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Vol. 16, Issue 9, 4398-4409, September 2005
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* Department of Medicine, Vanderbilt University, Nashville, TN 37232;
@ Department of Cell and Developmental Biology, Vanderbilt University, Nashville, TN 37232;
¶ Department of Cancer Biology, Vanderbilt University, Nashville, TN 37232;
Department of Genetics and Developmental Biology, University of Connecticut, Farmington, CT 06030;
Department of Bioengineering, University of California-Berkeley, Berkeley, CA 94720;
|| Laboratory of Translational Cancer Research, Cancer Institute and Hospital, Chinese Academy of Medical Sciences, Beijing 100021, China;
# Department of Research Medicine, Veterans Administration Hospital, Nashville, TN 37232; and
Department of Cardiology, First Affiliated Hospital, Sun Yat-sen University, Guangzhou 510080, China
Submitted November 22, 2004;
Revised May 16, 2005;
Accepted June 15, 2005
Monitoring Editor: Keith Mostov
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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; Zerres et al., 1998). The major clinical manifestations of this disease are ectasia of renal collecting and hepatic biliary ducts as well as fibrosis of both the liver and kidneys (Potter, 1972; Bosniak and Ambos, 1975). By genetic linkage analysis, the gene responsible for this disease, termed polycystic kidney and hepatic disease 1 (PKHD1), was mapped to human chromosome 6p (Mucher et al., 1994; Guay-Woodford et al., 1995) and was recently cloned by several independent groups (Onuchic et al., 2002; Ward et al., 2002; Xiong et al., 2002). PKHD1 is speculated to encode a receptor-like protein (Ward et al., 2002) and may be composed of distinct exons that generate a number of isoforms (Onuchic et al., 2002; Xiong et al., 2002). The longest ORF of PKHD1 (PKHD1-FP) contains 67 exons and encodes a 4074-amino acid (AA) membrane-associated protein, termed fibrocystin/polyductin (FPC). In addition, our group identified a short form of PKHD1, named PKHD1-tentative (PKHD1-T), which encodes a 3396-AA protein, termed tigmin (FPT; Xiong et al., 2002).
PKHD1 encodes a very large and complex protein with only a few recognizable motifs and a single predicted transmembrane domain (Onuchic et al., 2002; Ward et al., 2002; Xiong et al., 2002). Using a panel of antibodies, it was shown that PFC is widely expressed in epithelial derivatives, which form the primary duct system during embryogenesis and organogenesis (Nagasawa et al., 2002; Zhang et al., 2004). In the kidney, FPC predominantly localizes to the apical domain of renal tubule cells, where it associates with the basal bodies/primary cilia (Masyuk et al., 2003; Ward et al., 2003; Menezes et al., 2004; Wang et al., 2004; Zhang et al., 2004).
A well known approach in studying the tubulomorphogenic processes is the use of three-dimensional (3-D) epithelial cell cultures in extracellular matrix (ECM) gels (Zegers et al., 2003). To unravel the role of FPC in regulating renal epithelial function, we used stable Pkhd1-silenced IMCD cells to characterize its functions in tubulomorphogenesis, cell proliferation, apoptosis, motility, polarization, and cytoskeletal organization. Our results indicate that the expression of normal PFC is required to sustain renal tubulogenesis/tubulomaturation in 3-D cultures of IMCD cells. Lack of this protein resulted in abnormalities in cell-cell contact, actin cytoskeleton organization, cell-ECM interactions, cell proliferation, and apoptosis. Pkhd1-silenced IMCD cells showed delayed and diminished ERK1/2 and FAK pY861 activation after cell adhesion to ECM, suggesting that FPC may mediate tubulomorphogenesis of IMCD cells via ERK and FAK signaling pathways. The observations made in vitro may illuminate the characteristics of ARPKD phenotypes in vivo.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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-tubulin (Sigma), p-Akt/total Akt (Cell Signaling Technology, Beverly, MA), p-ERK/total ERK (Cell Signaling Technology), FAKpY397, 407, 576, 577, 861 (BioSource, Camarillo, CA), and total FAK (BD Transduction Laboratories) antibodies were purchased. Polyclonal and monoclonal antibodies against FPC, including hAR-Np, hAR-C2p, hAR-Nm3G12, hAR-Cm3G6, and hAR-C2m3C10, were described in our previous study (Zhang et al., 2004). Secondary antibodies included Cy3-conjugated rabbit anti-mouse IgG, Cy2-conjugated goat anti-rabbit IgG (Jackson ImmunoResearch Laboratories, West Grove, PA), Alexa Fluor 488 goat anti-mouse IgG, and Alexa Fluor 594 goat anti-rabbit IgG (Molecular Probes, Eugene, OR).
METHODS
Cell Culture and Transfection. Culture conditions for IMCD cells and the tubulogenesis assay for the IMCD-derived cells in 3-D ECM gels were as previously described (Zent et al., 2001; Chen et al., 2004). The collagen I (CI) gels were composed of 1 mg/ml CI in Dulbecco's minimal essential medium containing 20 mM HEPES (pH 7.2). The Matrigel/collagen I gels (MG) is 1:1 mixture of CI and MG with a final concentration of 0.5 mg/ml for CI and 0.5 mg/ml for Matrigel (Chen et al., 2004). Ten percent fetal calf serum (FCS) was used for GI and MG gel cultures. The tubule formation was determined in five randomly picked high-power fields (Figure 1, A-D). The cells used to establish normal cell-cell contacts were plated and grown to confluence for at least 3 d on 12-mm transwell plates (Costar, Cambridge, MA). Transient transfection assays were performed in 24- or 6-well culture plates (Costar) with Lipofectamine 2000 (Invitrogen, Carlsbad, CA). The procedures for transfection were performed according to the manufacturer's protocol. All cell culture reagents were purchased from Life Technologies (Invitrogen).
Small Interfering RNA and shRNA Stable Cell Lines. Double-stranded small interfering RNAs (siRNAs) were synthesized by Dharmacon Research (Boulder, CO; Table 1). Subconfluent populations of IMCD cells were transfected using Lipofectamine 2000 according to the manufacturer's instructions. Tubulogenesis assays were performed 24 h after the transfection. To establish stable Pkhd1-silenced and control IMCD cell lines, Pkhd1 siRNA3 and other control constructs (Figure 2A) were transiently transfected into subconfluent IMCD cells. Twenty-four hours later, 5% FCS DMEM/F12 along with G418 at a concentration of 1 mg/ml was used to select for G418-resistant clones. After a week of G418-selective culture, the remaining cells were resuspended and seeded in 100-mm2 culture plates (Costar) with a cell density of 103 per plate. When G418-selected colonies were formed, single colonies were picked using inverted microscopy and transferred into a new set of 24-well cultured plates (Costar).
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RNA Isolation and Quantitative PCR. Total RNA was isolated from the IMCD-derived transient transfected cells and stable cell lines using Trizol reagents (Invitrogen) according to the manufacturer's instructions. Quantitative PCR was performed using the iCycler iQ Real Time PCR Detection System (Bio-Rad, Richmond, CA). A pair of primers was designed to bridge exons 34 and 35 in Pkhd1 cDNA. The forward primer, 5'-GGC TTT CCT ATG TGA CCT G-3', and reverse primer, 5'-trichloroacetic acid CAC TCC ATC TCT GCC TC-3' were used for real-time PCR with the iQ SYBR Green Supermix kit (Bio-Rad).
Western Blotting, Immunofluorescence Staining, and Flow Cytometry. The different IMCD cell populations were serum starved for 12 h and detached from the plates with trypsin. The trypsin was inactivated by the addition of 1 mg chicken egg white trypsin inhibitor/ml (Sigma). Cells were then centrifuged and resuspended in serum-free medium. One part of the cells was kept in suspension for 30 min at 37°C, while the other part was replated on collagen I (10 µg/ml) for 10 and 30 min at 37°C, as previously described (Hanks et al., 1992; Cai et al., 2005). Cells were then washed twice with phosphate-buffered saline (PBS) and lysed using RIPA buffer (50 mM Tris-HCl, pH 7.4, 1% Triton X-100, 0.25% sodium deoxycholate, 150 mM NaCl, 1 mM EDTA, protease inhibitor cocktail, 1 mM Na3VO4, 1 mM NaF) for 20 min. Lysates were clarified by centrifugation at 12,000 x g for 10 min at 4°C. Total protein, 20 µg, was run onto an SDS gel and subsequently transferred to nitrocellulose membranes. Membranes were blocked in 5% milk/tris-buffered saline Tween and then incubated with the different primary antibodies followed by the appropriate HRP-conjugated secondary antibodies. Immunoreactive bands were identified using enhanced chemiluminescence according to the manufacturer's instructions.
For immunofluorescence (IF), cultured cells were first washed with 1x PBS twice and fixed with 4% paraformaldehyde at 4°C for 30 min. The cells were washed twice more with 1x PBS and incubated with 1% bovine serum albumin (BSA) at room temperature for 1 h. The cells were then incubated with primary antibodies for 2 h and washed again with 1x PBS three times. Washed cells were treated with fluorescence-conjugated secondary antibodies for 1 h.
For flow cytometry, a suspension of tested cells was incubated with fluorescent reagents provided in the Apoptosis Kits (BioVision Research Products). Flow cytometry was performed with a FACScan instrument (Becton Dickinson, San Jose, CA).
Confocal Microscopy, Time-lapse Phase-Contrast Microscopy, and Scanning Electron Microscopy. For confocal microscopy, the IF images were collected as Z-series sections using a Zeiss LSM 510 confocal microscope system (Thornwood, NY) with a 10x or 40x oil objective. Multiple sections (
0.3-0.5 µm thick) were projected onto one plane for presentation.
Time-lapse microscopy video was performed using a Nikon inverted microscope (TE300; Melville, NY) equipped with a 10x objective. A temperature hood was built around the microscope to keep the temperature at 37°C during experiments. Phase-contrast images were collected using a Hamamatsu Orca100 cooled digital CCD camera (Bridgewater, NJ) at 20-min intervals for 20-24 h and transferred directly from a frame grabber to computer storage using C-Imaging System software (Compix, Cranberry Township, PA). Cell outlines were visualized by phase contrast microscopy with a 10x objective. A scanning stage allowed image collection from different areas of the sample automatically.
For scanning electron microscopy, samples were fixed in 2% glutaraldehyde in 0.1 M sodium cacodylate for 1 h followed by treatment with 1% osmium tetroxide in 0.1 M sodium cacodylate buffer. After an ethanol dehydration series, the samples were critical-point dried and sputter-coated with 40% gold, 60% palladium microparticles to a thickness of 15-17 nm. The images of the samples were collected by using an Electroscan E3 Environmental Scanning Electron Microscope (Hitachi S-5000, Hitachi Science, Ibaraki, Japan).
Cell proliferation and Apoptosis. IMCD cells (3 x 103) were embedded in the 3-D gels (100 µl final volume) in 96-well plates as described above and incubated in DMEM/F12 containing 5% FCS. After 2 d in culture, cells were pulsed for an additional 48 h with [3H]thymidine (1 µCi/well). The gels were then removed from the plates and dialyzed against PBS for 24 h to remove free [3H]thymidine. The cells in the gels were then lysed in 1% SDS (100 µl final volume) and the lysates were measured with a 
counter. In addition, the Quick Cell Proliferation Assay Kit (BioVision Research Products) was used to analyze the proliferation rate among the cell lines. This assay is based on cleavage of the tetrazolium salt WST-1 to formazan by cellular mitochondrial dehydrogenases. Expansion of the number of viable cells results in an increase in the activity of the mitochondrial dehydrogenases (Berridge et al., 1996), leading to an increase in the amount of formazan dye, which can be detected by spectrometry.
For apoptosis studies, the cells were incubated in 3-D gels as described above. After 7 d in culture, the gels were fixed in 4% paraformaldehyde for 30 min, followed by dimethyl sulfoxide/methanol at a ratio of 1:1 for 30 min. Apoptosis was detected using the Apoptag Apoptosis Detection Kit as described by the manufacturer (Serologicals, Norcross, GA). The gels were also stained with DAPI to identify all the cell nuclei. Apoptotic cells were counted in at least 10 different microscopic fields of a fluorescence microscope and the apoptotic index, expressed as (number of positive apoptotic cells/200 counted cells) x 100, was determined. We also quantified apoptosis using an Annexin V-PE Apoptosis Kit (BioVision Research Products), in which Annexin V is conjugated to PE as the florescent marker (An and Huang, 2004) to evaluate apoptotic rates by flow cytometry.
Cell Adhesion and Migration Assays. The 96-well cell culture plates (Nunc, Napierville, IL) were coated with CI at the indicated concentrations in PBS for 12 h at 4°C. The negative controls were performed on plates coated with 1% BSA. Positive controls were the cells plated onto tissue culture plates in the presence of 10% FCS. The plates were then washed with PBS and incubated with PBS containing 1% heat-denatured BSA for 60 min to block nonspecific adhesion. Aliquots (100 µl) of single-cell suspensions (106 cells/ml) in serum-free DMEM/F12 containing 0.1% BSA were added in triplicate to 96-well plates with 0.01-2 µg/ml CI and incubated for 60 min at 37°C. Nonadherent cells were removed by washing the wells with PBS. Cells were then fixed with 4% formaldehyde, stained with 1% crystal violet, solubilized in 20% acetic acid, and the OD of the cell lysates was read at 570 nm. Cells bound to FCS were used as a positive control to indicate maximal cell adhesion, and the amount of cells bound to CI-uncoated wells (BSA only) was used as the background, and this OD was subtracted from that obtained with serum or ECM proteins. We evaluated the cell-matrix adhesion by the formula: (OD value of tested cells minus OD of background/OD of positive controls minus OD of background) x 100.
Transwell filter migration assay was performed in polyvinylpyrolidone-free polycarbonate filters with 8-µm pores (Costar). The underside of the transwell was precoated with 5 µg/ml CI overnight at 4°C and the filter was subsequently blocked with 1% BSA for an hour at 37°C to inhibit nonspecific migration. Aliquots (100 µl) of cell suspension (1 x 106 cells/ml) in serum-free medium were added to the wells, and cells were allowed to migrate into the matrix coated on the underside of the transwell for 3 h. Cells on the top of the filter were removed by wiping and the filter was then fixed in 4% formaldehyde in PBS. Migrating cells were stained with 1% crystal violet and five randomly chosen fields from triplicate wells were counted at 200x magnification.
A wound-healing assay was used to test the cell migration in the IMCD cell lines and the cells (104 cells/well) were seeded in 24-well culture plates with low serum (0.5%). After 2 d of culture, the confluent monolayers were wounded by scraping a 10-15-µm line with a flattened needle across the cell monolayer. The status of the wound line was then documented by time-lapse microscopy at 20-min intervals for >12 h.
Histological Sectioning of Cultured Gels. The 3-D cultured samples were washed with 1x PBS twice, then fixed with 4% paraformaldehyde, and incubated at room temperature for 20 min. After a regular ethanol dehydration series, the samples were treated twice with xylene for 20 min each time and then soaked in 1:1 xylene/paraffin for 30 min followed by paraffin alone for 30 min. The paraffin-embedded samples were then sectioned to a thickness of 6 µm using a Leica 2135 microtome (Deerfield, IL).
Transepithelial Resistance. The tested cells were plated on transwell filters (12 well, 0.4-µm pore size, Corning Costar) with 1 x 105 cells per well and allowed to attach overnight, to form a confluent monolayer in normal culture medium. Transepithelial resistance (TER) was measured after 24 h of plating and every 24 h thereafter using an EVOM/STX2 (World Precision Instruments, Sarasota, FL) electrical resistance measurement system with the results expressed in 
/cm2.
Statistics. All assays were repeated at least three times in duplicate or triplicate and the graphic data were presented as the mean ± SD. Statistical analysis was performed where appropriate using the Student's t test or one-way analysis of variance (ANOVA) followed by the Tukey's Multiple Comparison Test. Differences with p < 0.05 were considered statistically significant.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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). To test whether silencing FPC would induce abnormal tubulomorphogenesis in 3-D IMCD cell cultures, we selected at least three siRNA duplexes based on the mouse Pkhd1 cDNA sequence using Oligoengine (Table 1; Zhang et al., 2004). The cDNA fragments for these siRNA duplexes were first used as the query sequence in a BLAST search on the NCBI database to confirm the absence of homology for either humans or mice. Each duplex was then transiently transfected into IMCD cells and 24 h later the cells were placed into 3-D ECM gels for the tubulogenesis assay. The cells transfected with any of the individual siRNA duplexes (Table 1) exhibited decreased Pkhd1-mRNA levels with Pkhd1siRNA3 resulting in the most Pkhd1 inhibition (Figure 1G). A combination of all three siRNA duplexes (siRNA1-3) showed the largest amount of inhibition when compared with individual transfections. Tubulogenesis failure (Figure 1, C and D) was present in 10% in wild-type IMCD cells; however, it was 85% in Pkhd1siRNA1 cultures, 50% in Pkhd1siRNA2 cultures, 85% in Pkhd1siRNA3 cultures, and 90% in combined Pkhd1siRNA1-3 cultures (Figure 1H). Similar results were seen when the cells were seeded in MG gels (Figure 1I). These findings support the idea that silencing Pkhd1 arrests tubulogenesis in 3-D cultured IMCD cells.
Establishment and Characterization of Stable Pkhd1-silenced IMCD Cell Lines
As inhibition of Pkhd1 mRNA in the IMCD cells transiently transfected with the Pkhd1siRNA3 duplex were profound (Figure 1G), this duplex was selected for constructing a Pkhd1shRNA vector, designated the Pkhd1shRNA3 clone (Figure 2A). In parallel, we constructed a control vector, Pkhd1shRNA3M, in which two nucleotides in the middle of the duplex were randomly mutagenized from GC to AA (Figure 2A). We transfected the Pkhd1shRNA3, Pkhd1shRNA3M constructs as well as an empty shRNA vector into IMCD cells to produce stable cell lines designated as IMCDshRNA3, IMCDshRNA3M, and IMCDsh (Figure 2A).
We then isolated 12 single G418-resistant clones and performed Western blot analysis and IF staining to select clones of IMCDPkhd1shRNA3 transfected cells in which the expression level of FPC was reduced. Of the 12 selected IMCDPkhd1shRNA3 cell clones, where the Pkhd1shRNA3 construct was confirmed by PCR, at least four demonstrated significantly decreased expression of Pkhd1 mRNA. Two of them, e11 (IMCDshRNA3e11) and e23 (IMCDshRNA3e23), were chosen for further analysis and were eventually used for the functional studies of FPC. Using a similar approach, two control cell lines were established from the IMCDshRNA3M transfected cells, named IMCDshRNA3M5 and IMCDshRNA3M9, and another two empty-vector control cell lines were also established and named IMCDsh15 and IMCDsh21 (Figure 2A).
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), IF cell staining showed that the two Pkhd1-silenced IMCD cell lines, IMCDshRNA3e11 and IMCDshRNA3e23, exhibited markedly less staining of the basal bodies/cilium than the IMCD wild-type cells (IMCDWT; Figure 2, B-H). The staining patterns in the vector control cell lines (IMCDsh15 and IMCDsh21) were similar to the pattern in IMCDWT (Figure 2, B and E). Western blot analysis using both mono- and polyclonal antibodies showed that these two stable Pkhd1-silenced cell lines exhibited greatly decreased levels of FPC (Figure 2L). These results provide evidence that FPC was down-regulated in the stable cell lines mediated by the shRNA approach. Consistent with results from Western analyses, a 4-5-fold decrease in the Pkhd1 mRNA level was also seen in the IMCDshRNA3e11 and IMCDshRNA3e23 cells by quantitative PCR, but not in the control cell lines (p > 0.05; Figure 2M). In contrast, there was only a 25% decrease in Pkhd1 mRNA levels (Figure 2M) and relatively less Pkhd1-staining (Figure 2F) in IMCDshRNA3M5 cells. This result implies that the mutagenized Pkhd1-shRNA construct may produce micro-RNA (miRNA) against Pkhd1 because of its nine identical nucleotides for Pkhd1 in the construct (Figure 2A). Given that the empty shRNA-vector cell lines did not demonstrate any inhibition effects, whereas mutagenized Pkhd1-shRNA and true Pkhd1-silenced constructs (Pkhd1shRNA3) resulted in minor and strong inhibition, respectively, we can conclude Pkhd1shRNA3 silencing activity is specific.
FPC Is Required for Normal Tubulogenesis in 3-D Culture of IMCD Cells
To confirm the finding from the Pkhd1siRNA transient transfection assays (Figure 1, H and I), the stably Pkhd1-silenced IMCD cell lines were used for tubulogenesis assays. Loss of tubulogenesis was consistently seen in the cultures of Pkhd1-silenced cells. Cells failed to establish tubulogenesis in 2-4% of the IMCDWT, 85-95% of the IMCDshRNA3e11, 75-85% of the IMCDshRNA3e23, and <10% of the IMCDshRNA3M5 and IMCDshRNA3M9 of the CI (Figure 3A) and MG (Figure 3B) 3-D cultures. The significant inhibition of tubulogenesis observed in the Pkhd1-silenced cell lines, together with the results from our transient Pkhd1-dsRNA-mediated assays, strongly suggest that reduction of FPC expression prevents tubule formation and arrests normal branching morphogenesis in 3-D cultures.
Lack of Pkhd1 Impairs Cell-Cell Contacts
The establishment of intercellular junctions and normal cytoskeletal assembly is essential for epithelial polarity and tubule formation (Higashiyama et al., 1995; Matter and Balda, 2003; Zegers et al., 2003). We therefore assessed the effects of Pkhd1-silencing on these cell biological processes.
The subcellular localization and distribution of E-cadherin and ZO-1 were compared between wild-type/control and Pkhd1-silenced IMCD cells utilizing immunofluorescence staining. In the wild-type and empty-vector control cell lines, E-cadherin was predominantly seen at the cell-cell junctions (Figure 4A, a and b), whereas in the Pkhd1-silenced cells, the junctional staining was markedly indistinguishable and more cytosolic in their subcellular distribution (Figure 4A, d and e). In the mutagenized IMCDshRNA3M clones, the E-cadherin was also predominantly concentrated at the cell-cell junctions, but some of the cell-cell junctions exhibited a slightly diffused staining pattern (Figure 4Ac). These data from the serial cell lines provide evidence that silencing Pkhd1 alters the distribution of E-cadherin and impairs the formation of adherent junctions.
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). ZO-1 was predominantly found at cell-cell junctions in wild-type cells (Figure 4Ba). In the Pkhd1-silenced IMCD cells, junctional staining of ZO-1 showed a diffuse submembranous distribution pattern (Figure 4Bd). As with E-cadherin, ZO-1 staining in the mutagenized IMCDshRNA3M clones was different from the wild-type and empty-vector control cell lines (Figure 4Bc). These results indicate that lack of FPC may also affect the structure of tight junctions. Given the fact that IF staining showed different distribution patterns for E-cadherin and ZO-1 in wild-type/control versus Pkhd1-silenced IMCD cells, we performed a Western blot analysis to determine whether there was a variation in the E-cadherin or ZO-1 expression levels among the different cell lines. There was no detectable immunoreactive change of both proteins between cells with and without down-regulation of FPC (unpublished data), suggesting that FPC alters the cellular distribution of these proteins rather than their levels of expression.
To characterize the functional consequences of these alterations in cell-cell interactions, we measured cell TER. In contrast to the wild-type and control cell lines, TER was low in Pkhd1-silenced cells after 3 d of transwell culture (p < 0.01) and this persisted out to day 6 (Figure 4C). These results suggest that Pkhd1-silenced cells lose the ability to establish normal cell-cell contacts.
Because the cell-cell interactions in the cultured Pkhd1-silenced IMCD cells appeared to be abnormal, we hypothesized that down-regulation of FPC might alter the cytoskeleton of these cells. We therefore performed rhodamine-phalloidin staining on the cells. As predicted, we found that the inhibition of FPC disturbed the normal cortical distribution of actin cytoskeleton, resulting in lamellipodia formation (Figure 4D, a and b vs. c). These findings indicate that lack of FPC is able to alter the cytoskeletal organization and induce epithelial to mesenchymal transformation (EMT).
Down-regulation of Pkhd1 Induces Aberrant Cell Motility
Based on the morphological changes seen in the Pkhd1-silenced IMCD cells with respect to cell-cell interactions and the actin cytoskeleton, we examined the motility of the cells in a scratch wound assay on tissue culture plastic in the presence of 0.5% serum. Wound healing in the Pkhd1-silenced IMCD cells (IMCDshRNA3e11 and IMCDshRNA3e23) was only slightly faster than control cells (IMCDsh and IMCDshRNAM5) and only the IMCDshRNA3e11 clone was statistically significant (p < 0.05; Figure 5B). It was noted that Pkhd1-silenced cells in margins of the wound were markedly lamellipodial in appearance and did not grow in tight apposition to each other as seen in wild-type IMCD cell cultures (Figure 5A, e and f vs. g and h). To further determine the cell motility of these cell lines, time-lapse microscopy was performed to trace the movement of single cells on tissue culture plastic for 3 h. In this assay Pkhd1-silenced IMCD cells showed a faster migration rate than that of control cell lines (Figure 5C).
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To demonstrate if the inhibition of FPC induced aberrant migratory polarity in the Pkhd1-silenced cells, time-lapse microscopy was performed on cell colonies that were grown on 0.2% gelatin-coated culture plates and photographed at 20-min intervals for >20 h. Intriguingly, the Pkhd1-silenced IMCD cells exhibited spontaneous scattering of the cell colonies (Figure 5D, a-c, Supplementary Video S1), developed a fibroblast-like appearance, and migrated away from each other. In contrast, the wild-type and mutagenized control cells exhibited clustered-cell growth and collective cell migration (Figure 5D, d-i and Supplementary Videos S2 and S3). Taken together, these data suggest that the Pkhd1-silenced IMCD cells become less epithelial-like than IMCD controls and have undergone a mesenchymal transformation.
Inhibition of the Pkhd1 Gene Product Decreases Integrin-dependent Cell Adhesion
It is well known that cell-ECM interactions play a critical role in branching morphogenesis and perturbations of these interactions in IMCD cells result in decreased branching morphogenesis (Balda and Matter, 2003; Chen et al., 2004). For this reason we investigated the effects of silencing Pkhd1 in IMCD cells on integrin-dependent adhesion to CI. Cells in which Pkhd1 is silenced adhere less well than control cells at concentrations of CI from 0.1 to 2 µg/ml (Figure 6A). At 2 µg/ml, the IMCDshRNA3e11 and IMCDshRNA3e23 cells showed only 20-40% cell adhesion compared with >70% for all control cell lines (p < 0.001). To identify the importance of FPC on integrin-dependent cell migration, we performed transwell migration assays on CI. As shown in Figure 6B, the Pkhd1-silenced cells migrated significantly less than the control groups (p < 0.05). These results indicated that inhibition of FPC disrupts integrin-mediated cell interactions.
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Reduction of FPC Inhibits Cell Proliferation and Induces Apoptosis in Cultured IMCD Cells
During the time-lapse microscopy examination we observed that the Pkhd1-silenced cells proliferated significantly less than the control cells (Supplementary Videos S1-S3). To further validate this finding, we performed proliferation assays in the 3-D CI gels. Knocking down Pkhd1 resulted in a significant decrease in tritiated thymidine uptake (p < 0.01), indicating that the elimination of FPC inhibits cell proliferation (Figure 7A). Similar results were obtained with another proliferation assay, in which the activity of cellular mitochondrial dehydrogenases is measured based on cleavage of the tetrazolium salt WST-1 to formazan (Berridge et al., 1996; Figure 7B). Taken together these assays indicate that lack of FPC suppresses cell proliferation.
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) and polycystic kidney disease (Woo, 1995, Boletta, 2000), apoptosis of cells with and without Pkhd1-silencing were assessed and quantified by TUNEL assays in the 3-D CI gels. Only 5-10% of the wild-type and control cells underwent programmed cell death, whereas >25% of the Pkhd1-silenced IMCD cells underwent apoptosis (p < 0.001; Figure 7C). To further confirm the results from the TUNEL assay, we used Annexin V as an indictor to detect programmed cell death in all the tested cell lines (Figure 7D). In agreement with the results from the TUNEL assay, the apoptosis rate seen in the Pkhd1-silenced IMCD cell lines was significantly higher than wild-type and all the control cells (p < 0.001). These results suggest that inhibition of FPC, either directly or indirectly, promotes programmed cell death during tubulomorphogenesis in vitro.
Stable Pkhd1-silenced IMCD Cells Exhibit Aberrant Ciliogenesis
FPC has recently been demonstrated to localize to the primary cilium and/or basal bodies of renal tubular epithelia (Masyuk et al., 2003; Ward et al., 2003; Menezes et al., 2004; Wang et al., 2004; Zhang et al., 2004) and malformation of this cell organelle has been shown to induce cyst formation in the kidneys (Yoder et al., 2002; Lin et al., 2003). We therefore determined whether a lack of FPC arrests ciliogenesis in cultured renal epithelial cells. To this end, scanning electron microscopy (SEM) was used to examine the primary cilia in cells with or without Pkhd1-silencing. In contrast to wild-type/control cells (Figure 8A), ciliary formation is markedly suppressed in Pkhd1-silenced cells (Figure 8B). Additionally, when a common ciliary marker, acetylated anti--tubulin antibody, was used to determine the existence of ciliary malformation in cultured Pkhd1-silenced cells, 92% of wild-type IMCD cells showed ciliary staining, whereas Pkhd1-silenced IMCD cells displayed <10% staining (Figure 8, C and D). Compared with control cell lines IMCDshRNA3M5 and IMCDsh21 (Figure 8, E and F), Pkhd1-silenced IMCD cells also showed shorten ciliary structure and decreased ciliary staining (Figure 8G). The fact that severe ciliary shortness and absence were found in Pkhd1-silenced IMCD cells but not in wild-type/control cells (Figure 8H) suggests that lack of FPC may cause defects in ciliogenesis of renal epithelial cells in vitro.
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). To determine which putative signaling regulators are most likely to be involved in the processes of FPC-induced tubulomorphogenesis, we examined FAK, JNK/SAPK, PKB/Akt, and ERK1/2 phosphorylation activities, which could be responsible for aberrant cell migration, proliferation, and apoptosis in Pkhd1-silenced IMCD cells. Significant differences between wild-type/control and Pkhd1-silenced IMCD cells were only seen in ERK and FAK activation. ERK was strongly phosphorylated 30 min after adhesion to CI in the control cells (Figure 9A); however, ERK phosphorylation only occurred at 60 min in the Pkhd1-silenced IMCD cells. The phosphorylation in the Pkhd1-silenced IMCD cells never reached the levels of the wild-type/control cells.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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; Ward et al., 2002). To date, no studies pertaining to the functional roles of this large and complex protein have been reported. In this in vitro study, utilizing stable Pkhd1-silenced IMCD cells, we demonstrated that down-regulating Pkhd1 results in 1) the inhibition of tubulomorphogenesis; 2) impairment of cell-cell interactions and inducement of spontaneous cell scattering; 3) disorganization of the actin cytoskeleton and induction of EMT; 4) reduction of integrin-dependent adhesion; v) increased apoptosis and decreased cell proliferation; and 5) decreased FAKpY861 and ERK1/2 activation. Because FPC localizes at vicinity of the basobodies (centriole system) and the primary cilia (microtubule system) of epithelial cells (Masyuk et al., 2003; Ward et al., 2003; Menezes et al., 2004; Wang et al., 2004; Zhang et al., 2004), dysfunction of FPC may potentially disrupt the core pathway for cytoskeleton distribution/assembly and disable scaffold protein transport for cell polarity generation, which might account for the diversity of cell biological alterations observed in our in vitro study.
A prominent biological defect found in the Pkhd1-silenced IMCD cells was the disruption of normal cell-cell interactions. The loss of E-cadherin-mediated cell-cell adhesion blocks the assembly of adherens junctions which is required for all other intercellular junction formation in cultured MDCK cells (Matter and Balda, 2003). Nevertheless, normal assembly of these intercellular junctions is essential for tubulogenesis (Grisendi et al., 1998; Balda and Matter, 2003). In addition, defects in cell-cell adhesion were able to promote spontaneous cell scattering phenotypes and induce a failure in collective migration or migratory polarity of the renal epithelial cells, which is critical for normal tubule formation (Zegers et al., 2003).
The failure of renal epithelia to assemble primary cilia induces cystogenesis in the kidneys (Yoder et al., 2002; Lin et al., 2003). Our in vitro findings suggested that ciliary generation was disrupted in Pkhd1-silenced IMCD cells, implicating possible FPC involvement in ciliogenesis. This result is consistent with a report that transiently siRNA-mediated inhibition of Pkhd1 in chlolangiocytes resulted in shortening and decreased formation of cilia (Masyuk et al., 2003). However, a mouse model with an exon 40 deletion of Pkhd1, which caused distinct liver cysts, did not show any ciliary abnormality in chlolangiocytes, suggesting that a lack of functional PFC may not affect normal ciliogenesis in vivo (Moser et al., 2005). In spite of the spatial and environmental differences between the in vivo tissues and in vitro culture systems, the ciliary defect seen in cultured Pkhd1-silenced IMCD cells may be caused by arrestment of epithelial polarity, which may occur because of disruption of cytoskeleton organization, cell-cell/matrix contacts, and centriole arrangement.
The aberrant cell behaviors including abnormal proliferation, cell scattering, and disrupted cell-cell/matrix contacts can be mediated through several signal-transduction processes (Matter and Balda, 2003; Wozniak et al., 2004). The down-regulation of JNK/SAPK phosphorylation induces cell scattering and the activation of ERK and PKB/Akt regulates epithelial tubulogenesis in cultured cells (Paumelle et al., 2000; O'Brien et al., 2004; Lavenburg, 2003). In addition, focal adhesive kinase (FAK) has been indicated to play a role in reorganization of the cytoskeleton, assembly of cell adhesion structures, and regulation of cell membrane protrusions to cell migration (Schaller, 2004). In our studies, we found a significant difference in the ERK signaling pathway, which has been implicated as a key regulator for cell proliferation (Roux and Blenis, 2004) and epithelial tubule formation in vitro (O'Brien et al., 2004). Thus the aberrant proliferation and impairment of tubulomorphogenesis found in cultured Pkhd1-silenced IMCD cells may be mediated by disturbances of ERK signaling.
The observation that a difference in FAK phosphorylation was present at pY861 was intriguing. FAK is a protein tyrosine kinase that contains multiple critical tyrosine phosphorylation sites that could contribute to its central role in cell-cell/matrix contacts, cytoskeleton remodeling, cell scattering, and polarization (Mitra et al., 2005). FAK is required for normal mouse developmental morphogenesis (Ilic et al., 1995). Furthermore lack of FAK results in abnormalities in endothelial tubulogenesis in a 3-D cultured system (Ilic et al., 2003). The fact that FAKpY861 was less phosphorylated may unveil a novel pathway that links FAK/ERK signaling to tubulomorphogenesis (Juliano, 2002; O'Brien et al., 2004; Mitra and Hanson, 2005).
In addition, our results have also showed that normal cell-cell/matrix contacts in cultured Pkhd1-silenced IMCD cells have been impaired. It is feasible that loss of normal cell-cell/matrix contacts could disrupt junction-related gene expression (Balda and Matter, 2003). This disruption may dysregulate many cytoplasmic proteins, including transcriptional factors and/or other proteins that control cell-cycle progression, cytoskeleton remodeling, and epithelial polarization either directly or through other intermediary molecules, and eventually impair normal cell behaviors, including the multiple abnormalities observed in this study.
In our studies, we adopted a shRNA inhibition approach to knock down Pkhd1 expression. We utilized an empty-vector control, IMCDsh and a mutagenized control, IMCDshRNA3M, in which two base pairs in the middle of the hairpin duplexes were mutagenized to emphasize the specificity of our knockdown construct (Figure 2A). The use of the empty-vector IMCDsh cells was to prevent Pkhd1-specific micro-RNA effects, which may be produced by the IMCDshRNA3M construct (Ambros, 2004). This indeed appeared to be the case in our studies as the mutagenized IMCDshRNA3M cells exhibited a slightly decreased Pkhd1 expression (Figure 2, F and M), resulting in minor alterations of E-cadherin and ZO-1 staining patterns (Figure 4, Ac and Bc). However, these graded changes from Pkhd1shRNA3 to Pkhd1shRNA3M and finally to IMCDsh provided further evidence for the silencing specificity of our Pkhd1shRNA3 construct.
In summary, we have demonstrated that down-regulation of FPC disrupts normal cell-cell and cell-ECM interactions of epithelial cells and promotes spontaneous cell scattering. In addition, it results in alterations in the cell proliferation, apoptosis, polarity, and cytoskeleton organization, which may be mediated by FAK and ERK signaling. As a consequence of some or all of these changes in cell biological processes, Pkhd1-silenced IMCD cells results in inhibition of tubulomorphogenesis in vitro. Given that FPC is expressed in the organs with primary duct systems, including the kidney, lung/trachea, mammary gland, cardiovascular system, and gastrointestinal/urinary-genital tracts, and is predominantly distributed at the apical domain of epithelial cells (Nagasawa et al., 2002; Zhang et al., 2004), this protein may serve as a key molecule for creating and/or maintaining the lumen of tubules/ducts. Disruption of this functional molecule gives rise to a pathogenic path for cystogenesis of ARPKD, in which massive fusiform ectasia of renal tubules is seen.
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ACKNOWLEDGMENTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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Footnotes |
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The online version of this article contains supplemental material at MBC Online (http://www.molbiolcell.org). 
Address correspondence to: Guanqing Wu (guanqing.wu{at}vanderbilt.edu).
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