containing epitopes of either the extracellular part of polycystin-1 (fpEX5 and fpEX15, encoded by exons 5 and 15 of PKD1, respectively) or against epitopes of the cytoplasmic tail (fpAH4, encoded by exon 46), showed a punctate line at the lateral membranes of cells in contact with other cells and staining of the cytoplasm, as shown previously (23). No signal was detected at the cell-free borders of monolayer cultures. To study the precise localization of polycystin-1 in the cell membrane, we performed two-color immunofluorescence (Fig. 1A–C and I–L) and confocal laser scanning microscopy (Fig. 1D–H) experiments with polyclonal antibodies against polycystin-1 and monoclonal antibodies raised against proteins specific for cellular junction complexes: zona occludens (ZO-1) to visualize tight junctions, E-cadherin for adherens junctions and desmoplakin (I and II) for desmosomes.
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Antibodies against desmoplakin (Fig. 1B and F) showed a highly similar membrane staining pattern to that given by antibodies against polycystin-1 (Fig. 1A and E), consequently resulting in a punctate yellow line in merged pictures (Fig. 1C, D and G). Also a manually performed pixel shift of the red and green staining strongly support the highly similar staining profiles of polycystin-1 (fpAH4) and desmoplakin (Fig. 1H).
Double staining with anti-ZO-1 and anti-fpEX5 revealed an alternating pattern of red and green lines, with occasional yellow/orange spots (Fig. 1J). Because we could focus only at the level of one color with immunofluorescence microscopy, the signals tend to overlap now and then. However, we did not see co-localization with confocal laser scanning microscopy (data not shown), thus these proteins appeared to be distributed at different levels of the cell membrane.
Antibodies against E-cadherin demonstrated a different, much more diffuse, staining pattern compared with anti-fpEX5 staining (Fig. 1I). Also in these pictures, an orange color was observed but this can be explained by the diffuse staining profile of E-cadherin. Double staining of anti-fpAH4 and anti-ZO-1 or anti-E-cadherin showed similar staining patterns as for anti-fpEX5 (data not shown).
Polycystin-1 staining showed, in addition to the membrane localization, weak to moderate cytoplasmic staining with antiserum against fpEX5 (Fig. 1A). This was more abundantly seen with the antiserum and affinity-purified antibodies against fpAH4 (Fig. 1E) (23). Pre-immune serum against fpEX5 gave virtually no signal (Fig. 1L).
When cells were extracted with a buffer containing Triton X-100 prior to fixation, the co-distribution of desmoplakin and polycystin-1 at cell–cell contacts remained (Fig. 1K). However, the punctate green and yellow colors indicate that not all polycystin-1 remained co-localized in the proximity of desmoplakin but that a fraction of the protein had been extracted. From these results, we conclude that desmoplakin and at least a fraction of polycystin-1 is connected to the triton-insoluble cell fraction, which mainly consists of the cytoskeleton and nuclear matrix-associated proteins. After this treatment cytoplasmic staining of polycystin-1 was reduced as described previously (23). E-cadherin also maintained its membrane localization on extraction, but did not co-localize with polycystin-1 (data not shown).
To demonstrate specificity of the polyclonal antibodies raised against fpAH4, we previously showed precipitation of an in vitro transcription translation product of exons 45 and 46 of polycystin-1 in a eukaryotic cell lysate (30). Now we also show immunoprecipitation of polycystin-1 from [35S]methionine/cysteine-labeled MDCK cells (Fig. 2A). A band of 400 kDa (the size of the 400 kDa laminin marker) was precipitated in MDCK cells using affinity-purified antibodies (Fig. 2A, lanes 1 and 3) or serum (data not shown). Precipitation of this protein was completely blocked when the immunizing fusion protein (30) was added (Fig. 2A, lane 2). A non-specific protein, glutathione S-transferase (GST), did not affect the precipitation (Fig. 2A, lane 3). Non-specific rabbit IgG (Fig. 2A, lane 4) or the protein A–Sepharose beads alone (Fig. 2A, lane 5) did not precipitate the 400 kDa protein. Antiserum against fpEX5 also precipitated polycystin-1 but at very low amounts (data not shown). In Figure 2B, we show an immunoblot of lyzed bacterial pellets containing an exon 5 histidine-tagged fusion protein, stained with anti-fpEX5. A highly intense protein band was seen after induction with IPTG at 35 kDa concordant with the size of this fusion protein. No fusion protein was generated without induction. Similar results were obtained with antibodies against fpAH4 on induction of an exon 46 fusion protein (data not shown). We were not able to immunoblot polycystin-1 routinely, probably due to the low levels of expression in MDCK cells.
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Junction reassembly: the Ca2+ switch
In cells cultured in medium with low Ca2+ levels, the development of calcium-mediated cell–cell adhesion, and thus junction complex assembly, is prevented (21,24). To study reassembling of the different complexes in time and to investigate a possible role of polycystin-1, we performed Ca2+ switch experiments. This type of experiment involves culturing MDCK cells in medium with low Ca2+ (<5 µM) and then increasing the Ca2+ to normal levels (1.87 mM). The switch in Ca2+ concentration has been shown to stimulate the reassembly of the junction complexes and to lead to a recovery of cell–cell interactions (21).
Cells fixed at different time points were all double stained with antibodies against E-cadherin, desmoplakin or ZO-1 (green labeled) and anti-fpEX5 or anti-fpAH4 of polycystin-1 (red labeled). Figure 3 shows a compilation of the most important cellular changes for fpEX5. Similar results were obtained for fpAH4.
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At time point zero, cells were spherical (Fig. 3A) and all proteins analyzed were located in the cytoplasm (in vesicles and more diffusely distributed) and not at the cell surface. This spherical shape also explains the intense fluorescent signal in the cells. In the course of the experiment, cells flattened, spread on a larger surface and vesicles (sometimes large) could be better identified. Most vesicles stained either for polycystin-1 or for a junctional protein. After 0.5 h cells initiated contact with neighboring cells (Fig. 3B). Tight junction assembly started at this time point, as was demonstrated by the accumulation of a small amount of ZO-1 along the cytoplasmic membrane. At time point 1 h, ZO-1 staining showed a line between two contacting cells (Fig. 3C) which remained at later time points (Fig. 3D). In contrast at time point 1 h, E-cadherin, desmoplakin and polycystin-1 staining was confined to the cytoplasm although the signals were more directed towards the membrane compared with the start of the Ca2+ switch experiment (data not shown).
The next stage of junction assembly involved the adherens junctions, as could be monitored after 1.25 h by E-cadherin staining but was better seen after 1.5 h onwards (Fig. 3 E–H). Additionally, at 1.25 h more desmoplakin-containing vesicles were observed near the cellular border (data not shown). At time point 1.5 h, a characteristic zipper-like staining pattern corresponded to the start of desmosome formation (Fig. 3I). At that time point polycystin-1 was still not accumulated at the cell membrane (Fig. 3M). Finally, at time point 1.75 h, very small amounts of polycystin-1 were detected near the desmosomes and could be better seen after 2 h (Fig. 3N). Much more polycystin-1 and desmoplakin was located in the cytoplasm. At later time points, e.g. 4 h (Fig. 3O and K) and 24 h (Fig. 3P and L), the intensity and brightness of the polycystin-1 and desmoplakin signals at the cell membrane increased dramatically, cytoplasmic staining decreased and the cells flattened. The overlapping staining pattern is shown in the merged pictures of Figure 3Q–T. Slight differences in the time course were seen between different experiments and experiments reported by others (24,25).
Immunoelectron microscopic analysis of polycystin-1 localization
Detailed studies to establish the (sub)cellular distribution of polycystin-1 were performed using electron microscopy. Antibodies indirectly labeled with 10 nm gold, were used to stain polycystin-1 either in MDCK cells grown on coverslips using a pre-embedding method or in MDCK cells allowed to aggregate in suspension using a post-embedding method. The desmosomal plaque is an electron-dense structure which can be easily recognized by its distinct morphology (42). Gold-labeled anti-polycystin-1 was observed very close to the desmosomal plaque whereas no label was detected at the membranes adjacent to the desmosomes (Fig. 4A and B). Label was also detected along filamentous structures connected to the desmosomes (Fig. 4A) or partly dispersed throughout the cytoplasm by associated with membranous structures (Fig. 4B). In the aggregated MDCK cells additional large vesicles were present in which accumulation of gold was seen (Fig. 4C). Desmosomal localization was confirmed in double labeling experiments with polycystin-1 (10 nm gold; Fig. 4E and F) and proteins connected to the desmosomes, desmoplakin (5 nm gold; Fig. 4E) and keratin (5 nm gold; Fig. 4F). Gold particles of both sizes were found in close proximity at the desmosomal region. Overall, most of the polycystin-1 label was located in the cytoplasm or in cytoplasmic vesicles and less label could be observed near the cytoplasmic membrane. This is in agreement with the immunofluorescence data which showed that the membrane localization was confined to a very discrete band, the desmosomal structures, and that cytoplasmic vesicles were stained throughout the cell. Hardly any gold particles were seen in sections labeled with pre-immune serum (Fig. 4D).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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This study was aimed at gaining a better understanding of the expression and subcellular localization of polycystin-1 in MDCK cells, a well characterized renal epithelial cell line. We previously showed that antibodies against predicted intra- and extracellular epitopes placed polycystin-1 in the cytoplasm and in a punctate line along the plasmic membrane at cell–cell contacts (23). This is concordant with the predicted transmembrane structure of polycystin-1 (9).
In polarized epithelial cells, structural and functional organization is established and maintained by several junction complexes. To investigate whether polycystin-1 co-localizes with known cell–cell junction complexes we performed double label experiments with antibodies against polycystin-1 together with antibodies against components of the tight junctions, adherens junctions and desmosomes. We clearly observed co-localization of polycystin-1 with desmoplakin (I and II), proteins expressed in the cytoplasmic region of the desmosomal plaque, and not with proteins connected to the tight junctions or the adherens junctions. Even after extraction with a Triton X-100-containing buffer, a fraction of polycystin-1 remains anchored to the cytoskeleton together with desmoplakin. Electron microscopy confirmed polycystin-1 localization on or near to filamentous structures and at the cytoplasmic side of the desmosomal plaque region. Antibodies against the predicted extracellular epitope encoded by exon 5 also labeled the cytoplasmic side of the desmosome. This result could signify a temporary localization of polycystin-1, prior to incorporation in the membrane. Furthermore, anti-polycystin-1 label was observed near membranous structures in the cytoplasm and in cytoplasmic vesicles. Using immunoelectron microscopy, staining at the cell membrane (basolateral and apical) and/or around small subcellular membranous structures in 18-week-old fetal kidneys and G401 cells has been described by others (26,27).
The establishment of homotypic adhesion between neighboring cells is a stepwise process in which Ca2+-dependent interactions play an important role (24,28). To induce assembly of junctions in MDCK cells we performed Ca2+-switch experiments. In cells cultured in low Ca2+ medium, tight junction, adherens junction and desmosomal proteins were diffusely distributed in the cytoplasm and in vesicles. On switching the Ca2+ concentrations to normal levels, rebuilding of junctions was synchronously induced. A step-wise assembly into cell membranes was seen in time: tight junctions showed up after 0.5 h, adherens junctions followed after 1.25 h and desmosomes started to form after 1.5 h. At these time points assembling of the different complexes was in an initial phase, since we still observed cytoplasmic staining of all proteins tested. Our data are in agreement with experiments described by others previously (24,25). Polycystin-1 appeared at the lateral membrane, just after the assembly of desmosomes was started (Fig. 3). These data strongly suggest that desmosomal proteins are first transported to the cell surface, followed by polycystin-1. Our experiments also revealed hardly any cytoplasmic co-alignment with desmoplakin close to the membrane, suggesting a transport mechanism distinct from that for desmoplakin. Although these data argue against an involvement of polycystin-1 in the (initial) formation of desmosomes, a possible functional role cannot be excluded.
Studies on the function of polycystin-1 have proposed a role in signal transduction. It has been shown that the C-terminal domain of polycystin-1 can bind heterotrimic G proteins, suggesting a G protein-coupled receptor action (17). Interestingly, interaction of the C-terminus with a regulator of G protein signaling (RGS7) has also been described recently by Kim et al. (18). The precise signaling cascades influenced by polycystin-1 are not yet clear but the C-terminal region can elevate AP-1 promoter activity and can modulate Wnt signaling (15,16). These data together with the predicted protein-binding domains in the extracellular part of polycystin-1 suggest that a variety of intracellular reactions may be triggered on binding of a ligand.
We propose that polycystin-1 needs anchoring to the desmosomes in order to function as a signal transduction molecule rather than being a functional desmosomal component. Binding of membrane-bound heterotrimeric G proteins on activation of polycystin-1 could be a critical step. Alternatively, polycystin-1 may function as a (homotypic) adhesion molecule, as recently suggested by Ibraghimov-Beskrovnaya et al. (43), possibly supported by other adhesion complexes. Interestingly, co-localization or co-precipitation of polycystin-1 with E-cadherin has been observed in fetal renal tissue (40,41) and with the platelet endothelial cell adhesion molecule 1 (PECAM-1) in human umbilical vein endothelial cells (HUVECs) (29), supporting our hypothesis of adhesion complex association. Endothelial cells express different junction complexes compared with epithelial cells. These cells have no desmosomes but PECAM-1, belonging to the immunoglobulin superfamily of adhesion molecules, is a major constituent of the endothelial cell junctions. These data suggest that, depending on the cell type and stage of differentiation, polycystin-1 could be anchored to different junctional complexes.
Expression and subcellular localization studies of polycystin-1 in a large number of tissues and specific cell types seemed to lead to conflicting results. However, localization of polycystin-1 appears to be developmentally regulated and to be dependent on the differentiation state of cells. For example, mostly apical/basolateral distribution of polycystin-1 is observed in young fetal kidney tissues, whereas cytoplasmic staining is seen in more mature fetal and adult kidneys with the latter more reduced in intensity (26,27,29–35,40,41). The localization of polycystin-1 in the cytoplasm and in membranes either could be a reflection of a precursor pool of polycystin-1 transported towards the cell membrane, as we see during the Ca2+ switch, or could mean different functions of polycystin-1 in different cellular compartments.
In summary, this study shows that polycystin-1 is associated with desmosomes in MDCK cells, but we have found no evidence for an involvement in initial desmosome assembly. We postulate that polycystin-1 anchorage to adhesion complexes is needed for signal transduction, in order to maintain cell adhesion, protein sorting and cell polarity, all of which are features disrupted in cystic epithelium.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Cell culture
MDCK cells (ATCC, Rockville, MA) were cultured in DMEM/F12 medium supplemented with 100 U penicillin/streptomycin, Glutamax, 10 mM HEPES, 1 mM sodium pyruvate (Gibco BRL, Grand Island, NY) and 10% fetal bovine serum (Pan Systems, Aidenbach, Germany).
Antibodies
Anti-polycystin-1 polyclonal antibodies were raised in rabbits against three fusion proteins of gluthatione S-transferase (GST) and a part of exon 5 (fpEX5, amino acids 251–465, 1:1600), exon 15 (fpEX15, amino acids 1571–1979, 1:1600) and exon 46 (fpAH4, amino acids 4288–stop, 1:1600; affinity- purified, 1:2) have been described previously (23,30). In addition Ex5 and Ex46 were cloned in a pET28 expression vector containing a his tag (Novagen, Breda, The Netherlands). For affinity purification of fpAH4 we used the fusion construct pMAL-BD3 encoding amino acids 4097–4302 (29) (kindly provided by O. Ibraghimov-Beskrovnaya, Genzyme, Framingham, MA). The protein was purified on amylose column according to standard procedure (New England Biolabs, Leusden, The Netherlands).
For the detection of junction complexes rat monoclonal antibodies against ZO-1 (1:50; Zymed Laboratories, South San Francisco, CA), mouse monoclonal antibodies against E-cadherin (1:200; Thamer Diagnostica, Uithoorn, The Netherlands) and desmoplakin (I and II) clone 115F (1:50; kindly provided by D. Garrod, University of Manchester, UK) were used. For electron microscopy studies monoclonal antibodies against desmoplakin (described above) and clone 80 of keratin (Monosan, Uden, The Netherlands) were used.
The following secondary antibodies were used: sheep anti-mouse IgG–fluorescein isothiocyanate (FITC) (1:50; Sigma, Zwijndrecht, The Netherlands), Alexa 594-conjugated goat anti-rabbit (1:2000; Molecular Probes, Eugene, OR), goat anti-rat IgG–FITC (1:50; Sigma), 10 nm gold-conjugated protein A (kindly provided by J. Onderwater, Department of Electron Microscopy, Leiden University Medical Center), 10 nm gold-conjugated goat anti-rabbit (Amersham Pharmacia Biotech, Roosendaal, The Netherlands), 5 nm gold-conjugated goat anti-mouse (Amersham Pharmacia Biotech) and for western blotting donkey anti-rabbit Ig–horse radish peroxidase (Pharmacia Biotech, Uppsala, Sweden).
Immunoprecipitation
MDCK cells were labeled for 4 h with 0.1 mCi/ml 35S-labeled methionine/cysteine (Amersham) in methionine-free medium with dialyzed fetal bovine serum (FBS) (Gibco BRL). Cells were lyzed in buffer containing 150 mM NaCl, 1% (v/v) NP-40, 0.5% (m/v) DOC, 1% (v/v) SDS, 50 mM Tris–HCl pH 7.5 and subsequently centrifuged at 10 000 g for 15 min at 4°C. Lysates were pre-cleared by incubation with protein A–Sepharose (Pharmacia Biotech) overnight. After centrifugation, lysates were incubated with protein A–Sepharose beads conjugated to immune serum from rabbits immunized with fusion proteins against polycystin-1, pre-immune serum, total purified rabbit IgG (0.02 mg/ml), purified antibodies plus fusion protein (0.1 mg/ml) or GST for 6 h. Beads were removed by centrifugation at 5000 g for 30 s at 4°C and washed four times with lysis buffer.
Immunoprecipitates were separated on a 6% SDS–polyacrylamide gel and examined by autoradiography using a PhosphorImager (Molecular Dynamics, Sunnyvale, CA). Protein sizes were compared with a protein size marker (Benchmark; Gibco BRL) and laminin-1 (EHS-laminin; Sigma, St Louis, MO).
Ca2+ switch
The Ca2+ switch protocol was essentially performed as described by Rajasekaran et al. (24). Briefly, a single cell suspension (6 x 105) was seeded on APS (3-aminopropyltriethoxysilane)-coated glass coverslips (Sigma) in 24 well plates and allowed to attach in DMEM/F12 medium containing 1.8 mM Ca2+ for 2 h. Thereafter, cells were washed twice in SMEM medium (Gibco BRL), which lacks CaCl2 and contains <5 µM Ca2+, and subsequently incubated in SMEM medium containing 5% fetal calf serum [dialyzed against phosphate-buffered saline (PBS) and 0.2 mM EDTA] for 18 h. The Ca2+ switch was performed by replacing SMEM by DMEM/F12 medium. At different time points, cells were fixed with cold acetone:methanol (2:1) at 4°C for 10 min and processed for immunofluorescence microscopy as described below.
Immunofluorescence and confocal laser scanning microscopy
Cells were grown on coverslips in 24 well plates to 50–80% confluency, acetone:methanol fixed as described above, and washed with PBS (150 mM NaCl, 8 mM Na2HPO4, 1.5 mM KH2PO4, pH 7.4). Non-fat dry milk (5%) was used to block non-specific staining of fixed cells at room temperature for 1 h. Subsequently, cells were washed with PBS and incubated with specific primary antibodies in PBS/1% bovine serum albumin (BSA) overnight at 4°C. Bound antibodies were detected with Alexa 594 goat anti-rabbit (1:2000) IgG and sheep anti-mouse IgG–FITC (1:50) or goat anti-rat IgG–FITC (1:50) for 1 h and washed with PBS and distilled water. To analyze the Triton X-100 insoluble fraction, cells were treated with extraction buffer (10 mM Tris–HCl, 150 mM NaCl, 2 mM CaCl2, 1% Triton X-100, 1% NP-40, 1 µg/ml aprotinin, 1 mM PMSF, 10 µg/ml trypsin, 1 µg/ml leupeptin) for 15 min prior to fixation. Final preparations were embedded with Gelvatol and analyzed with a Leica Aristoplan fluorescence microscope and photographed using Cytovision (Photometrics, Osnabrück, Germany) digital system. Confocal laser scanning microscopy was performed on cells with the BioRad (Hercules, CA) MRC1024 ES krypton–argon ion laser scanning imaging system with a 585 EFLP and 522 DF35 emission filters.
Pre-embedding immunoelectron microscopy
Cells were grown on coverslips, fixed, blocked and incubated with specific primary antibodies overnight as described above. Cells were subsequently washed in PBS, incubated with 10 nm protein A gold electron-dense marker (2 h), washed in PBS, fixed in 1% glutaraldehyde in 0.1 M cacodylate buffer, pH 7.4, for 5 min at 4°C and postfixed in 1% osmium tetroxide for 10 min at room temperature. Fixed cells were dehydrated in a graduated series of ethanol (30–100%) and embedded using an Epon-filled gelatin capsule at 60°C for 24 h. Coverslips were released by immersing the capsule in liquid nitrogen. Ultra-thin sections were cut on a Reichert ultramicrotome FCS (Vienna, Austria), contrasted with uranyl acetate and lead citrate, and examined using a Philips CM-10 electron microscope at 60 kV.
Post-embedding, ultra-cryo immuno-electron microscopy
Cell aggregates were fixed in 4% paraformaldehyde, 0.05% glutaraldehyde in sodium bicarbonate buffer, pH 7.4, for 1 h at room temperature according to Litvinov et al. (36). Aggregates were sedimented at 60 g for 2 min and embedded in 10% gelatin, PBS. Small blocks of gelatin-embedded cells were soaked with 2.3 M sucrose, placed on a specimen holder and frozen in liquid nitrogen. Ultrathin cryosections were collected on carbon-coated pioloform films on copper grids. Grids were transferred to 2% gelatin (on ice), temperature raised to 40°C, sections incubated with 20 mM glycine in PBS for 20 min and pre-incubated in 1% BSA in PBS for 30 min. The sections were then incubated with primary antibodies and 10 nm gold-conjugated protein A or goat anti-rabbit and 5 nm gold-conjugated goat anti-mouse as previously described (37,38). Ultra-thin cryosections were washed and fixed in 1% glutaraldehyde in PBS for 5 min and stained and embedded with a mixture of 3% uranyl acetate diluted 1:10 with 2% methyl cellulose as previously described (39). Sections were examined using a Philips CM-10 electron microscope at 6 kV.
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ACKNOWLEDGEMENTS |
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The authors would like to thank G. Lamers and J. Slats for technical support as well as R. Giles for critical reading of the manuscript. This research was funded by the Dutch Kidney Foundation (grants C96.1578 and C95.1477).
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FOOTNOTES |
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+ To whom correspondence should be addressed. Tel: +31 715 276 048; Fax: +31 75 276 075; Email: d.j.m.peters@lumc.nl
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REFERENCES |
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