Cystogenesis in ARPKD results from increased apoptosis in collecting duct epithelial cells of Pkhd1 mutant kidneys

E XP E RI ME N T AL C ELL R ES E AR C H 3 1 7 ( 2 0 11 ) 1 7 3– 1 87

available at www.sciencedirect.com

www.elsevier.com/locate/yexcr

Research Article

Cystogenesis in ARPKD results from increased apoptosis in collecting duct epithelial cells of Pkhd1 mutant kidneys Bo Hua,b , Xiusheng Hea , Ao Lid , Qingchao Qiua,b , Cunxi Lib , Dan Liang b , Ping Zhaod , Jie Mad , Robert J. Coffeyb,c , Qimin Zhand , Guanqing Wub,c,d,⁎ a

Cancer Research Institute, University of South China, Hengyang, Hunan, 421001, China Department of Medicine, Vanderbilt University, Nashville, TN 37232, USA c Department of Cell and Developmental Biology, Vanderbilt University, Nashville, TN 37232, USA d Division of Translational Cancer Research and Therapy, State Key Laboratory of Molecular Oncology, Cancer Hospital and Institute, Chinese Academy of Medical Sciences, Beijing, 100021, China b

A R T I C L E I N F O R M A T I O N

A B S T R A C T

Article Chronology:

Mutations in the PKHD1 gene result in autosomal recessive polycystic kidney disease (ARPKD) in

Received 28 July 2010

humans. To determine the molecular mechanism of the cystogenesis in ARPKD, we recently

Revised version received

generated a mouse model for ARPKD that carries a targeted mutation in the mouse orthologue of

9 September 2010

human PKHD1. The homozygous mutant mice display hepatorenal cysts whose phenotypes are

Accepted 19 September 2010

similar to those of human ARPKD patients. By littermates of this mouse, we developed two

Available online 25 September 2010

immortalized renal collecting duct cell lines with Pkhd1 and two without. Under nonpermissive culture conditions, the Pkhd1−/− renal cells displayed aberrant cell–cell contacts and

Keywords:

tubulomorphogenesis. The Pkhd1−/− cells also showed significantly reduced cell proliferation

Pkhd1

and elevated apoptosis. To validate this finding in vivo, we examined proliferation and apoptosis in

Fibrocystin

the kidneys of Pkhd1−/− mice and their wildtype littermates. Using proliferation (PCNA and

ARPKD

Histone-3) and apoptosis (TUNEL and caspase-3) markers, similar results were obtained in the

Apoptosis

Pkhd1−/− kidney tissues as in the cells. To identify the molecular basis of these findings, we

Proliferation

analyzed the effect of Pkhd1 loss on multiple putative signaling regulators. We demonstrated that the loss of Pkhd1 disrupts multiple major phosphorylations of focal adhesion kinase (FAK), and these disruptions either inhibit the Ras/C-Raf pathways to suppress MEK/ERK activity and ultimately reduce cell proliferation, or suppress PDK1/AKT to upregulate Bax/caspase-9/caspase-3 and promote apoptosis. Our findings indicate that apoptosis may be a major player in the cyst formation in ARPKD, which may lead to new therapeutic strategies for human ARPKD. © 2010 Elsevier Inc. All rights reserved.

Introduction Autosomal recessive polycystic kidney disease (ARPKD) is one of the most common hereditary renal cystic diseases in infants and

children, with an estimated incidence of ~ 1 in 20,000 live births and a prevalence ~ 1 in 70 for heterozygosity [1,2]. The disease is caused by mutations in PKHD1, which encodes a 16-kb transcript, contains at least 86 exons, and spans 470 kb on chromosome 6p12

⁎ Corresponding author. Division of Genetic Medicine, Department of Medicine and Cell & Developmental Biology, Vanderbilt University, 539 LH, 2215 Garland Ave., Nashville, TN 37232, USA. Fax: + 1 615 936 2661. E-mail address: [email protected] (G. Wu). 0014-4827/$ – see front matter © 2010 Elsevier Inc. All rights reserved. doi:10.1016/j.yexcr.2010.09.012

174

E XP ER I ME NT AL CE L L R ES E AR CH 3 1 7 (2 0 11 ) 1 73 – 1 87

[3]. The longest ORF is predicted to have 66 exons and to yield a 4074-amino acid membrane-associated receptor-like protein, fibrocystin/polyductin (FPC) [4–7]. FPC has a predicted ~3900-amino acid extracellular region that sequentially contains an NH-terminal signal peptide and a minimum of 7 or 8 IPT/TIG and 5 TIG-like domains clustered in the first half of the protein, and 9 or 10 PbH1 repeats [8], which assemble into two transmembrane protein (TMEM) homology domains [9], in the COOH-terminal half of the protein [7,10]. In the membrane-proximal region, FPC's extracellular domain bears a proprotein convertase site, which may be cleaved to release the extracellular domain into the tubular lumen through the apical domain/primary cilium of renal epithelial cells [11,12]. Only one putative transmembrane domain is predicted in the COOH-terminal portion. The last 192 amino acids of FPC are predicted to reside in the intracellular cytoplasm [5,6], and can be proteolytically cleaved and transported into the nucleus of renal epithelial cells to regulate cell behavior [13]. Recently, a ciliary targeting sequence (CTS) was also identified in the intracellular portion of FPC. It is believed to associate with the regulatory protein Rab8 and to mediate FPC trafficking through the ER to the primary cilium of epithelial cells [14]. To study the functional roles of the PKHD1 gene product (FPC), various animal mutant models have been generated by gene targeting, and all of the mouse models exhibit cystogenesis in the liver/pancreas and/or kidneys [15–19]. By studying the pck rat, which carries a mutation in the rat orthologue of human PKHD1, Fischer et al. demonstrated that the disruption of FPC function causes aberrant planar cell polarity (PCP) in the epithelial cells of renal tubule/ducts, which could be responsible for inducing the renal cysts [20]. This cellular defect may be induced by disorganized spindle orientation in the FPC-deficient epithelial cells [21]. In addition, several reports demonstrated that disruption of the transcriptional activator hepatocyte nuclear factor-1β (HNF-1β) can induce cyst formation in the kidney [22]. The loss of HNF-1β can developmentally downregulate the transcription of Pkhd1 and other PKD-causal genes, such as Pkd2 and Bicc1, suggesting the HNF-1β-dependent PKD phenotypes may be due to the inhibition of these cystoproteins [23,24]. Intriguingly, a genetic link between FPC and polycystin-1 or -2 (PC1 or PC2), the products of the PKD1 and PKD2 genes, whose mutations cause autosomal dominant polycystic kidney disease (ADPKD), was also demonstrated [16,18,25]. Other studies indicated that FPC can interact with PC2 (also known as TRPP2) to modulate the PC2-associated cation channel [26,27] and that the disruption of FPC impairs the channel function [18,28]. Our previous study indicated that the downregulation of Pkhd1 promotes apoptosis and reduces proliferation in a renal collecting duct cell line [29]. However, the observation was controversial, and the molecular mechanisms underlying the decreased proliferation and increased apoptosis have remained unknown [28,30– 32]. In the current study, to clarify these issues, we developed a panel of immortalized renal collecting duct cell lines with and without Pkhd1. The null-Pkhd1 cell lines displayed significantly decreased proliferation and increased apoptosis, compared to littermate-derived wildtype cell lines. Consistent with our in vitro finding, the kidneys of the Pkhd1−/− mice also exhibited very low cell proliferation and significantly upregulated apoptosis in vivo. Using the cell lines, we found that the loss of FPC downregulates Ras/C-Raf signaling and reduces MEK/ERK activity, leading to decreased proliferation in renal epithelial cells. We also found that

the absence of FPC significantly promotes apoptosis by downregulation of the FAK/PI3K/AKT/Bax pathway, suggesting that apoptosis may be a major contributor to cyst formation in ARPKD. The novel contribution of these signaling pathways to the aberrant proliferation and apoptosis was mediated by the dysfunction of multiple major phosphorylation sites of focal adhesion kinase (FAK), whose disruption can induce many cell-processing defects. Our findings indicate that the apoptotic loss of renal epithelia and tissues may be a major player in the cystogenesis of ARPKD and that cell proliferation may not be necessary for it. This new concept for the pathogenic mechanism of ARPKD cystogenesis should lead to new therapeutic strategies for human ARPKD.

Results Establishment and characteristics of renal collecting duct cell lines bearing null-Pkhd1 alleles, from Pkhd1−/− mutant kidneys To examine the molecular mechanisms of cystogenesis in ARPKD, we recently generated a mutant mouse model for Pkhd1. The homozygous mice (Pkhd1−/−) display hepatorenal cysts, whose phenotypes are similar to those of human ARPKD patients [18]. We then mated the Pkhd1+/− mice with Immorto mice (Im) (both with C57/Bl6 congenic background) [33] to produce Im::Pkhd1−/− mice and their Im::WT (wildtype) littermates. To establish null-Pkhd1 cell lines, the kidneys from an 8-week-old Im::Pkhd1−/− mouse and its Im::WT littermate were removed, and a Dolichus biflorus agglutinin (DBA)based isolation approach was used to develop immortalized renal collecting duct cell lines of both genotypes [34]. After a limiting dilution, 38 immortalized renal collecting duct cell colonies were isolated from each of the Im::Pkhd1−/− and Im::WT cell pools. To identify the origin of the cell lines, we used E-cadherin and cytokeratin as epithelial markers and DBA as a collecting duct marker (Fig. 1A–C). By these biomarkers, 26 collecting duct cell lines with the Im::Pkhd1−/− genotype were selected from the Im::Pkhd1−/− cell pool, and 21 Im::WT collecting duct cell lines were selected from the Im::WT cell pool. Of these, two randomly selected lines from each pool (W10B6 and W10B2 for wildtype and M10H2 and M10C7 for Pkhd1−/−) were used as the genotype-representative cell lines for further analysis (Fig. 1). Quantitative PCR verified that the wildtype cell lines expressed Pkhd1 and the Pkhd1−/− cell lines did not (Fig. 1D–E). To further confirm the genotypes of these cell lines, an anti-FPC monoclonal antibody hAR-C2m4E12, which is a subclone from hAR-C2m3C10 [35], was used to detect the FPC expression levels in the cell lines by western blot. Consistent with the quantitative PCR results, the Pkhd1−/− cell lines did not express any detectable FPC, while the wildtype cell lines expressed it (Fig. 1F). These results indicated that the Pkhd1−/− cell lines were Pkhd1-deficient cells. We next characterized the cell lines, by first performing experiments in 3-D Matrigel culture to examine whether the loss of FPC induced abnormal tubulomorphogenesis in vitro. Most of the wildtype cells formed normal tubular structures in the 3-D culture, and only 5–10% of them failed to exhibit tubulogenesis. In sharp contrast, ~95% of the 3-D cultured Pkhd1−/− cells failed to undergo tubulogenesis (Suppl. Fig. 1A–D). In addition, less than 5% of the colonies in the Pkhd1−/− cell cultures had 3 or more branches,

E XP E RI ME N T AL C ELL R ES E AR C H 3 1 7 ( 2 0 11 ) 1 7 3– 1 87

175

Fig. 1 – Characteristics of collecting duct cell lines derived from the kidneys of an 8-week-old Pkhd1−/− mouse and its wildtype littermate. The cell lines with Pkhd1 (W10B6 and W10B2) and without it (M10H2 and M10C7) were characterized by staining with epithelial cell markers, cytokeratin (A) and E-cadherin (B); and a collecting duct cell marker, Dolichos biflorus agglutinin (DBA) (C). Positive staining for all three markers was seen in the tested cell lines. DAPI, a dye for nucleic acids (blue) is used in C. (D) RT-PCR was used to determine the Pkhd1 mRNA levels in the null-Pkhd1 and wildtype littermate cells. No Pkhd1 mRNA was detected in the M10H2 and M10C7 cells. (E) Quantitative PCR yielded similar results as the RT-PCR assays. The wildtype cell line W10B6 was used for the 100% control. (F) Western blot analysis using an anti-FPC monoclonal antibody hAR-C2m4E12 agreed with the PCR results; the W10B6 and W10B2 cell lines showed positive immunoreactivity for FPC, while no band was detected in the M10H2 and M10C7 cell lines. Bar = 5 μm in A–C.

whereas ~60% of the wildtype tubular structures did (Suppl. Fig. 1D). Thus, the loss of FPC interrupts normal tubulomorphogenesis in vitro. We also analyzed the cell–cell contacts in these lines. The Pkhd1−/− and wildtype-littermate cells were stained with antibodies against E-cadherin and ZO-1, which are putative cell–cell junctional markers

[36]. Although E-cadherin and ZO-1 were predominantly found at the cell–cell junctions of the wildtype cells, the Pkhd1−/− cells showed discontinuous and diffuse sub-membranous distribution patterns for these proteins (Suppl. Fig. 1E). However, western blots did not show a detectable difference in the expression of E-cadherin

176

E XP ER I ME NT AL CE L L R ES E AR CH 3 1 7 (2 0 11 ) 1 73 – 1 87

and ZO-1 between the Pkhd1−/− and wildtype cells (Suppl. Fig. 1F). To confirm the altered cell–cell interactions, we measured the cell transepithelial resistance (TER) to determine if the integrity of the cell–cell contacts was impaired. The TER was significantly lower in

the null-Pkhd1 cells than in the wildtype ones after 3 days of transwell culture (*P < 0.05), and this difference persisted to day 7 (Suppl. Fig. 1G). Collectively, these findings suggested that the loss of FPC interrupts normal cell–cell contacts.

E XP E RI ME N T AL C ELL R ES E AR C H 3 1 7 ( 2 0 11 ) 1 7 3– 1 87

Loss of FPC promotes renal epithelial apoptosis but not proliferation in vitro and in vivo Renal cyst formation is closely associated with the proliferation and apoptosis of tubular epithelial cells, but it is usually considered to be induced by increased renal epithelial proliferation [37–41]. Although our previous in vitro study with Pkhd1-silenced inner medullary collecting duct (IMCD) cells indicated that a lack of FPC downregulates renal epithelial proliferation [29], this finding remained controversial. We therefore subjected the renal epithelial cell lines from our Pkhd1 knockout mouse model to a tritiated thymidine proliferation assay, to determine whether the loss of FPC caused a decrease in renal epithelial proliferation. Compared to the W10B6 and W10B2 wildtype-littermate cells, the M10H2 and M10C7 (Pkhd1−/−) cells showed a significant decrease in tritiated thymidine uptake (*P < 0.05), suggesting that the loss of Pkhd1 inhibits cell proliferation (Fig. 2A). Similar results were obtained using a Phospho-Histone H3 staining assay (*P < 0.05) (Fig. 2B), which reflects cell mitotic and meiotic activity. We also performed western analyses using lysates from the Pkhd1−/− and wildtype cell lines with an anti-Phospho-Histone H3 antibody. We found that the loss of Pkhd1 significantly reduced Phospho-Histone H3 expression (Fig. 2C upper panel and D left panel). To confirm these results, we examined the proliferating cell nuclear antigen (PCNA) protein, another putative proliferation marker, to evaluate the proliferation rate in the same cells. Compared to the wildtype cell lines, western analyses with an anti-PCNA antibody showed significant PCNA downregulation in lysates from the null-Pkhd1 cells (Fig. 2C lower panel and D

177

right panel). All the results of these assays support that the loss of FPC suppresses renal collecting duct cell proliferation in vitro. Since increased proliferation does not occur in ARPKD epithelial cells, we reasoned that apoptosis might serve as a major player for the disease-related cystogenesis. We examined the apoptosis rates for the Pkhd1−/− and wildtype cell lines using TUNEL (Terminal deoxynucleotidyl Transferase Biotin-dUTP Nick End Labeling) assays. Under routine culture conditions, approximately 5% of the wildtype cells and around 14% of the null-Pkhd1 cells were apoptotic (*P<0.05) (Fig. 2E). To confirm this result, we assayed active caspase-3 as an indictor of apoptosis in the same cell lines. In agreement with the TUNEL assay results, the percentage of apoptotic Pkhd1−/− cells was significantly higher than that of the wildtype cells (*P<0.05) (Fig. 2F). These results suggested that normal Pkhd1 expression can prevent programmed cell death of the renal epithelial cells in vitro. To examine whether this decreased proliferation and increased apoptosis also occurred in vivo, we performed immunohistochemical (IHC) staining of kidneys from a 6-week-old Pkhd1−/− and its wildtype littermate using antibodies against Phospho-Histone H3 and PCNA (for proliferation). The epithelial cells of the Pkhd1−/− kidneys clearly showed fewer positive spots of Phospho-Histone H3 staining compared to those of the wildtype-littermates (Fig. 2G). Similar results were obtained for the staining of PCNA (Fig. 2H). These in vivo data strongly supported the idea that the loss of Pkhd1 downregulates epithelial proliferation in the ARPKD mouse kidneys. In addition, we also performed immunofluorescent (IF) staining of kidneys from a set of 6-month-old mice using anti-PCNA antibody. Less positive cells were seen in the Pkhd1−/−

Fig. 2 – Loss of Pkhd1 inhibits cell proliferation and promotes apoptosis. (A) Pkhd1−/− cells M10H2 and M10C7 and their wildtype littermate cells W10B6 and W10B2 were incubated with 3 H-thymidine, then the rate of 3 H-thymidine incorporation was determined as described in Materials and methods, at least three times. The 3H-thymidine values were significantly greater in the cells with Pkhd1 than in those without (*P < 0.05). (B) The same cell lines were subjected to Phospho-Histone H3 staining to evaluate cell proliferation. The percentage of cells positive for Phospho-Histone H3 was significantly greater in the cells with Pkhd1 than in those without (*P < 0.05). (C) Western analyses with Phospho-Histone H3 and PCNA antibodies showed that both these proliferation markers were significantly downregulated in the null-Pkhd1 cells (M10H2 and M10C7), compared with their wildtype littermate cells W10B6 and W10B2. Anti-β-actin was used as a protein loading control. (D) Normalized quantitative analysis of the densitometry values of Phospho-Histone H3 and PCNA using the results from at least three western blots. The loss of Pkhd1 significantly downregulated the Phospho-Histone H3 and PCNA expression levels (*P < 0.05). (E) The same cell lines were cultured in 24-well plates, then fixed and subjected to TUNEL assays to assess apoptosis three times. The values represent the percentage of apoptotic cells relative to the total number of cells. Significantly more (>3 times) null-Pkhd1 cells M10H2 and M10C7 were apoptotic than wildtype cells W10B6 and W10B2 (*P < 0.05). (F) The same cell lines were cultured in 24-well plates with the same number of cells per well for 24 h. A PE Caspase-3 Active Apoptosis kit was used to measure the apoptosis rate, according to the manufacturer's instructions. The values represent the percentage of apoptotic cells relative to the total number of cells. Significantly more (~3 times) null-Pkhd1 cells were positive for active apoptosis (*P < 0.05). (G) IHC staining of Phospho-Histone H3 in the kidney of a 6-week-old Pkhd1−/− mouse and its wildtype littermate. Many positive-stained cells were seen in the wildtype kidneys, but only a few appeared in the corresponding region of Pkhd1−/− kidneys (arrows). (H) Similar observations were obtained using a PCNA antibody (arrows). (I) The kidney sections of a 6-month-old Pkhd1−/− mouse and its wildtype littermate were co-stained with PCNA (red) and DBA (collecting duct marker, green) or LTL (proximal tubule marker, green). Much less PCNA-positive (red) cells were seen in Pkhd1−/− kidneys than its wildtype littermate (arrows). (J) The same 6-month-old kidney sections were also co-stained with caspase-3 (red) and DBA (green) or LTL (green). Increased caspase-3-positive (red) cells were seen in the Pkhd1−/− kidneys than its wildtype littermate (arrows). (K) The merged microscopic images from immunofluorescence staining and light illumination are showed. A TUNEL kit was used for the immunofluorescence staining of kidney sections from 12-month-old wildtype (WT) and Pkhd1−/− littermates. Very few positively stained cells (arrow) were seen in the WT kidneys, while several were seen in the corresponding cortical region of the Pkhd1−/− kidneys (arrows). Interestingly, many positive spots appeared inside the tubular lumen of the Pkhd1−/− kidneys (lower arrows in left panel), suggesting that the dilated renal tubules in the Pkhd1−/− kidneys might result from apoptotic loss. Bars = 20 μm in G-H; 5 μm in I–J, 15 μm in K.

178

E XP ER I ME NT AL CE L L R ES E AR CH 3 1 7 (2 0 11 ) 1 73 – 1 87

kidney than its wildtype littermate (Fig. 2I). To examine apoptosis, the same kidneys were subjected to caspase-3 staining. More positive staining was seen in the Pkhd1−/− than in the wildtype kidneys (Fig. 2J). Since older Pkhd1−/− mice exhibit severer cystic kidney [18], the kidneys of 12-month-old Pkhd1−/− and wildtype mice were also examined by TUNEL staining. Increased positive staining was seen in the Pkhd1−/− than in the wildtype kidneys (Fig. 2K). Notably, much apoptotic debris was observed in the lumen of the Pkhd1−/− renal tubules. Our in vitro and in vivo findings suggested that apoptotic loss may be a major contributor to the cystogenesis observed in ARPKD kidneys.

Downregulation of Akt phosphorylation results in elevated apoptosis in Pkhd-null cells To determine the molecular mechanisms by which the loss of Pkhd1 induces aberrant apoptosis and proliferation, we examined the Akt signaling pathway, which can regulate cell proliferation or apoptosis [42]. Unlike ADPKD cell lines, which show upregulated Akt activity [43], we found that phosphorylated Akt (pS473) was significantly reduced in the null-Pkhd1 cells compared to the wildtype-littermate cells after collagen I (CI) induction (Fig 3A,C). Because Bax and Bcl2 are putative downstream factors of Akt, and their dysfunction can also induce abnormal apoptosis [44], we next compared their factors between the null-Pkhd1 and wildtype cells. Western analysis showed no significant difference in Bcl2 expression

between the cells with and without Pkhd1, at all time points examined after CI induction (data not shown). In contrast, Bax expression was significantly higher in the null-Pkhd1 cells compared to the wildtype cells (Fig. 3B). The basal and collagen I (CI)-inducible levels of Bax were both much lower in the null-Pkhd1 cells than in the wildtype cells (Fig 3D). Since both caspase-9 and -3 are putative downstream factors of Bax, we next examined their expression levels in the same cell lines. Western analyses showed that the caspase-9 and -3 were significantly elevated in the null-Pkhd1 cells compared to the wildtype cells (Fig. 3B, E–F). These findings indicated that the loss of Pkhd1 downregulates Akt-pS473 phosphorylation, which upregulates the Bax/caspase-9/-3 pathway to trigger apoptosis in the renal epithelial cells.

Reduced proliferation of null-Pkhd1 cells is induced by aberrant Ras/MAPK signaling To investigate the molecular mechanism by which the absence of Pkhd1 causes reduced proliferation, we used western analyses to examine the Akt-downstream factor B-Raf, which is responsible for aberrant proliferation in many PKD model systems [43]. Surprisingly, there was no significant difference in the B-Raf phosphorylation in the M10H2 and M10C7 (Pkhd1−/−) cell lines versus the wildtypelittermate W10B6 and W10B2 cell lines (Fig. 4A–B). We therefore shifted our focus to c-Raf, which is another important regulator of cell proliferation [45]. Compared to the wildtype cell lines, western results

Fig. 3 – Loss of FPC downregulates Akt phosphorylation and induces the Bax/Caspase 9 pathway. The null-Pkhd1 cells M10H2 and M10C7 and their wildtype littermate cells W10B6 and W10B2 were serum starved for 12 h, trypsinized, and left in suspension or replated on 10 μg/ml collagen I (CI) for 0, 10, 30, or 60 min. Equal amounts of cell lysate were separated by 10% SDS-PAGE and transferred to nitrocellulose. (A) The membranes were immunoblotted with antibodies to phospho-Akt (pAkt) and total-Akt (tAkt). (B) Western blots using the same cell lysates were performed with anti-Bax, -Caspase 9 (Casp-9), and -Caspase 3 (Casp-3) antibodies. Anti-β-actin was used as a protein loading control. (C) A normalized quantitative analysis at the indicated times was performed using the densitometry values from the western blots for pAkt/tAkt (A). Normalized quantitative analyses were also performed for the western blots for Bax/β-actin (D), Casp-9/β-actin (E) and Casp-3/β-actin (F).

E XP E RI ME N T AL C ELL R ES E AR C H 3 1 7 ( 2 0 11 ) 1 7 3– 1 87

showed that the null-Pkhd1 cells exhibited a significant downregulation of phosphorylated c-Raf, in both the basal and induced conditions, indicating that the loss of Pkhd1 reduces renal epithelial proliferation via the dysregulation of c-Raf phosphorylation, rather than that of B-Raf, in contrast to many other PKD cells (Fig. 4A, C). Given that Raf is the first effector that is positioned downstream of Ras [46], we next investigated if the aberrant c-Raf phosphorylation was caused by Ras dysregulation. Western analyses of lysates from the Pkhd1−/− and wildtype cell lines showed that the level of Ras was significantly downregulated in the basal condition and after 30 min of CI induction in the null-Pkhd1 cells (Fig. 4A, D), indicating that the alteration in c-Raf phosphorylation may be due to a decrease in Ras. To confirm that the downregulation of Ras/c-Raf induces the decreased proliferation of the renal epithelial cells, we examined the Ras/c-Raf downstream factors MEK and ERK, both of which are key markers for cell proliferation [47], using lysates from the Pkhd1−/− and wildtype cell lines. The results showed that the phosphorylated MEK and ERK were significantly decreased in the null-Pkhd1 cells at all time points after CI induction (Fig. 4A, E–F), suggesting that the inhibition of MEK and ERK phosphorylation caused by the loss of Pkhd1 is the reason for the reduced proliferation. These findings indicate that downregulation of the Ras/c-Raf/MEK/ERK cascade may be the molecular mechanism underlying the decreased renal epithelial proliferation in ARPKD.

Aberrant apoptosis and proliferation in null-Pkhd1 renal epithelial cells may be correlated with abnormal FAK phosphorylation Focal adhesion kinase (FAK) contributes to many cellular functions in epithelial cells, including cell growth, apoptosis, adhesion, and migration, through its multiple phosphorylation sites (FAKpY397, 576–577, 861, and 925) [48]. In our Pkhd1-silenced IMCD cells, in which Pkhd1 is still expressed at one fourth of the normal level, we found that only FAKpY861 was significantly reduced [29]. It is challenged that the multiple cellular defects seen in Pkhd1silenced IMCD cells [29] are caused by inactivation of the single phosphorylation site FAKpY861 [48]. We therefore examined whether the Pkhd1 depletion could result in the dysfunction of all the major FAK phosphorylation sites, which regulate multiple cellular processes from cell migration and polarity to proliferation and apoptosis [49]. To this end, we assessed the FAK phosphorylations and their related cellular behaviors (such as integrindependent cell adhesion and migration) using our null-Pkhd1 and wildtype littermate cell lines. Besides FAKpY861, which we reported previously, the phosphorylations of FAKpY397, 576, and 925 were also significantly decreased in the M10H2 and M10C7 (Pkhd1−/−) cells compared to the wildtype-littermate W10B6 and W10B2 lines after CI induction (Fig. 5A–E). These findings indicated that the loss of Pkhd1 wipes out the major FAK phosphorylation-dependent activities, to disrupt multiple normal cellular processes. To validate the idea that the loss of Pkhd1 results in severe FAK dysfunction, we investigated the effect of null-Pkhd1 on the integrin-dependent adhesion to CI. The Pkhd1−/− cells adhered less well than the wildtype cells at concentrations of CI from 0.125 to 2 μg/ml (Fig. 5F). At 2 μg/ml CI, the null-Pkhd1 cells showed only 40% cell adhesion compared to over 90% for the control cell lines (P < 0.05). Since FAK defects can inhibit integrin-dependent cell migration, we also tested the transwell migration capability upon

179

CI between the cells with and without Pkhd1. As shown in Fig. 5G, significantly fewer null-Pkhd1 cells than wildtype cells migrated through the filter (P < 0.05). These results supported the idea that the dysfunction of FAK phosphorylation-induced activities may be responsible for the multiple aberrant cellular behaviors of the nullPkhd1 renal epithelial cells.

FAK-induced aberrant apoptosis may be mediated by the PI3K/PDK1 cascade To determine if elevated apoptosis also results from the downregulation of FAK phosphorylations, we examined several mediators that could potentially link FAK to the aberrant Akt/Bax/caspase-9 apoptotic pathway (Fig. 3). We hypothesized that the downregulation of FAK phosphorylations would disrupt PI3K activity and further dysregulate its downstream factor, PDK1. The inhibition of PDK1 phosphorylation would suppress Akt signaling, thereby provoking the apoptotic pathway [42,50]. We therefore examined lysates from our null-Pkhd1 and wildtype cell lines for changes in the phosphorylation of PI3K and PDK1. We first analyzed the PI3K phosphorylation using anti-PI3K class III and PI3Kp110α antibodies. We observed no difference in PI3K class III phosphorylation between the cells with and without Pkhd1 (Fig. 6A–B). In contrast, the basal and induced levels of PI3Kp110α were much lower in the null-Pkhd1 M10H2 and M10C7 cells than in the wildtype-littermate W10B6 and W10B2 cells (Fig. 6A). Notably, in the null-Pkhd1 cells, a low level of PI3Kp110α was observed in the basal and CI-induced condition (Fig. 6C), indicating that the loss of Pkhd1 resulted in a failure to induce PI3Kp100α phosphorylation. Since PDK1 is a downstream factor of PI3K that is activated by PI3K's phosphorylation on p100α, we analyzed the phosphorylation of PDK1 between the null-Pkhd1 and wildtype cells. Similar to PI3Kp100α, both the basal and induced pPDK1 levels were significantly lower in the null-Pkhd1 cells than in the wildtype cells (Fig. 6A, D). Given that pPDK1 can positively regulate Akt phosphorylation [42], our findings suggest that a decrease in FAK phosphorylation can promote Bax and provoke apoptosis in null-Pkhd1 cells via the PI3Kp110α/PDK1 inactivation. Thus, both the aberrant cell proliferation and the apoptosis in null-Pkhd1 cells can be linked to the dysregulation of FAK phosphorylation (Fig. 7).

Discussion In general, cystogenesis of PKD is thought to be closely associated with cell proliferation [37,39–41]. In this study, using our Pkhd1 knockout mouse model and renal collecting duct cell lines derived from this model, we demonstrated that the cystogenesis of ARPKD is predominantly caused by the apoptosis, rather than the proliferation, of renal epithelial cells. This finding represents a new understanding of the pathogenesis of human ARPKD and gives rise to a novel approach for therapeutic intervention in this disease. Increased proliferation rates have been widely reported for ADPKD epithelial cells and tissues, and the molecular mechanism by which the lack of polycystins promotes renal epithelial proliferation has been intensively investigated [34,37,51]. Given that PC1 physically interacts with PC2, and PC2 is considered to be a member of the Trp superfamily (TRPP2) [52,53], it has been proposed that PC2-associated Ca++ inhibits adenylyl cyclase IV (AC-IV) activity, thus suppressing the conversion of ATP to cAMP.

180

E XP ER I ME NT AL CE L L R ES E AR CH 3 1 7 (2 0 11 ) 1 73 – 1 87

The loss of both polycystin functions reduces channel activities and increases the cAMP level through enhanced AC-IV activity [41,54]. The high cAMP level can promote Ras/B-Raf signaling and

induce MAPK/ERK, and this dysregulation in turn upregulates the cyst-cell proliferation [55]. In the current study, we found that the loss of FPC in ARPKD downregulates the Ras/c-Raf signaling

E XP E RI ME N T AL C ELL R ES E AR C H 3 1 7 ( 2 0 11 ) 1 7 3– 1 87

181

Fig. 5 – Loss of FPC disrupts the functional activities of multiple FAK phosphorylation sites. (A) Protein lysates from the cell lines were immunoblotted with antibodies to FAKpY861, pY397, pY576, and pY925, and total-FAK (tFAK). (B) A normalized quantitative analysis at the indicated times was performed using the densitometry values from the western blots for FAKpY861/tFAK. Normalized quantitative analyses were also performed for the western blots for FAKpY397/tFAK (C), FAKpY576/tFAK (D), and FAKpY925/tFAK (E). All the tested FAK phosphorylation sites exhibited much lower activity in the null-Pkhd1 cells (M10H2 and M10C7) than in the wildtype littermate cells (W10B6 and W10B2) (P < 0.05) after CI induction. (F) Adhesion assays were performed on CI-coated plates. The values shown represent the mean and SD of at least three independent experiments. Significant differences in CI-dependent cell adhesion (at a CI concentration of 0.125 μg/ml and higher) were seen between the Pkhd1−/− and wildtype cells (P < 0.05). (G) Transwell migration assay results indicating the absolute number of cells that migrated to the underside of the transwell. There was a significant decrease in the number of migrated Pkhd1−/− cells compared to wildtype cells (W10B6 and W10B2) (*P < 0.05).

pathway, but has no effect on B-Raf. The c-Raf downregulation can further inhibit the MAPK/ERK activities, ultimately reducing epithelial cell proliferation. Our finding that Raf signaling is

different between ADPKD and ARPKD provides a molecular basis for explaining how the cystogenesis of ARPKD may not be associated with cell proliferation.

Fig. 4 – Loss of FPC downregulates Ras/MAPK. (A) Protein lysates from the cell lines were immunoblotted with antibodies to phospho-B-Raf (pB-Raf) and total-B-Raf (tB-Raf); phospho-c-Raf (pc-Raf) and total-c-Raf (tc-Raf); Pan-Ras and β-actin (a protein loading control); phospho-MEK1/2 (pMEK1/2) and total-MEK1/2 (tMEK1/2); and phospho-ERK1/2 (pERK1/2) and total-ERK1/2 (tERK1/2). (B) A normalized quantitative analysis was performed using the densitometry values from the western blots for pB-Raf/ tB-Raf. Normalized quantitative analyses were also performed for the western blots for pc-Raf/tc-Raf (C), Pan-Ras/β-actin (D), pMEK1/2/tMEK1/2 (E), and pERK1/2/tERK1/2 (F).

182

E XP ER I ME NT AL CE L L R ES E AR CH 3 1 7 (2 0 11 ) 1 73 – 1 87

Fig. 6 – FPC-loss-dependent Akt downregulation may result from FAK-induced PI3K/PDK1 inhibition. (A) Western analyses of the cell lines were performed with antibodies to PI3K ClassIII, PI3Kp110α, phospho-PDK1 (pPDK1), total-PDK1 (tPDK1), and β-actin. (B–C) Normalized quantitative analysis at the indicated times using the densitometry values from the western blots for PI3K ClassIII/β-actin and PI3Kp110α/β-actin. (D) The same normalized quantitative analysis was performed for the western blots for pPDK1/tPKD1. There was no difference in PI3K ClassIII/β-actin between the cells with and without Pkhd1, but significant differences were seen in the PI3Kp110α/β-actin and pPDK1/tPDK1 analyses, indicating that the loss of FPC downregulates the pPDK1 activity by downregulating PI3Kp110α, rather than PI3K ClassIII.

Fig. 7 – Schematic illustrating the molecular signaling by which the downregulation of FAK phosphorylation activities is linked to decreased cell proliferation and increased apoptosis, to induce cystogenesis in ARPKD. Dark circles indicate upregulation and white circles indicate downregulation.

E XP E RI ME N T AL C ELL R ES E AR C H 3 1 7 ( 2 0 11 ) 1 7 3– 1 87

Another proposed cause for cyst formation in PKD is the apoptosis of renal epithelial cells. Over a decade ago, Woo provided evidence that the apoptotic loss of renal epithelial cells might contribute to cyst formation in cpk and pcy mice, which exhibit massive renal cysts [56]. Thereafter, many other studies showed that the cystogenesis of PKD is associated with apoptosis, but the apoptosis was usually thought to be regulated cooperatively with cell proliferation [31,57–59]. No study to date has provided strong evidence showing that apoptotic loss plays a major role in inducing cyst formation in human ARPKD. In the present study, using in vitro and in vivo model systems, we showed that the cystic phenotype in human ARPKD may result from apoptosis, rather than from cell proliferation, in the affected organs. This finding represents a new concept for the pathogenesis of ARPKD. To seek the molecular mechanism by which the cyst formation in ARPKD was predominantly associated with apoptotic cell death rather than cell proliferation, we examined the serial signaling regulators that are most likely to be involved in the Pkhd1-induced cell behaviors. In our previous Pkhd1-silenced IMCD cell study, we found that a single FAK phosphorylation site (pY861) was dysfunctional [29]. Since FAK is a protein tyrosine kinase that contains multiple critical tyrosine phosphorylation sites, whose activation can regulate many different cellular processes [48], it was important to determine whether the abnormality of only the single FAK pY861 function could account for multiple cellular behavior changes [60,61]. To answer this question, we examined all the major sites of FAK phosphorylation-induced activation (pY397, 576, 861, 925) in our cell lines. Interestingly, all of these sites showed markedly delayed and decreased phosphorylation in the null-Pkhd1 cells compared to the wildtype littermate cells. This finding leads to a reasonable explanation for the multiple aberrant cellular changes in null-Pkhd1 cells. That is, the low Pkhd1 expression we previously observed in the Pkhd1-silenced IMCD cells may have resulted in only FAK pY861 dysfunction, because the small amount of remaining FPC might be able to compromise FAK phosphorylation at the other FAK activation sites. By our analyses, we linked FAK signaling to the aberrant proliferation and apoptosis observed in the null-Pkhd1 cells. FAK is a 125-kDa non-receptor protein tyrosine kinase that associates with focal adhesions and is phosphorylated in an integrindependent manner and in response to v-Src-mediated transformation [49]. However, FAK phosphorylation and activation is not only induced by integrin-based triggering, but also by diverse cellular responses to many extracellular stimuli, including epidermal growth factor (EGF) and G protein-coupled receptor agonists [49]. FAK can be initially autophosphorylated at Y397 to establish a binding site for various SH2 domain-containing molecules, such as Src kinase itself and PI3K [50,62]. When Src tyrosine kinase binds to FAK, a conformational change of the molecule can be induced and Src is activated. The activation of Src subsequently phosphorylates other sites on FAK, including Y576/Y577 which can enhance the FAK kinase activity [63]; Y861, which regulates an antiapoptotic response [61]; and Y925, which mediates the activation of Ras-MAPK signaling [64]. Dysfunction of the FAK pY397, 576, 861, and 925 phosphorylation sites in null-Pkhd1 cells would affect diverse downstream signaling pathways, to induce aberrant cell adhesion, migration, apoptosis, and proliferation, as highlighted in Fig. 7. It should be noted here that we could not exclude other molecular signaling, except the FAK-induced signaling, involve aberrant cellular behaviors in ARPKD cells and tissues.

183

Through our characterization of the mouse mutant model for Pkhd1 and its null-Pkhd1 collecting duct cell lines, we demonstrated that the cystogenesis of ARPKD predominantly results from apoptotic loss in the affected tissues or organs, and does not associate with cell proliferation. We also demonstrated the molecular mechanism by which these aberrant cell behaviors could be induced. These in vitro and in vivo findings reveal a new concept for the pathogenesis of ARPKD, and may lead to new strategies for therapeutic intervention in ARPKD patients.

Materials and methods Materials Our monoclonal antibody against the human FPC COOH-terminus (hAR-C2m4E12) is a subclone of hAR-C2m3C10 [35]. The commercial reagents and antibodies used in this study were as follows: antiE-cadherin antibody, anti-SV40 and total B-Raf/total FAK (BD Transduction Laboratories); anti-Bax, anti-PCNA (Abcam), p-Akt (ser473)/total Akt, p-c-Raf(Ser338)/total c-Raf, p-B-Raf(Ser445), PI3Kp110α, PI3K Class III, p-PDK1(Ser241)/total PDK1, p-MEK1/2 (Ser217/221)/total MEK, Cleaved caspase-3, Cleaved caspase-9, PHistone H3(SER10), p-ERK/total ERK (Cell Signaling Technology); anti-pan-Ras, anti-cytokeratin 7 (Santa Cruz Biotechnology); FAKpY397, 407, 576, 861, 925 (BioSource); Rat type I collagen (CI) (Becton Dickinson); gelatin, anti-β-actin, paraformaldehyde (Sigma); Dolichos Biflorus Agglutinin lectin (DBA), Lotus Tetragonolobus Lectin (LTL), Rhodamine-phalloidin (Vector Laboratories); anti-ZO-1 antibody (Zymed Lab, Inc.); and DAPI (Invitrogen). Secondary antibodies included Cy3-conjugated rabbit anti-mouse IgG and Cy2-conjugated goat anti-rabbit IgG (Jackson Laboratories).

Mouse strains Pkhd1 mutant mice (Pkhd1−/−) were previously generated by us [18]. Pkhd1+/− mice on a C57/Bl6 congenic background were mated with the same congenic Immortomouse (Im) [33] to obtain Im: Pkhd1−/− mice and their wildtype littermates.

Renal epithelial cell lines and their culture To establish cell lines with and without Pkhd1, kidneys from an 8week-old Im:Pkhd1−/− mouse and its wildtype littermate were removed and minced finely with a scalpel. A Dolichus biflorus agglutinin (DBA)-based isolation approach was used to develop immortalized renal collecting duct cell lines from the kidneys [34]. The collecting duct cell lines with and without Pkhd1 were selected from the Im:Pkhd1−/− and its wildtype littermate cell pool. After using Ecadherin and cytokeratin as epithelial markers and DBA as the collecting duct marker to identify their origin, we randomly chose two positive cell lines for each genotype (M10H2 and M10C7 for nullPkhd1; W10B6 and W10B2 for their wildtype control), for further analyses. The cell lines were cultured using previously described conditions [34]. The 3-D tubulogenesis assay was performed as previously described [29]. Briefly, the Matrigel (MG)/collagen I (CI) gels contained 0.5 mg/ml CI, 0.5 mg/ml MG, and 10% fetal calf serum (FCS). Tubule formation was determined in five randomly chosen high-power fields.

184

E XP ER I ME NT AL CE L L R ES E AR CH 3 1 7 (2 0 11 ) 1 73 – 1 87

For the cell lines used to well establish cell–cell contacts, the cells were grown to confluence for at least 4 days on either 12-mm transwell plates or 24-well culture plates (Costar). Before performing any cellbased assays, all the established cell lines were cultured under nonpermissive conditions (37 °C without γ-INF) for at least three days to turn off SV40 transgene activities (Suppl. Fig. 2).

RNA extraction and real-time PCR Total RNA was isolated from the tested lines using Trizol reagents (Invitrogen), according to the manufacturer's instructions. The cells were harvested 3 days after attaining confluence. Quantitative PCR was performed using the iCycler iQ Real Time PCR Detection System (Bio-Rad) [65]. The relative expression level of Pkhd1 was normalized using the 2− ΔΔCT method [66]. A pair of primers was designed to bridge exons 17 and 20 of Pkhd1 cDNA. The forward primer 5'-CAA ATG CCA CAG CCC AAC AG-3', and reverse primer 5'-CAG AAT GGT TAG GGG TGG GA -3' were used for quantitative PCR with the iQ SYBR Green Supermix kit (BioRad, Richmond, CA). PCR was performed for 35 cycles, each consisting of denaturation at 94 °C for 30 s, annealing at 56 °C for 30 s, and extension at 72 °C for 1 min. GAPDH (forward primer 5'GAC CAC AGT CCA TGC CAT CAC T-3', reverse primer 5'-TCC ACC ACC CTG TTG CTG TA-3') was amplified to confirm that the samples contained an equal amount of total mRNA, and to normalize samples.

Protein extraction and Western blotting For western blot analysis, cell lysates from 3-day confluent cultures were extracted in lysis buffer (0.5% Nonidet P-40/5% sodium deoxycholate/50 μM NaCl/10 μM Tris/HCl (pH 7.5)/1% BSA) and centrifuged at 4 °C for 15 min. For collagen I induction, the cell lysates were prepared as previously described [29]. The supernatant was mixed with 2× loading buffer and boiled for 5 min, then the samples were separated using 4%–10% gradient SDS/PAGE gels (BioRad) and transferred to nitrocellulose membranes. The membranes were incubated with 5% fat-free milk at room temperature for 1 h, blotted with primary antibodies at room temperature for 2 h, washed, and then incubated with a peroxidase-conjugate secondary Ab. Immunoreactivity was detected with enhanced chemiluminescence (Amersham Pharmacia). Quantitative analyses of western blot results were obtained from the densitometry values of the immunoreactive bands for targeted proteins normalized to the total β-actin (loading control) or for the phosphorylated form of a protein versus its total protein amount.

Histology, immunofluorescence staining, and microscopy Detailed procedures for the histology, and immunohistochemistry (IHC) experiments were published previously [18,34,35]. For immunofluorescence (IF) staining, cultured cells were washed with 1× PBS twice and fixed with 4% paraformaldehyde at 4 °C for 30 min. The cells were permeabilized in 0.1% Triton-X100 on ice for 5 min. After blocking in 2% BSA solution for 1 h, the cells were incubated with primary Abs for 2 h and washed again with 1× PBS three times. The washed cells were treated with fluorescence-conjugated secondary Abs for 1 h. For microscopy, the antibody-stained images were collected under 10×, 20×, and 40× objectives of a Zeiss Axioplan 2 IE microscope with the Zeiss Axiovision 3.1 digital color camera system.

Cell proliferation and apoptosis Cells (40,000 per well) were placed in 24-well tissue culture plates. After 5 days in culture, the cells were pulsed for 24 h with 3 H-thymidine (1 μCi/well). The medium was then removed and the plates washed with PBS 3 times to remove the free 3 Hthymidine. The cells were lysed in 0.2% NaOH after being fixed with 10% trichloroacetic acid, and the lysates were measured with a β-counter. For cell-proliferation assays, cells (20,000 per well) were placed on glass coverslips in 24-well plates and cultured for 3 days. The cells were then stained with an anti-phospho-histoneH3 and anti-PCNA antibodies. The positively stained cells were counted from three randomly chosen high-power fields (20–40×). For apoptosis assays, the cells were cultured in 24-well plates and grown to sub-confluence in 7% FCS DMEM/F-12 (1:1) medium (Gibco) under 5% CO2 at 37 °C. The cells were incubated with 0.5 μM Ionomycin (Alexis Corp.) under serum-free conditions. Six or 12 h later, the TUNEL assay (DeadEnd™fluorometric TUNEL system, Promega) or Caspase-3 Active Apoptosis Kit I (BD Biosciences) was used according to the manufacturer's manual, respectively. The apoptotic cells were counted in three randomly chosen high-power fields (40×).

Cell adhesion and migration For cell adhesion assays, 96-well cell culture plates (Nunc) 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 cells plated onto tissue culture plates in the presence of 10% FCS. The plates were washed with PBS and incubated with PBS containing 1% BSA for 60 min to block nonspecific binding. Aliquots (100 μl) of single-cell suspensions (106 cells/ml) in serumfree DMEM/F12 containing 0.1% BSA were added in triplicate to the 96-well plates with 0.03–2 μg/ml CI and incubated for 60 min at 37 °C. Non-adherent cells were removed by washing the wells with PBS. The cells were then fixed with 4% formaldehyde, stained with 1% crystal violet, and 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. 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 control minus OD of background) × 100. For the transwell filter migration assay, the cells were placed on polyvinylpyrrolidone-free polycarbonate filters with 8-μm pores (Costar). Aliquots (100 μl) of cell suspension (5 × 105 cells/ml) in 7% FCS medium were added to the wells, and the cells were allowed to migrate to the underside of the transwell for 36 h. Cells on the top of the filter were removed by wiping, and the filter was then fixed in 4% formaldehyde in PBS. The migrating cells were stained with 1% crystal violet, and five randomly chosen fields from triplicate wells were counted at 20× magnification.

Transepithelial resistance Cells were plated on transwell filters (12-well, 0.4-μm pore size, Corning Costar) at 5 × 104 cells/well and allowed to attach overnight, to form a confluent monolayer in normal culture medium. The transepithelial resistance (TER) was measured 24 h after plating and

E XP E RI ME N T AL C ELL R ES E AR C H 3 1 7 ( 2 0 11 ) 1 7 3– 1 87

every 24 h thereafter using an EVOM/STX2 (World Precision Instruments) electrical resistance measurement system, for 6 days, and the results were expressed in Ω/cm2.

Statistics All assays were repeated at least 3 times in duplicate or triplicate, and the graphic data were presented as the mean ± SD unless otherwise stated. Statistical analysis was performed where appropriate using the Student's t-test or one-way analysis of variance (ANOVA) followed by Tukey's Multiple Comparison Test. Differences with P-values < 0.05 were considered statistically significant. Supplementary materials related to this article can be found online at doi:10.1016/j.yexcr.2010.09.012.

Acknowledgments This work was supported by the National Institutes of Health of the USA (DK062373 and DK71090), the National Cancer Institute Specialized Programs of Research Excellence (5P50 CA095103), NIH, USA, and the National Natural Science Foundation of China (30672483 and 30870501), Changjiang Scholarship of China, State Key Laboratory of Molecular Oncology of China, to G.W.

REFERENCES

[1] P.C. Harris, V.E. Torres, Polycystic kidney disease, Annu. Rev. Med. 60 (2009) 321–337. [2] L.M. Guay-Woodford, Autosomal recessive PKD in the early years, Nephrol. News Issues 21 (2007) 39. [3] A.M. Sharp, L.M. Messiaen, G. Page, C. Antignac, M.C. Gubler, L.F. Onuchic, S. Somlo, G.G. Germino, L.M. Guay-Woodford, Comprehensive genomic analysis of PKHD1 mutations in ARPKD cohorts, J. Med. Genet. 42 (2005) 336–349. [4] C. Bergmann, V. Frank, F. Kupper, C. Schmidt, J. Senderek, K. Zerres, Functional analysis of PKHD1 splicing in autosomal recessive polycystic kidney disease, J. Hum. Genet. 51 (2006) 788–793. [5] L.F. Onuchic, L. Furu, Y. Nagasawa, X. Hou, T. Eggermann, Z. Ren, C. Bergmann, J. Senderek, E. Esquivel, R. Zeltner, S. Rudnik-Schoneborn, M. Mrug, W. Sweeney, E.D. Avner, K. Zerres, L.M. Guay-Woodford, S. Somlo, G.G. Germino, PKHD1, the polycystic kidney and hepatic disease 1 gene, encodes a novel large protein containing multiple immunoglobulin-like plexin-transcription-factor domains and parallel beta–helix 1 repeats, Am. J. Hum. Genet. 70 (2002) 1305–1317. [6] C.J. Ward, M.C. Hogan, S. Rossetti, D. Walker, T. Sneddon, X. Wang, V. Kubly, J.M. Cunningham, R. Bacallao, M. Ishibashi, D.S. Milliner, V.E. Torres, P.C. Harris, The gene mutated in autosomal recessive polycystic kidney disease encodes a large, receptor-like protein, Nat. Genet. 30 (2002) 259–269. [7] H. Xiong, Y. Chen, Y. Yi, K. Tsuchiya, G. Moeckel, J. Cheung, D. Liang, K. Tham, X. Xu, X.Z. Chen, Y. Pei, Z.J. Zhao, G. Wu, A novel gene encoding a TIG multiple domain protein is a positional candidate for autosomal recessive polycystic kidney disease, Genomics 80 (2002) 96–104. [8] S. Heffron, G.R. Moe, V. Sieber, J. Mengaud, P. Cossart, J. Vitali, F. Jurnak, Sequence profile of the parallel beta helix in the pectate lyase superfamily, J. Struct. Biol. 122 (1998) 223–235.

185

[9] D.A. Scott, S. Drury, R.A. Sundstrom, J. Bishop, R.E. Swiderski, R. Carmi, A. Ramesh, K. Elbedour, C.R. Srikumari Srisailapathy, B.J. Keats, V.C. Sheffield, R.J. Smith, Refining the DFNB7-DFNB11 deafness locus using intragenic polymorphisms in a novel gene, TMEM2, Gene 246 (2000) 265–274. [10] P.C. Harris, S. Rossetti, Molecular genetics of autosomal recessive polycystic kidney disease, Mol. Genet. Metab. 81 (2004) 75–85. [11] J.Y. Kaimori, Y. Nagasawa, L.F. Menezes, M.A. Garcia-Gonzalez, J. Deng, E. Imai, L.F. Onuchic, L.M. Guay-Woodford, G.G. Germino, Polyductin undergoes notch-like processing and regulated release from primary cilia, Hum. Mol. Genet. 16 (2007) 942–956. [12] M.C. Hogan, L. Manganelli, J.R. Woollard, A.I. Masyuk, T.V. Masyuk, R. Tammachote, B.Q. Huang, A.A. Leontovich, T.G. Beito, B.J. Madden, M.C. Charlesworth, V.E. Torres, N.F. LaRusso, P.C. Harris, C.J. Ward, Characterization of PKD protein-positive exosome-like vesicles, J. Am. Soc. Nephrol. 20 (2009) 278–288. [13] T. Hiesberger, E. Gourley, A. Erickson, P. Koulen, C.J. Ward, T.V. Masyuk, N.F. Larusso, P.C. Harris, P. Igarashi, Proteolytic cleavage and nuclear translocation of fibrocystin is regulated by intracellular Ca2+ and activation of protein kinase C, J. Biol. Chem. 281 (2006) 34357–34364. [14] J.A. Follit, L. Li, Y. Vucica, G.J. Pazour, The cytoplasmic tail of fibrocystin contains a ciliary targeting sequence, J. Cell Biol. 188 (2010) 21–28. [15] M. Moser, S. Matthiesen, J. Kirfel, H. Schorle, C. Bergmann, J. Senderek, S. Rudnik-Schoneborn, K. Zerres, R. Buettner, A mouse model for cystic biliary dysgenesis in autosomal recessive polycystic kidney disease (ARPKD), Hepatology 41 (2005) 1113–1121. [16] M.A. Garcia-Gonzalez, L.F. Menezes, K.B. Piontek, J. Kaimori, D.L. Huso, T. Watnick, L.F. Onuchic, L.M. Guay-Woodford, G.G. Germino, Genetic interaction studies link autosomal dominant and recessive polycystic kidney disease in a common pathway, Hum. Mol. Genet. 16 (2007) 1940–1950. [17] A.R. Gallagher, E.L. Esquivel, T.S. Briere, X. Tian, M. Mitobe, L.F. Menezes, G.S. Markowitz, D. Jain, L.F. Onuchic, S. Somlo, Biliary and pancreatic dysgenesis in mice harboring a mutation in Pkhd1, Am. J. Pathol. 172 (2008) 417–429. [18] I. Kim, Y. Fu, K. Hui, G. Moeckel, W. Mai, C. Li, D. Liang, P. Zhao, J. Ma, X.Z. Chen, A.L. George Jr., R.J. Coffey, Z.P. Feng, G. Wu, Fibrocystin/polyductin modulates renal tubular formation by regulating polycystin-2 expression and function, J. Am. Soc. Nephrol. 19 (2008) 455–468. [19] S.S. Williams, P. Cobo-Stark, L.R. James, S. Somlo, P. Igarashi, Kidney cysts, pancreatic cysts, and biliary disease in a mouse model of autosomal recessive polycystic kidney disease, Pediatr. Nephrol. 23 (2008) 733–741. [20] E. Fischer, E. Legue, A. Doyen, F. Nato, J.F. Nicolas, V. Torres, M. Yaniv, M. Pontoglio, Defective planar cell polarity in polycystic kidney disease, Nat. Genet. 38 (2006) 21–23. [21] J. Zhang, M. Wu, S. Wang, J.V. Shah, P.D. Wilson, J. Zhou, Polycystic kidney disease protein fibrocystin localizes to the mitotic spindle and regulates spindle bipolarity, Hum. Mol. Genet. (2010). [22] T. Hiesberger, Y. Bai, X. Shao, B.T. McNally, A.M. Sinclair, X. Tian, S. Somlo, P. Igarashi, Mutation of hepatocyte nuclear factor-1beta inhibits Pkhd1 gene expression and produces renal cysts in mice, J. Clin. Invest. 113 (2004) 814–825. [23] T. Hiesberger, X. Shao, E. Gourley, A. Reimann, M. Pontoglio, P. Igarashi, Role of the hepatocyte nuclear factor-1beta (HNF-1beta) C-terminal domain in Pkhd1 (ARPKD) gene transcription and renal cystogenesis, J. Biol. Chem. 280 (2005) 10578–10586. [24] F. Verdeguer, S. Le Corre, E. Fischer, C. Callens, S. Garbay, A. Doyen, P. Igarashi, F. Terzi, M. Pontoglio, A mitotic transcriptional switch in polycystic kidney disease, Nat. Med. 16 (2009) 106–110. [25] M. Vujic, C.M. Heyer, E. Ars, K. Hopp, A. Markoff, C. Orndal, B. Rudenhed, S.H. Nasr, V.E. Torres, R. Torra, N. Bogdanova, P.C. Harris, Incompletely penetrant PKD1 alleles mimic the renal manifestations of ARPKD, J. Am. Soc. Nephrol. 21 (2010) 1097–1102.

186

E XP ER I ME NT AL CE L L R ES E AR CH 3 1 7 (2 0 11 ) 1 73 – 1 87

[26] Y. Wu, X.Q. Dai, Q. Li, C.X. Chen, W. Mai, Z. Hussain, W. Long, N. Montalbetti, G. Li, R. Glynne, S. Wang, H.F. Cantiello, G. Wu, X.Z. Chen, Kinesin-2 mediates physical and functional interactions between polycystin-2 and fibrocystin, Hum. Mol. Genet. 15 (2006) 3280–3292. [27] I. Kim, C. Li, D. Liang, X.Z. Chen, R.J. Coffy, J. Ma, P. Zhao, G. Wu, Polycystin-2 expression is regulated by a PC2-binding domain in the intracellular portion of fibrocystin, J. Biol. Chem. 283 (2008) 31559–31566. [28] R. Rohatgi, L. Battini, P. Kim, S. Israeli, P.D. Wilson, G.L. Gusella, L. M. Satlin, Mechanoregulation of intracellular Ca2+ in human autosomal recessive polycystic kidney disease cyst-lining renal epithelial cells, Am. J. Physiol. Renal. Physiol. 294 (2008) F890–F899. [29] W. Mai, D. Chen, T. Ding, I. Kim, S. Park, S.Y. Cho, J.S. Chu, D. Liang, N. Wang, D. Wu, S. Li, P. Zhao, R. Zent, G. Wu, Inhibition of Pkhd1 impairs tubulomorphogenesis of cultured IMCD cells, Mol. Biol. Cell 16 (2005) 4398–4409. [30] S.A. Gradilone, T.V. Masyuk, B.Q. Huang, J.M. Banales, G.L. Lehmann, B.N. Radtke, A. Stroope, A.I. Masyuk, P.L. Splinter, N.F. Larusso, Activation of transient receptor potential vanilloid 4 reduces the hyperproliferative phenotype of cystic cholangiocytes from an animal model of autosomal-recessive polycystic kidney disease, Gastroenterology (2010). [31] B. Goilav, L.M. Satlin, P.D. Wilson, Pathways of apoptosis in human autosomal recessive and autosomal dominant polycystic kidney diseases, Pediatr. Nephrol. 23 (2008) 1473–1482. [32] J.U. Becker, A.O. Saez, K. Zerres, O. Witzke, P.F. Hoyer, K.W. Schmid, A. Kribben, C. Bergmann, J. Nurnberger, The mTOR pathway is activated in human autosomal-recessive polycystic kidney disease, Kidney Blood Press. Res. 33 (2010) 129–138. [33] R.H. Whitehead, P.E. Van Eeden, M.D. Noble, P. Ataliotis, P.S. Jat, Establishment of conditionally immortalized epithelial cell lines from both colon and small intestine of adult H-2Kb-tsA58 transgenic mice, Proc. Natl Acad. Sci. USA 90 (1993) 587–591. [34] I. Kim, T. Ding, Y. Fu, C. Li, L. Cui, A. Li, P. Lian, D. Liang, D.W. Wang, C. Guo, J. Ma, P. Zhao, R.J. Coffey, Q. Zhan, G. Wu, Conditional mutation of Pkd2 causes cystogenesis and upregulates beta-catenin, J. Am. Soc. Nephrol. 20 (2009) 2556–2569. [35] M.Z. Zhang, W. Mai, C. Li, S.Y. Cho, C. Hao, G. Moeckel, R. Zhao, I. Kim, J. Wang, H. Xiong, H. Wang, Y. Sato, Y. Wu, Y. Nakanuma, M. Lilova, Y. Pei, R.C. Harris, S. Li, R.J. Coffey, L. Sun, D. Wu, X.Z. Chen, M.D. Breyer, Z.J. Zhao, J.A. McKanna, G. Wu, PKHD1 protein encoded by the gene for autosomal recessive polycystic kidney disease associates with basal bodies and primary cilia in renal epithelial cells, Proc. Natl Acad. Sci. USA 101 (2004) 2311–2316. [36] K. Matter, M.S. Balda, Signalling to and from tight junctions, Nat. Rev. Mol. Cell Biol. 4 (2003) 225–236. [37] P.D. Wilson, B. Goilav, Cystic disease of the kidney, Annu. Rev. Pathol. 2 (2007) 341. [38] D.D. Woo, S.Y. Miao, J.C. Pelayo, A.S. Woolf, Taxol inhibits progression of congenital polycystic kidney disease, Nature 368 (1994) 750–753. [39] A.K. Bhunia, K. Piontek, A. Boletta, L. Liu, F. Qian, P.N. Xu, F.J. Germino, G.G. Germino, PKD1 induces p21(waf1) and regulation of the cell cycle via direct activation of the JAK-STAT signaling pathway in a process requiring PKD2, Cell 109 (2002) 157–168. [40] X. Li, Y. Luo, P.G. Starremans, C.A. McNamara, Y. Pei, J. Zhou, Polycystin-1 and polycystin-2 regulate the cell cycle through the helix–loop–helix inhibitor Id2, Nat. Cell Biol. 7 (2005) 1102–1112. [41] F.A. Belibi, G. Reif, D.P. Wallace, T. Yamaguchi, L. Olsen, H. Li, G.M. Helmkamp Jr., J.J. Grantham, Cyclic AMP promotes growth and secretion in human polycystic kidney epithelial cells, Kidney Int. 66 (2004) 964–973. [42] T.F. Franke, PI3K/Akt: getting it right matters, Oncogene 27 (2008) 6473–6488.

[43] T. Yamaguchi, D.P. Wallace, B.S. Magenheimer, S.J. Hempson, J.J. Grantham, J.P. Calvet, Calcium restriction allows cAMP activation of the B-Raf/ERK pathway, switching cells to a cAMP-dependent growth-stimulated phenotype, J. Biol. Chem. 279 (2004) 40419–40430. [44] B.L. Stiles, PI-3-K and AKT: onto the mitochondria, Adv. Drug Deliv. Rev. 61 (2009) 1276–1282. [45] R. Schreck, U.R. Rapp, Raf kinases: oncogenesis and drug discovery, Int. J. Cancer 119 (2006) 2261–2271. [46] X.F. Zhang, J. Settleman, J.M. Kyriakis, E. Takeuchi-Suzuki, S.J. Elledge, M.S. Marshall, J.T. Bruder, U.R. Rapp, J. Avruch, Normal and oncogenic p21ras proteins bind to the amino-terminal regulatory domain of c-Raf-1, Nature 364 (1993) 308–313. [47] P.P. Roux, J. Blenis, ERK and p38 MAPK-activated protein kinases: a family of protein kinases with diverse biological functions, Microbiol. Mol. Biol. Rev. 68 (2004) 320–344. [48] M.A. Wozniak, K. Modzelewska, L. Kwong, P.J. Keely, Focal adhesion regulation of cell behavior, Biochim. Biophys. Acta 1692 (2004) 103–119. [49] S.K. Mitra, D.A. Hanson, D.D. Schlaepfer, Focal adhesion kinase: in command and control of cell motility, Nat. Rev. Mol. Cell Biol. 6 (2005) 56–68. [50] H.C. Chen, P.A. Appeddu, H. Isoda, J.L. Guan, Phosphorylation of tyrosine 397 in focal adhesion kinase is required for binding phosphatidylinositol 3-kinase, J. Biol. Chem. 271 (1996) 26329–26334. [51] S. Nishio, M. Hatano, M. Nagata, S. Horie, T. Koike, T. Tokuhisa, T. Mochizuki, Pkd1 regulates immortalized proliferation of renal tubular epithelial cells through p53 induction and JNK activation, J. Clin. Invest. 115 (2005) 910–918. [52] F. Qian, F.J. Germino, Y. Cai, X. Zhang, S. Somlo, G.G. Germino, PKD1 interacts with PKD2 through a probable coiled-coil domain, Nat. Genet. 16 (1997) 179–183. [53] C. Montell, L. Birnbaumer, V. Flockerzi, The TRP channels, a remarkably functional family, Cell 108 (2002) 595. [54] T. Yamaguchi, S.J. Hempson, G.A. Reif, A.M. Hedge, D.P. Wallace, Calcium restores a normal proliferation phenotype in human polycystic kidney disease epithelial cells, J. Am. Soc. Nephrol. 17 (2006) 178–187. [55] T. Yamaguchi, J.C. Pelling, N.T. Ramaswamy, J.W. Eppler, D.P. Wallace, S. Nagao, L.A. Rome, L.P. Sullivan, J.J. Grantham, cAMP stimulates the in vitro proliferation of renal cyst epithelial cells by activating the extracellular signal-regulated kinase pathway, Kidney Int. 57 (2000) 1460–1471. [56] D.D. Woo, A.P. Tabancay Jr., C.J. Wang, Microtubule active taxanes inhibit polycystic kidney disease progression in cpk mice, Kidney Int. 51 (1997) 1613–1618. [57] Y. Tao, J. Kim, S. Faubel, J.C. Wu, S.A. Falk, R.W. Schrier, C.L. Edelstein, Caspase inhibition reduces tubular apoptosis and proliferation and slows disease progression in polycystic kidney disease, Proc. Natl Acad. Sci. USA 102 (2005) 6954–6959. [58] A. Boletta, F. Qian, L.F. Onuchic, A.K. Bhunia, B. Phakdeekitcharoen, K. Hanaoka, W. Guggino, L. Monaco, G.G. Germino, Polycystin-1, the gene product of PKD1, induces resistance to apoptosis and spontaneous tubulogenesis in MDCK cells, Mol. Cell 6 (2000) 1267–1273. [59] T. Wegierski, D. Steffl, C. Kopp, R. Tauber, B. Buchholz, R. Nitschke, E.W. Kuehn, G. Walz, M. Köttgen, TRPP2 channels regulate apoptosis through the Ca(2+) concentration in the endoplasmic reticulum, EMBO J. 28 (2009) 490–499. [60] Y. Lim, I. Han, J. Jeon, H. Park, Y.Y. Bahk, E.S. Oh, Phosphorylation of focal adhesion kinase at tyrosine 861 is crucial for Ras transformation of fibroblasts, J. Biol. Chem. 279 (2004) 29060–29065. [61] J.A. Lunn, R. Jacamo, E. Rozengurt, Preferential phosphorylation of focal adhesion kinase tyrosine 861 is critical for mediating an anti-apoptotic response to hyperosmotic stress, J. Biol. Chem. 282 (2007) 10370–10379.

E XP E RI ME N T AL C ELL R ES E AR C H 3 1 7 ( 2 0 11 ) 1 7 3– 1 87

[62] H.C. Chen, J.L. Guan, Association of focal adhesion kinase with its potential substrate phosphatidylinositol 3-kinase, Proc. Natl Acad. Sci. USA 91 (1994) 10148–10152. [63] M.B. Calalb, T.R. Polte, S.K. Hanks, Tyrosine phosphorylation of focal adhesion kinase at sites in the catalytic domain regulates kinase activity: a role for Src family kinases, Mol. Cell. Biol. 15 (1995) 954–963. [64] D.D. Schlaepfer, S.K. Hanks, T. Hunter, P. van der Geer, Integrin-mediated signal transduction linked to Ras pathway by

187

GRB2 binding to focal adhesion kinase, Nature 372 (1994) 786–791. [65] Y. Fu, I. Kim, P. Lian, A. Li, L. Zhou, C. Li, D. Liang, R.J. Coffey, J. Ma, P. Zhao, Q. Zhan, G. Wu, Loss of Bicc1 impairs tubulomorphogenesis of cultured IMCD cells by disrupting E-cadherin-based cell–cell adhesion, Eur. J. Cell Biol. 89 (2010) 428–436. [66] T.D. Schmittgen, K.J. Livak, Analyzing real-time PCR data by the comparative C(T) method, Nat. Protoc. 3 (2008) 1101–1108.