Fibrocystin/Polyductin Modulates Renal Tubular Formation by Regulating Polycystin-2 Expression and Function
Ingyu Kim*,
Yulong Fu,
Kwokyin Hui,
Gilbert Moeckel,
Weiyi Mai*,||,
Cunxi Li*,¶,
Dan Liang*,
Ping Zhao,
Jie Ma,
Xing-Zhen Chen**,
Alfred L. George, Jr.*,
Robert J. Coffey*,¶,
Zhong-Ping Feng and
Guanqing Wu*,,¶
Departments of * Medicine, Pathology, and ¶ Cell and Developmental Biology, Vanderbilt University, Nashville, Tennessee; Division of Translational Cancer Research and Therapy, Cancer Hospital and Institute, Chinese Academy of Medical Sciences, Beijing, China; Department of Physiology, University of Toronto, Toronto, Ontario, Canada; ** Department of Physiology, University of Alberta, Edmonton, Alberta, Canada; and || Department of Internal Medicine, First Affiliated Hospital, Sun Yat-sen University, Guangzhou, China
Correspondence: Dr. Guanqing Wu, Division of Genetic Medicine, Department of Medicine and Cell and Developmental Biology, Vanderbilt University, 539 LH, 2215 Garland Avenue, Nashville, TN 37232. Phone: 615-936-1761; Fax: 615-936-2661; E-mail: guanqing.wu{at}vanderbilt.edu
Received for publication July 16, 2007.
Accepted for publication September 4, 2007.
Autosomal recessive polycystic kidney disease is caused by mutationsin PKHD1, which encodes the membrane-associated receptor-likeprotein fibrocystin/polyductin (FPC). FPC associates with theprimary cilia of epithelial cells and co-localizes with thePkd2 gene product polycystin-2 (PC2), suggesting that thesetwo proteins may function in a common molecular pathway. Forinvestigation of this, a mouse model with a gene-targeted mutationin Pkhd1 that recapitulates phenotypic characteristics of humanautosomal recessive polycystic kidney disease was produced.The absence of FPC is associated with aberrant ciliogenesisin the kidneys of Pkhd1-deficient mice. It was found that theCOOH-terminus of FPC and the NH2-terminus of PC2 interact andthat lack of FPC reduced PC2 expression but not vice versa,suggesting that PC2 may function immediately downstream of FPCin vivo. PC2-channel activities were dysregulated in culturedrenal epithelial cells derived from Pkhd1 mutant mice, furthersupporting that both cystoproteins function in a common pathway.In addition, mice with mutations in both Pkhd1 and Pkd2 hada more severe renal cystic phenotype than mice with single mutations,suggesting that FPC acts as a genetic modifier for disease severityin autosomal dominant polycystic kidney disease that resultsfrom Pkd2 mutations. It is concluded that a functional and molecularinteraction exists between FPC and PC2 in vivo.
Autosomal dominant polycystic kidney disease (ADPKD) and autosomalrecessive polycystic kidney disease (ARPKD) are common humangenetic disorders that are characterized by numerous, expandingfluid-filled cysts in both kidneys and other duct/tubule-containingorgans.1,2 ADPKD is inherited as a dominant trait, occurringrelatively late in life, and is characterized by focal outpouchingsof spherical cysts in the renal tubules. In contrast, ARPKDis inherited as a recessive trait, usually presents during perinatallife, and is characterized by numerous spindle-shaped renalcysts. Because of these differences, ADPKD and ARPKD are usuallyconsidered two distinct diseases in clinical practice.3
ADPKD affects 1 in every 500 to 1000 people and results frommutations in either of at least two causal genes, PKD1 and PKD2,leading to nearly identical clinical manifestations. Mutationsin PKD2, which maps to chromosome 4q21–23, are responsiblefor approximately 15% of familial ADPKD cases.4,5PKD2 has anapproximately 5.4-kb transcript and encodes the 968–aminoacid gene product polycystin-2 (PC2). PC2 is predicted to bean integral membrane protein with six putative transmembranedomains and intracellular NH2- and COOH-termini.4 PC2 has beenreported to be a receptor-operated, nonselective cation channel;it is also referred to as TRPP2, because it is considered amember of the trp superfamily.6,7 Conversely, mutations in PKD1are responsible for approximately 85% of ADPKD cases. PKD1 liesin an approximately 53-kb region on chromosome 16p13.3 and yieldsa 14-kb transcript that encodes a 4303–amino acid integralmembrane protein.8 The gene product of PKD1, polycystin-1 (PC1),was reported to interact with PC2 and regulate Pkd2-channelactivity.9–12
ARPKD is one of the common hereditary renal cystic diseasesin infants and children.13 The estimated incidence of ARPKDis approximately 1 in 20,000 live births.14 The clinical characteristicsof ARPKD include ectasia of the renal collecting ducts and hepaticbiliary ducts with associated renal and hepatic fibrosis.15Approximately 50% of patients with ARPKD present with theirdisease as neonates16 and are born with two very large kidneyswith 60 to 90% of the renal tubules being ectatic.15 These neonatessuffer a 30% mortality rate as a result of respiratory and/orrenal dysfunction.17 Patients who have ARPKD and survive theirfirst year of life have a more optimistic prognosis: 75% reachage 5, and only half develop ESRD.16–18 In rare cases,patients with ARPKD survive beyond age 60.19
The most common complications of ARPKD include hypertension(60 to 100%), portal hypertension owing to severe hepatic fibrosisor Caroli disease (30 to 75%), and chronic lung disease (approximately11%).17 In addition, growth retardation,15 intracranial aneurysms,20,21and adrenal insufficiency22 can be seen in patients with ARPKD.
ARPKD is caused by mutations in PKHD1. This gene consists ofat least 86 exons spanning 470 kb on chromosome 6p12 and producesa 16-kb transcript. The longest open reading frame is predictedto include 66 exons and to encode the 4074–amino acidmembrane-associated receptor-like protein fibrocystin/polyductin(FPC).23–26 It was shown that FPC is associated with thebasal bodies/primary cilia of epithelial cells27–30 andco-localizes with PC2 within the cell.31 These observationssuggest the possibility that FPC and PC2 may function in a commonmolecular pathway in vivo.
To investigate a potential functional relationship between FPCand PC2, we generated and characterized a mouse model with agene-targeted mutation in Pkhd1. We discovered that lack ofFPC was associated with the downregulation of PC2 expressionin vivo. Renal cyst formation is more severe in mice that aretransmutant in Pkhd1 and Pkd2 (Pkd2tm2Som)32,33 than in micethat are heterozygous for the Pkd2 mutation alone. In addition,we found that Pkd2-channel activities were dysregulated in primarycultures of renal epithelial cells derived from Pkhd1 mutantmice. The in vivo observations demonstrated that FPC and PC2functionally interacted and resided in a common molecular pathway.
Generation of Pkhd1-Deficient Mutant Mice
Because the Northern probe containing exons 15 to 16 of PKHD1displayed highly mRNA expression signal in the kidney tissue,we designed a targeting construct that not only disrupted exon15 but also deleted exon 16, resulting in a mutant allele designatedPkhd1e15GFP16 (Pkhd1–) (Figure 1, A through D). Becauseour gene-targeting construct included an in-frame green fluorescenceprotein (GFP) reporter gene with a predicted fusion proteinof 130 kD (Figure 1A), anti-GFP immunoprecipitation assays showedthat only the mutant mice expressed the GFP reporter protein(Figure 1E). In addition, with the use of a panel of previouslygenerated anti-FPC antibodies,31 Western blot analyses wereable to detect the expected immunoreactive bands at >400kD in the wild-type (WT) and heterozygous mouse kidneys (Figure 1F).These immunoreactive bands were not detected in protein lysatefrom Pkhd1–/– mouse kidneys, suggesting that thehomozygous mice lacked full-length, functional FPC.
Figure 1.Pkhd1 gene-targeting construct and molecular analysis of the specific targeting event at the Pkhd1 locus. (A) Schematic representation of the gene-targeting strategy. (I) Partial genomic map showing exons 15 to 21 of Pkhd1. (II) Pkhd1e15GFP16 targeting vector in which exon 16 was deleted and exon 15 was disrupted. (III) Partial map of the mutant Pkhd1 allele. Nh, NheI; S, SphI; P, PstI; B, BamHI; K, KpnI. (B) Tail biopsy DNA from mice with the germline targeted mutation in Pkhd1 were digested with PstI and hybridized with probe 1 from outside the targeted region (AIII). The expected 3.3-kb WT band was observed in WT and Pkhd1 heterozygous mice. A mutant 2.2-kb band was seen in Pkhd1 heterozygous and homozygous mice. (C) Tail biopsy DNA were digested with BamHI and were hybridized with the Pkhd1 exon 16 probe. The radioactive signal was not observed in Pkhd1 homozygous mice. (D) A 3.8-kb cDNA fragment containing exons 37 to 43 of Pkhd1 served as a Northern probe to detect Pkhd1 in the total RNA of adult kidneys. Pkhd1–/– mice showed an absence of Pkhd1 mRNA (D, top). The 28S rRNA band images provided a total RNA loading control (D, bottom). (E) Because the Pkhd1 mutant alleles contained an in-frame GFP reporter gene, we used an anti-GFP antibody to detect GFP expression. GFP immunoreactivity was detected at the expected size of 130 kD in the adult kidneys of Pkhd1–/– and Pkhd1+/– mice. (F) Western blot detection of FPC. Antibodies hAR-Cm3C10 and hAR-Nm3G12, which recognize the COOH- and NH2-portions of FPC, respectively, were used to detect immunoreactivity to FPC in the adult kidneys of WT, Pkhd1+/–, and Pkhd1–/– mice. FPC expression was significantly reduced in Pkhd1–/– kidneys compared with WT and Pkhd1+/–.
Pkhd1-Deficient Mice Exhibit the Phenotypic Characteristics of Human ARPKD
To determine whether the Pkhd1-deficient mice would be a suitablemodel for ARPKD, we intercrossed heterozygous Pkhd1 mice toproduce homozygous (Pkhd1–/–) progeny. Pkhd1–/–mice were born at a frequency of 18%, lower than the expectedMendelian ratio, indicating a loss of approximately 5 to 10%of the Pkhd1–/– mice during the embryonic or perinatalperiod (Figure 2A). This suggests that the Pkhd1–/–genotype may lead to embryonic lethality. Mice that reachedadulthood were used in a cohort study. Kaplan-Meier analysisshowed that only 25% of Pkhd1–/– mice survived beyond12 mo compared with 60% of Pkhd1+/– mice (P < 0.001)and 90% of WT mice (P < 0.001), suggesting that some adultPkhd1–/– mice died as a result of their diseasephenotypes (Figure 2B). Although there was a trend toward increasedmortality in Pkhd1+/– mice compared with WT mice, thisdid not reach statistical significance (P = 0.075).
Figure 2. Survival analyses of Pkhd1 mutant mice. (A) Genotype and survival rate of Pkhd1 mutant mice from embryo to adulthood. (B) Kaplan-Meier survival curves for WT, Pkhd1+/–, and Pkhd1–/– mice are shown. The mice in these cohorts were observed for >1 yr. Survival of the Pkhd1–/– mice differed significantly from that of the WT and Pkhd1+/– cohorts (P < 0.001).
Pkhd1–/– mice that escaped embryonic lethality andsurvived into adulthood exhibited mild to severe tubular dilationor cyst formation in the kidney and liver accompanied by fibrosisand necrosis (Figure 3). The severity of cysts and the age atthe onset of disease varied among individual Pkhd1–/–mice. Approximately 10% of the mice had early-onset, microscopicmalformations in the renal tubules (Figure 3, B through G),whereas others (approximately 60%) survived beyond 1 yr andhad late-onset cystic phenotypes. The extent of cystic changein the liver of Pkhd1–/– mice was generally moresevere than that observed in the kidneys (Figure 3, H throughK). These observations indicate that the targeted mutation inPkhd1 induces cystogenesis in the kidneys and livers of themice. Cystic or dilated-duct phenotypes were also seen in thepancreas (Figure 4, A versus B) and brain (Figure 4, C versusD) of Pkhd1–/– mice. In the gastrointestinal tract,hemorrhagic, and ulcer-like lesions were observed (Figure 4,E through I).
Figure 3. Hepatorenal cysts and tubular ectasia in Pkhd1–/– mice. (A) A kidney tissue section shows normal glomeruli and a normal tubulointerstitial compartment in a 2-wk-old WT mouse. (B) Patchy dilation of proximal tubules and focal dilation of papillary collecting ducts were seen in a Pkhd1–/– littermate. (C) There was a marked increase in the degree of tubular dilation with flattening of the tubular epithelial cells (arrow) in the kidney of a 4-wk-old Pkhd1–/– mouse. (D) At 2 mo, there was a further increase in the severity of tubular dilation, in both the cortical proximal tubules and the medullary collecting duct tubules. Moreover, there was an increased expansion of Bowman's space (arrow), an increase in mesangial cellularity, and a reduction of glomerular capillary loop formation. (E) At 4 mo, in the whole-mount Pkhd1–/– kidney, there was a dramatic increase in tubular dilation, involving >80% of the cortex and medulla. (F) A high-power view of E at the cortical region of the kidney shows persistent expansion of Bowman's space with a mild decrease in glomerular size and segmental mesangial hypercellularity. The proximal tubules were dilated. (G) The medullary sections of a higher-power view of E also showed occasional small cysts lined by a single layer of epithelium. Red blood cells were seen within the tubular lumen. (H) Liver section showed normal histology of the 2-wk-old WT mouse. (I) Liver section of the homozygous littermates showed dilation of the biliary ducts. (J) Transillumination of an affected liver in the 2-mo-old Pkhd1–/– mice showed cyst formation (left arrow) and dilated ducts (right arrow). (K) There were focally enlarged cysts with areas of hemorrhage and necrosis (arrow) within the liver parenchyma in the same mouse. Bar = 20 µm in A, B, H, and I; 10 µm in C, D, F, G, and K; and 1 mm in E and J.
Figure 4. Abnormal phenotypes were seen in the extrahepatorenal organs of Pkhd1–/– mice. (A) Pancreatic sections from a 6-mo-old WT mouse showed normal histology. (B) In the same homozygous littermates, there was dilation of both the small and large pancreatic ducts (upper arrow) with a marked increase in interstitial fibrosis (lower arrow). (C) Normal brain section from a 2-mo-old WT mouse. (D) Brain tissue sections showed multiple, diffuse, small- to medium-sized vacuoles in a homozygous littermate (arrow). (E) The stomach mucosa of a 2-mo-old homozygous mouse showed an ulcerated lesion (arrow) that was raised with beaded borders. (F) The colon of the same mouse showed small areas of subserosal hemorrhage. (G) The mucosal aspect of the colon from a 2-mo-old homozygous mouse showed a small, ulcerated lesion with hemorrhage (arrow). (H) The microscopic colon section of the sample in G revealed a submucosal lymphocytic infiltrate with focal necrosis and erosion of the overlying epithelium. (I) Gross view of the small intestinal mucosa with mild superficial hemorrhage (arrow). Histologic sections of the sample in I showed submucosal edema, lymphocytic infiltrate, hemorrhage, and erosion of the overlying mucosa. Bar = 25 µm in A through D; 5 µm in H; 1 mM in E through G and I; and 10 µm in I inset.
Because the Pkhd1 mutant allele carries an in-frame GFP reportergene in exon 15, we used immunohistochemistry (IHC) and immunofluorescence(IF) staining with anti-GFP antibodies to detect GFP distributionin the organs of homozygous mutant mice. GFP-positive signals,defining the expression pattern of Pkhd1, were observed in thetubular epithelia of the kidneys (Figure 5, A, B, and D), gastrointestinaltract (Figure 5, A and C), bronchioles/trachea (Figure 5, Eand F), ependymal cells lining the ventricles of the brain (Figure 5G),and hepatobiliary epithelial cells (Figure 5, H through J).
Figure 5. GFP expression in Pkhd1 mutant mice. (A) E15.5 Pkhd1–/– kidney, liver, adrenal gland, and gastrointestinal tract stained with IHC using an anti-GFP polyclonal antibody. Positive signals were seen in the cortical adrenal cells (left arrow), periportal liver cells (right arrow), gastrointestinal tract (white box), and weakly positive staining was seen in the renal tubules (black box). (B) Higher magnification of the black box in A shows positive staining of the renal epithelia (arrow). (C) Higher magnification of the white box in A shows strong positive staining in the mucosal cells of the colon (arrow). (D) The differential interface contrast view showed GFP-positive staining (arrows) in the renal tubule epithelia of 4-mo-old Pkhd1+/– (D, top) and Pkhd1–/– (D, bottom) littermates. (E) Positive GFP staining was detected in an alveolar bronchiole of the lung in a 2-mo-old homozygous mouse (arrow). (F) Positive GFP was also seen in the tracheal epithelium of the same mouse (arrow). (G) GFP-positive staining (red) appeared in the ependymal cells lining the ventricles of the brain in a 2-mo-old homozygous mouse (arrow). Yo-pro (green) was used to stain the nuclei of cells. (H) GFP-positive staining (red) was observed in the diseased liver of a 2-mo-old homozygous mouse (arrows). (I) Cytokeratin 7–positive staining (green), a marker for epithelial cells, outlined the biliary epithelial structures (arrows). (J) The merged confocal image showed that GFP co-localized with cytokeratin 7. Bar = 30 µm in A; 15 µm in B and C; and 10 µm in D through J.
Lack of FPC Exhibits Aberrant Ciliogenesis in the Renal Epithelial Cells
FPC was demonstrated to localize to the primary cilium and/orbasal bodies of renal tubular epithelia,27–31 and malformationof the cilia was shown to induce cyst formation in the kidneys34,35;therefore, we began to determine whether the lack of FPC alsodisrupts ciliogenesis in Pkhd1-deficient mice. We used IF withan anti-acetylated -tubulin antibody to examine the number andmorphology of renal primary cilia in 6-mo-old littermates. Comparedwith WT mice, there were far fewer primary cilia in the renaltubular epithelia of Pkhd1–/– mice (Figure 6, Aversus B), indicating that lack of FPC might reduce ciliogenesisin the kidneys. The ciliary defects seem to be most severe inthe cortical proximal tubules of the Pkhd1–/– kidneys.In addition, confocal images also displayed similar ciliarychanges in corresponding regions of littermate kidneys (Figure 6,C versus D). Scanning electron microscopy confirmed these resultsin 3-mo-old littermates with or without the targeted Pkhd1 mutation(Figure 6, E versus F). For further validation of these findings,primary culture of renal epithelial cells derived from 2-mo-oldPkhd1–/– and WT littermates was used to determinewhether the ciliary malformation could be observed in vitro.Compared with the cultured WT cells, a shortened ciliary structureand decreased ciliary staining were seen in the Pkhd1–/–cells (Figure 6, G versus H). The cilia stained in approximately94% of WT cells and in fewer than 34% of Pkhd1–/–cells (P < 0.001; Figure 6I). The mean length of primarycilia was 10 µm in cultured WT cells and was <5 µmin Pkhd1–/– littermate cells (P < 0.001; Figure 6J).That short or absent cilia were observed in Pkhd1–/–cells but not in WT cells suggests that lack of FPC causes defectsin ciliogenesis in renal epithelial cells in vivo.
Figure 6. Lack of FPC induces aberrant ciliogenesis in the Pkhd1–/– kidneys. (A) A common ciliary marker, anti-acetylated -tubulin antibody, was used for IF staining of kidney sections from 6-mo-old WT and Pkhd1–/– littermates. Ciliary structures (arrows) were abundantly observed in the WT kidneys. (B) Decreased ciliary staining (arrows) was seen in the corresponding cortical region of Pkhd1–/– mouse kidneys. (C) The confocal images also showed normal ciliary structures in the 4-mo-old WT kidneys (arrows). Lotus Tetragonolobus Lectin (green) was used to stain renal proximal tubules. (D) The ciliary structures in the Pkhd1–/– littermates were reduced in number and shorter (arrow) than WT controls. (E) Scanning electron microscopy showed the normal primary cilia (arrows) of a 3-mo-old WT kidney. (F) The ciliary structures in the Pkhd1–/– kidney are shorter than those in the WT littermates, in the similar regions of the kidney. (G) Primary cultures of renal epithelia derived from the 2-mo-old WT and Pkhd1–/– kidneys were stained with an anti-acetylated -tubulin antibody. The confocal images showed normal ciliary structures (arrows) in both top (G, top) and lateral views (G, bottom). Blue To-pro was used to stain nuclei. (H) In the primary cultured epithelial cells from the Pkhd1–/– littermate, confocal images showed ciliary structures that were shorter and fewer in number than those in control cells (arrows). The confocal top and lateral views were composed of multiple sections (approximately 0.5 µm thick and up to 16 layers) that were projected onto one plane to present the ciliary staining patterns. (I) One hundred individual primary cultured cells from five random high-power fields (x1000) were numbered; the cell number and positive cilium-staining rates are shown in I. In WT cells, 94% of cells stained positive for cilia, compared with 34% of Pkhd1–/– littermate cells (P < 0.001). (J) The length of 50 individual primary cilia of cultured cells from three random high-power fields was measured using lateral views of the confocal images; the average length of the primary cilia was calculated and showed in J. The primary cilium length is approximately 10 µm in WT and <5 µm in Pkhd1–/– littermate cells (P < 0.005). Bar = 10 µm in A through D; 2 µm in E and F; and 5 µm in G and H.
Trans-Mutant Mice in Pkhd1 and Pkd2 Accelerate Renal Cyst Disease Progression
Several in vitro studies from our group and others have shownthat FPC and PC2 co-localize to the basal bodies/primary ciliaof renal epithelial cells and are able to form a molecular complexand function in the same signaling pathway.31,36 For furthervalidation of this finding in vivo, the phenotypic effects oftransheterozygosity for Pkhd1 and Pkd2 were examined. We intercrossedPkhd1+/– and Pkd2+/– mutant mice to produce cohortsof age-matched littermates. The four genotypes of interest (Pkhd1–/–,Pkhd1–/–/Pkd2+/–, and WT) were obtained fromlive-born progeny. Cohorts were killed at 1 mo. Gross inspectiondid not reveal cysts on the surface of the kidneys in any ofthe genotypes.
Because spherical renal cysts (at least three times the normaldiameter of the proximal tubule) are characteristic of ADPKDand massive, dilated, spindle-shaped renal tubules are moretypical of the ARPKD phenotype, we used sphere-shape cysts torepresent ADPKD-like renal cysts.1–3 In mice with Pkhd1–/–/Pkd2+/–alleles, spherical cysts were distributed at the medullary andcortical regions on a background of Pkhd1–/–-specificdilated tubules (Figure 7A). By statistical analysis, Pkhd1–/–/Pkd2+/–mice had significantly greater numbers of spherical cysts thanother genotypes (P < 0.05; Figure 7B). Western blot analysesof lysates from Pkd2+/– and Pkhd1–/–/Pkd2+/–littermates demonstrated that loss of FPC in the adult mousekidney reduced PC2 expression in vivo (Figure 7C). These studiesindicate that lack of FPC exacerbates the severity of ADPKD.
Figure 7. The cystic phenotype of mice trans-mutant for Pkhd1 and Pkd2. (A) In a representative hematoxylin- and eosin-stained section, spherical renal cysts (arrows; diameter >50 µm was considered as renal cyst) were identified in a 1-mo-old Pkhd1–/–/Pkd2+/– double-mutant mouse. (B) Numbers of cysts are presented as means ± SD for four genotypes at the age of 1 mo (n = number of animals in each group). The increase in spherical renal cysts in Pkhd1–/–/Pkd2+/–trans-mutant mice was significantly higher than other genotypes (*P < 0.05). (C) Western blot of protein lysates from Pkd2+/– 1-mo-old mouse kidney with or without the Pkhd1–/– mutation was performed using the anti-PC2 antibody hPKD2-Cm1A11. A significant decrease in immunoreactivity in lysates from the Pkhd1–/– mutant mice indicates that lack of FPC reduces PC2 expression in vivo. Bar = 30 µm in A.
FPC and PC2 Interact In Vivo and Form a Molecular Complex in Renal Epithelia
Because a lack of FPC inhibited the expression of PC2 and inducedsevere cystic phenotypes in the Pkhd1–/–/Pkd2+/–mouse model (Figure 7), we decided to analyze the molecularrelationship between FPC and PC2. We used our mAb hPKD2-Cm1A11,which is directed against the intracellular COOH-terminus ofPC2, to examine protein level in tissue lysates from WT, Pkhd1+/–,and Pkhd1–/– littermates at embryonic day 13.5 (E13.5).Compared with WT embryos, PC2 expression was decreased in thePkhd1–/– embryos, suggesting that lack of FPC downregulatesPC2 expression in vivo (Figure 8A). In addition, IHC stainingwith the anti-PC2 polyclonal antibody hPKD2-Cp was used to examineexpression level of PC2 in the Pkhd1–/– 1-mo-oldkidneys. In the renal cortex, significantly less staining wasobserved in Pkhd1–/– kidneys than in WT littermates(Figure 8B), giving further evidence that lack of FPC disruptsnormal PC2 expression in vivo; however, quantitative PCR didnot reveal a significant difference in Pkd2 expression amongPkhd1–/–, Pkhd1+/–, and WT 1-mo-old littermatekidneys (data not shown), suggesting that lack of FPC may affectonly the synthesis and/or stability of PC2 in vivo.
Figure 8. Molecular relationship between FPC and PC2. (A) Using the anti-PC2 mAb hPKD2-Cm1A11, Western blot of duplicate protein lysates from WT, Pkhd1+/–, and Pkhd1–/– E13.5 littermates showed a significant downregulation of PC2 in Pkhd1–/– embryos, indicating that lack of FPC inhibits PC2 expression in vivo. An anti–β-actin antibody was used for a protein-loading control. (B) In comparison with the WT littermate (left), IHC staining with the anti-PC2 polyclonal antibody hPKD2-Cp showed a significant decrease in PC2 expression in the cortical region of the 1-mo-old Pkhd1–/– kidney (right). (C) Duplicate lysates from E13.5 WT, Pkhd1+/–, and Pkhd1–/– littermates were used to perform a co-IP Western using the anti-FPC antibody hAR-Nm3G12 to IP and the anti-PC2 antibody hPKD2-Cm1A11 to detect PC2 expression. Positive immunoreactivity was seen in the WT embryo, and progressively reduced immunoreactivities were seen in the Pkhd1+/– and Pkhd1–/– littermates, suggesting that FPC binds to PC2 in vivo. (D) Lysates from E13.5 WT, Pkd2+/–, and Pkd2–/– littermates were used to perform a co-IP Western using the anti-PC2 antibody to IP and the anti-FPC antibody to detect FPC expression. Positive immunoreactivity was seen in the WT and Pkd2+/– littermates, but no immunoreactivity was observed in the Pkd2–/– littermate, providing further evidence that FPC physically interacts with PC2 in vivo. (E) There was no change in FPC expression in Western blot analysis among the WT and Pkd2–/– littermates, indicating that the downregulation of PC2 does not affect FPC expression. (F) HA- and Flag-tagged expression vectors, in which the COOH-terminus of FPC (FPC-C-Flag) and the NH2-terminus of PC2 (PC2-N-HA) were constructed in-frame, were transiently co-transfected into HEK293 cells. Using an anti-Flag antibody to IP and an anti-HA antibody to detect the NH2-terminus of PC2, positive immunoreactivity was seen only in the co-transfected sample, indicating that the COOH-terminus of FPC physically interacts with the NH2-terminus of PC2 in vitro. (G) The same FPC-C-Flag expression vector was transiently co-transfected into HEK293 cells with an expression vector containing the human full-length PKD2 cDNA (PC2-Full). The anti-PC2 antibody hPKD2-Cm1A11 was used for IP and an anti-Flag antibody was used to detect the COOH-terminus of FPC. Strong positive immunoreactivity was seen only in the co-transfected sample, and weak immunoreactivity was detected in the FPC-C-Flag single-transfected sample, indicating that either exogenously transfected or endogenously expressed PC2 immunoprecipitates with FPC-C-Flag construct. This further confirms that the COOH-terminus of FPC physically interacts with the NH2-terminus of PC2. (H) Using the hAR-C2p antibody against the COOH-terminus of FPC to preincubate with FPC-C-Flag single-transfected protein lysates, positive immunoreactivity was seen only in the nonpreincubated co-IP sample, whereas the immunoreactivity was missing in the preincubated co-IP sample. Bar = 30 µm in B.
To investigate whether FPC and PC2 physically interact, we usedtissue lysates from E13.5 WT, Pkhd1+/–, and Pkhd1–/–littermates to perform a co-immunoprecipitation assay (co-IP)with antibodies against FPC and PC2. We found that FPC immunoprecipitatedwith PC2 in tissue from WT and Pkhd1+/– littermates butnot from the negative control Pkhd1–/– littermates(Figure 8C). In addition, we used tissue lysates from an E13.5Pkd2-mutant set in another co-IP assay. Similar to the previousresult, FPC immunoprecipitated with PC2 in tissue from WT andPkd2+/– littermates but not in tissue from Pkd2–/–littermates (Figure 8D). Immunoreactivity was stronger in WTthan in heterozygous littermates and was not detected in homozygouslittermate controls, providing strong evidence that FPC mayphysically interact with PC2 in vivo. It is interesting thatno reduction of FPC expression was seen in Pkd2–/–littermates by Western analysis, suggesting that lack of PC2does not affect the level FPC in vivo (Figure 8E).
We constructed serial Hemagglutinin (HA) and Flag-tagged expressionvectors that contain intracellular portions of FPC and PC2,respectively. Positive immunoreactivity was seen in co-IP assaysbetween the COOH-terminal portion of FPC (FPC-C-Flag) and theNH2-terminal portion of PC2 (PC2-N-HA; Figure 8F). Immunoreactivitywas not observed between COOH-terminal portions of FPC (FPC-C-Flag)and PC2 (PC2-C-HA; data not shown). This suggests that the interactionbetween FPC and PC2 occurs via the intracellular COOH-terminaltail of FPC and NH2-terminal portion of PC2. For confirmationof this, HEK293 cells were transiently co-transfected with bothan expression vector containing the full-length human PKD2 cDNA(PC2-Full) and the aforementioned FPC-C-Flag vector. With theuse of an anti-PC2 antibody to immunoprecipitate and an anti-Flagantibody for detection by Western blot, a strong band was seenin lysate from the co-transfected cells. Weak immunoreactivitywas observed in lysate from cells that were transfected onlywith FPC-C-Flag. These data suggest that FPC-C-Flag can be co-immunoprecipitatedwith either PC2 introduced exogenously or PC2 endogenous toHEK293 cells (Figure 8G). In addition, when lysates from FPC-C-Flag–transfectedHEK293 cells were preincubated with an antibody against theCOOH-terminal tail of FPC (hAR-C2p), the detection of PC2-N-HAand FPC-C-Flag co-IP was blocked (Figure 8H). This result providesfurther evidence of a specific interaction between the C-terminaltail of FPC and the N-terminal portion of PC2.
Lack of Pkhd1 Disrupts Pkd2-Induced Cation Channel Activities
To study further whether FPC and PC2 functionally interact,we used whole-cell patch clamp recordings to characterize Pkd2-channelactivities in Pkhd1-deficient cells. We first tested whetherPkd2-induced whole-cell current could be detected under publishedconditions37,38 and then investigated whether the recorded channel-likecurrent is Pkd2-specific. First, primary cultured fibroblastsderived from E13.5 WT and Pkd2–/– embryos were testedto determine whether whole-cell current densities were dysregulatedby the lack of Pkd2. The recorded current from the Pkd2–/–fibroblasts was significantly lower than that from the WT fibroblastsat all voltages tested, indicating that reduction of functionalchannel activities is caused by missing Pkd2 expression (P <0.005; Figure 9A). Next, we used the same approach to test thePkhd1-silenced inner medullary collecting duct (IMCD) cells(IMCDshRNA3e23) and WT control cells (IMCDsh15) that we hadgenerated previously.39 It is interesting that Pkhd1-silencedIMCD cells exhibited a similar channel reduction, indicatingthat inhibition of FPC also reduces Pkd2-channel activities(P < 0.001; Figure 9, Bb versus Bc). For verification thatthe functional channel changes were induced by missing Pkd2,the anti-PC2 polyclonal antibody hPKD2-Cp, which recognizesthe intracellular COOH-terminal region of PC2, was transferredinto WT control IMCDsh15 cells via pipette solution. The currentdensity was significantly inhibited in IMCDsh15 cells that weretreated with hPKD2-Cp antibody (P < 0.02; Figure 9, Bb versusBe), suggesting that the current change is Pkd2 specific. Tovalidate our findings further, we produced primary culturesof renal epithelia from 2-mo-old WT and Pkhd1–/–kidneys. Whole-cell current densities were significantly decreasedin Pkhd1–/– cells compared with WT cells (P <0.003; Figure 9C), indicating that lack of FPC also inhibitsPkd2-specific channel activities. For clarification that thedownregulated current in Pkhd1–/– primary culturedcells was due to the Pkd2-specific channel, the whole-cell currentof WT primary cultured cells was recorded in the presence ofeither the anti-PC2 antibody hPKD2-Cp or an anti-actin antibody.As representative current records and I-V curves in Figure 9Dshow, the anti-PC2 antibody reduced the current density, butthe anti-actin antibody did not, confirming that the whole-cellcurrent was conducted through Pkd2 channels.
Figure 9. Analysis of whole-cell current recordings in Pkhd1-deficient cells. (A) Average current density (pA/pF) and voltage relation (I-V curve) of primary cultured fibroblast cells from E13.5 embryos show a statistically significant difference between WT and Pkd2–/– littermates, suggesting that the cation channel activity is induced by lack of PC2 (n = 10; *P < 0.005). (B) A Pkhd1-knockdown stable cell line IMCDshRNA3e23 and its WT–controlled cell line IMCDsh15 were used to perform the whole-cell current recording assays under the same conditions. Currents were elicited by stepping from a holding potential of 0 mV to various test potentials (Ba). The current densities between IMCDshRNA3e23 (n = 12) and IMCDsh15 (n = 7) cells exhibited a statistically significant difference (Bb versus Bc; *P < 0.001). The control IMCDsh15 cells displayed a steep inwardly rectifying current, which was significantly reduced by the intracellular introduction of the anti-PC2 C-terminal antibody hPKD2-Cp via pipette solution at a dilution of 1:200 (n = 8; Bb versus Be; #P < 0.02), suggesting that the current changes are PC2 specific. (C) Primary cultured renal epithelial cells derived from 2-mo-old WT (WT) and Pkhd1–/– littermates were used to perform the same whole-cell current recording assays. The control WT cells (n = 15) displayed a steep inwardly rectifying current, which was substantially reduced in Pkhd1–/– cells (n = 7). This indicates that the whole-cell current density in cells from the Pkhd1–/– mice is significantly lower than in those from the WT littermates (*P < 0.003). (D) hPKD2-Cp and a negative control anti-actin antibody were each added intracellularly into separate WT cells via pipette solution. Representative current traces were not clearly reduced in the cells with anti-actin antibody (n = 6), but a substantially smaller current amplitude was seen in the cells with hPKD2-Cp antibody (n = 8; #P < 0.01), further suggesting that the current alteration between WT and Pkhd2–/– cells in C is Pkd2 specific. (E) For unequivocal validation that the inhibited current density seen in the Pkhd1–/– cells is due to lack of FPC expression, a human full-length PKHD1 cDNA was in-frame constructed into a GFP-tagged expression vector and was transiently transfected into the Pkhd1–/– cells. GFP-positive cells were chosen to perform the whole-cell current recording (n = 4), and current density-voltage plots showed that the re-expression of FPC rescues the inhibited current density seen in the Pkhd1–/– cells (Pkhd1–/–-C versus Pkhd1–/–; #P < 0.01). The dashed line represents the mean I-V curve from the Pkhd1–/– cells in C, and the dotted line shows the means I-V curve from the WT cells in C. Currents were measured 220 ms after stepping to the test potential. Data are means ± SEM from the tested independent cells. Statistical difference at *+80 and #+100 mV between the tested cell groups. The data were fitted with standard Boltzmann equation (I = Imax/1 – e(V50 – V)/k) + C).
To obtain unequivocal evidence that the whole-cell current changeswere induced by downregulation of FPC, we further investigatedwhether re-expression of FPC could rescue the reduced channelactivities seen in Pkhd1–/– cells (Figure 9C). Wetransiently transfected primary cultured Pkhd1–/–cells with a mammalian expression vector, pcDNA3.1/Hygro, containingthe full-length ORF cDNA of human PKHD1 with GFP fused in-frame.GFP-positive Pkhd1–/– cells, named Pkhd1–/–-C,were chosen for the whole-cell current recording assay. As shownin Figure 9E, both the current density and I-V relation of Pkhd1–/–-Ccells were used to compare the currents recorded from WT (dottedline) and Pkhd1–/– cells (dashed line; Figure 9C).Unlike Pkhd1–/– cells, the current density of Pkhd1–/–-Ccells was similar to WT cells (Figure 9E), indicating that thecurrent changes between WT and Pkhd1–/– cells aredue to downregulation of FPC. That inhibition of Pkhd1 expressiondisrupts Pkd2-specific channel activities gives further evidenceto demonstrate FPC and PC2 function in the same molecular pathway.
Although the gene responsible for ARPKD, PKHD1, has been identified23–25and its gene product, FPC, has been initially characterized,27,28,30,31,39,40the mechanisms by which PKHD1 causes disease phenotypes remainlargely unknown. To study the disease mechanism and pathogenesisof ARPKD, we created a mouse that allows manipulation of Pkhd1,an animal model that recapitulates the human ARPKD phenotype.Through genetic and biochemical studies, we demonstrated thatthe COOH-terminus of FPC physically interacts with the NH2-terminusof PC2 and that lack of FPC leads to downregulation of PC2 expressionin vivo. Transmutant mice for Pkhd1 and Pkd2 displayed a significantlymore severe renal cystic phenotype than single-mutant mice,suggesting that Pkhd1 serves as a disease modifier for ADPKD.For functional validation of these findings, Pkd2-channel activitieswere examined both in primary cultures of renal epithelia derivedfrom Pkhd1 mutant mice and in Pkhd1-silenced IMCD cells.39Pkd2-channelactivities were significantly dysregulated in Pkhd1-deficientcells, indicating that FPC and PC2 reside in the same molecularpathway. During submission of our article, another group reportedthat there are genetic interactions between Pkhd1 and Pkd1.41Because the gene product of PKD1, PC1, was reported to interactwith PC2 and regulate Pkd2-channel activity,9–11 our findingscombined with theirs draw the conclusion that ADPKD and ARPKDmay reside in the same pathogenic pathway.
In this study, the Pkhd1–/– genotype seemed to causeembryonic lethality; however, a phenotypic analysis of thesePkhd1–/– embryos did not show any significant cardiacdefects and only mild edema, which was prevalent in the Pkd2mutant mice. Because Pkd2–/– mice are embryonicallylethal, the finding that a lack of FPC promotes a significantdownregulation of PC2 expression raises the possibility thatpartial embryonic lethality in Pkhd1–/– mice maybe caused by this reduction of PC2.
We previously reported that FPC and PC2 co-localize to the samesubcellular organelles, the basal bodies/primary cilia of renalepithelial cells,31 suggesting that these two cystoproteinsmay form a molecular complex. Recent reports have shown thatthe same chemical molecule (a vasopressin V2 receptor antagonist,OPC31260) can inhibit cyst progression in both a rat geneticmodel of ARPKD and a mouse genetic model of ADPKD, suggestingthat both causal gene products for ADPKD and ARPKD may residein a common molecular pathway.42 Recently, our company studydemonstrated that FPC and PC2 indirectly interact via theirCOOH-termini and that this is mediated by KIF3B, a motor subunitof the heterotrimer kinesin-2.36Pkd2-channel activities weresignificantly altered when the FPC–PC2 complex was disrupted.This in vitro study provides a molecular basis for a functionallink between FPC and PC2. Another recent report that supportsa functional link suggests that FPC regulates mechanotransducedCa2+ responses, which may be induced by PC2, in cultured Pkhd1-knockdowncells.43 In this study, the evidence that the COOH-terminusof FPC directly interacts with the NH2-terminus of PC2 suggeststhat FPC and PC2 are able physically to form a heterodimericcomplex in vivo. Lack of FPC downregulates Pkd2-channel activitiesin either the Pkhd1-knockout renal epithelial cells in primaryculture or the Pkhd1-knockdown IMCD cells that we generatedpreviously.39 Given that FPC and PC2 physically interact andthat the lack of FPC downregulates PC2 expression in vivo butPC2 does not downregulate FPC, we speculate that PC2 may functionimmediately downstream of FPC.
The disruption of ciliary formation in renal epithelia inducescystogenesis in the kidneys.34,35 We recently reported thatdownregulation of Pkhd1 significantly decreases ciliary formationin cultured Pkhd1-silenced IMCD cells, suggesting that lackof FPC might disrupt ciliogenesis in renal epithelial cells.This result is consistent with studies in which transient smallinterference RNA–mediated inhibition of Pkhd1 in cholangiocytesresulted in shortening and decreased formation of cilia,29 butspatial and environmental differences between in vivo tissuesand in vitro cell culture may lead to different results. Mousemodels with a deletion of Pkhd1 exon 4044 and gene-targetedmutations in Pkd2 that cause distinct liver and/or kidney cysts33,45,46do not exhibit defects in ciliary structure in the affectedepithelial cells, suggesting that the failure of renal epitheliato assemble primary cilia may not be the only factor leadingto cyst formation in the kidneys. For example, a recent studyshowed that disruption of the extracellular matrix protein laminin5, which is a major component of the tubular and glomerularbasement membranes, also produces cystic kidneys in Lama5 mutantmice.47
By carefully examining ciliogenesis in our Pkhd1 mutant kidneys,we found that primary cilia of the cortical tubular epitheliawere reduced in number and were shorter than controls, documentingthat disruption of FPC expression induces malformation of theprimary cilia in the kidneys in vivo. Recently, an elegant studydemonstrated that disruption of FPC expression causes defectiveplanar cell polarity in another Pkhd1 genetic model, the pckrat. Spindle orientation in the renal epithelial cells of thepck rat is aberrant, suggesting that cell polarity is disruptedduring mitosis of renal epithelial cells.48 Our previous invitro study also demonstrated that renal epithelial IMCD cellswith downregulated FPC exhibit aberrant migratory polarity andlose the ability to drive collective cell migration, suggestingthat the planar cell polarity also might be disrupted.39 Aberrantplanar cell polarity seen in cells with Pkhd1 defects suggeststhat ciliary defects in our Pkhd1 mutant mice might be causedby impeding orientally centriole arrangement and disabling theestablishment of epithelial polarity.39,49
Because our Pkhd1 mutant mice bear an in-frame GFP reporter,immunostaining with anti-GFP antibodies provided an FPC expressionprofile in affected tissues. Positive GFP expression was detectedin the apical domain of epithelial derivatives, including therenal, hepatic, pulmonary, and gastrointestinal epithelia aswell as ependymal cells lining the ventricles of the brain.These findings are consistent with our previous report31 andindicate that FPC may modulate the morphogenesis and maintenanceof tubular/ductal architectures in organs generated from theprimary duct system.50 Several recent studies indicated thatFPC is involved in notch-like processing and that its extremelylarge extracellular domain is released from the primary ciliaof renal epithelial cells.51 That GFP-positive signals weredetected at the microvilli of tubular epithelia in our Pkhd1–/–mice agrees with the report that FPC may be released into thetubular/ductal lumen.
In summary, we produced a mouse model for Pkhd1 that recapitulatesthe phenotypic characteristics of human ARPKD. Using this modelalong with a Pkd2 mutant mouse, we were able to demonstratethe importance of FPC expression for normal PC2 function. Thefinding of aberrant ciliogenesis in the kidneys of Pkhd1-deficientmice indicates that FPC disrupts the process of ciliogenesis.That FPC physically interacts with PC2 and that lack of FPCdestabilizes PC2 expression in vivo but not vice versa suggestthat PC2, also known as TRPP2, functions immediately downstreamof FPC. In addition, because inhibition of FPC expression reducesPkd2-channel activity, we conclude that a functional and molecularinteraction exists between FPC and PC2 in vivo. Extracellularbiochemical and/or physical signals may activate the receptor-likeprotein FPC, which then triggers TRPP2 channel to transmit signalsthat affect intracellular processes. Our in vivo study revealsan intriguing molecular relationship between FPC and PC2.
Mouse Strains
The gene-targeted mouse model for Pkd2 (Pkd2tm2Som) was previouslygenerated by us.32,33 To produce mutant mice for Pkhd1, we designeda targeting construct disrupting its 15th coding exon (Figure 1).We found 620 embryonic stem cell colonies resistant to G418,with one (W4A5) identified by PCR screening using a pair ofoutside-construct and cassette-based primers. This cell linewas further confirmed by Southern blot analysis and injectedinto C57Bl/6 blastocysts at the gene targeting and transgenicfacility of University of Connecticut Health Center.
Southern and Northern Blotting and Quantitative PCR
Southern analysis was used to genotype Pkhd1 and Pkd2 mutantmice with our published approaches.32,52 For Northern analysis,total RNA was isolated from embryos or kidneys using Trizolreagent (Invitrogen, Carlsbad, CA) following the manufacturer'sinstructions. Probes were labeled using the RadPrime DNA-labelingsystem (Invitrogen) with -32P-dCTP (PerkinElmer, Waltham, MA)and were hybridized with total RNA blots (25 µg/lane).Images of 28-S rRNA bands in these same blots were used as atotal RNA loading control.
Quantitative PCR was performed using the iCycler iQ Real-TimePCR Detection System with iQ SYBR Green Supermix kit (Bio-Rad,Hercules, CA). Two pairs of primers were designed from eachcDNA sequence of Pkhd1 and Pkd2 (Table 1).
Antibodies
Polyclonal antibodies and mAb against FPC (including hAR-Np,hAR-C2p, hAR-Nm3G12, and hAR-C2m3C10) and antibodies againsthuman PC2 (hPKD2-Cp and hPKD2-Cm1A11, formerly named PKD2A11)were described in our previous studies.31,36 In addition, otherpolyclonal antibodies and mAb were purchased: Anti-acetylated-tubulin, anti–-tubulin, anti–β-actin, anti-Flag,and anti-HA mAb (Sigma, St. Louis, MO); anti-GFP polyclonalantibodies (ab6556; Abcam, Cambridge, MA); fluorescein lotustetragonolobus lectin (Vector Laboratories, Burlingame, CA);anti–cytokeratin 7 polyclonal antibody (Santa Cruz Biotechnology,Santa Cruz, CA); and the nucleic acid dyes To-pro and Yo-pro(Molecular Probes, Eugene, OR).
Western Blotting and Immunoprecipitation
For Western analysis and immunoprecipitation, the detailed approacheswere similar to our previous publications.31,36,39 For performanceof co-IP and Western analysis, the entire intracellular terminiof FPC and PC2 were constructed into Flag-tagged and HA-taggedpCMV expression vectors53 and were named FPC-C-HA (amino acids3872 to 4074), PC2-N-HA (amino acids 1 to 221), and PC2-C-Flag(amino acids 682 to 968).
Histology, IF Staining, IHC, Confocal Microscopy, and scanning electron microscopy
To index cystic disease severity, we calculated the total numberof cysts using four histologic sections from each kidney. Detailedprocedures for histology, IF, and IHC were published previously.31For confocal microscopy, antibody-stained images were collectedas Z-series sections using a Zeiss LSM 510 confocal microscopesystem with x40, x63, and x100 oil objectives. For scanningelectron microscopy, mice were perfused as described previously.31The kidneys were removed, sectioned longitudinally, washed by1x PBS three times, and fixed in 2.5% paraformaldehyde. Afteran ethanol dehydration series, the samples were critical-pointdried and sputter-coated with 40% gold/60% palladium microparticlesto a thickness of 15 to 17 mm. Images of the samples were obtainedusing an Electroscan E3 Environmental Scanning Electron Microscope.
Cell Lines and Mouse Kidney Primary Epithelial Cultures
All cell lines used in this study were cultured under previouslydescribed conditions.39 To generate primary cultures of renalepithelia, we removed kidneys from 2-mo-old WT or Pkhd1 mutantmice and minced them finely with a scalpel. The minced tissuewas incubated with 0.5% collagenase type IV at 37°C for45 min and pipetted vigorously. The undigested tissue was removedby filtration through a 40-µ mesh filter. The remainingsingle cells and small organoids were washed three times withPBS containing 5 mM glucose. The cells were incubated with 10µg/ml biotinylated Dolichos biflorus agglutinin (Vector;B-1035) at 4°C for 60 min. Then the cells were washed againwith PBS before incubation with 50 µl of CELLectin Biotinbinder Dynabeads (Invitrogen) at 4°C for 30 min. BecauseDynabeads are superparamagnetic polystyrene beads, the incubatedmixtures were washed twice with PBS containing 5 mM glucoseusing a magnetic rack. The cells were eluted with release bufferand were plated on 24-well dishes with 10% FCS DMEM under 5%CO2 at 37°C.
Statistical Analyses
The survival rate was determined by observing the cohorts ofmice daily. We analyzed the survival using the Kaplan-Meierfunction in the R Software. Graphical data are presented asmeans ± SD. Statistical analysis was performed whereappropriate using the t test or one-way ANOVA followed by Tukeymultiple comparison test. Differences with P < 0.05 wereconsidered statistically significant.
This work was supported by grants from the National Institutesof Health (DK062373 and DK071090) to G.W.
We kindly thank Dr. Stefan Somlo for allowing us to use thePKD2 mutant model (Pkd2tm2Som) that was generated in his laboratory.We also thank Dr. James P. Smith for excellent advice and suggestions;Dr. Caiying Guo for the embryonic stem cell work at Universityof Connecticut; Dr. Chun Li for statistical analysis; and Drs.Sae-youll Cho, Shun-Wei Huang, Aijun Zuo, and Hong Wang fortechnical assistance.
Footnotes
Published online ahead of print. Publication date availableat www.jasn.org.
I.K. and Y.F. contributed equally to this work.
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Top
Abstract
Introduction
16 (Pkhd1–) (Figure 1, A through D). Because our gene-targeting construct included an in-frame green fluorescence protein (GFP) reporter gene with a predicted fusion protein of 130 kD (Figure 1A), anti-GFP immunoprecipitation assays showed that only the mutant mice expressed the GFP reporter protein (Figure 1E). In addition, with the use of a panel of previously generated anti-FPC antibodies,31 Western blot analyses were able to detect the expected immunoreactive bands at >400 kD in the wild-type (WT) and heterozygous mouse kidneys (Figure 1F). These immunoreactive bands were not detected in protein lysate from Pkhd1–/– mouse kidneys, suggesting that the homozygous mice lacked full-length, functional FPC. 
-tubulin antibody to examine the number and morphology of renal primary cilia in 6-mo-old littermates. Compared with WT mice, there were far fewer primary cilia in the renal tubular epithelia of Pkhd1–/– mice (Figure 6, A versus B), indicating that lack of FPC might reduce ciliogenesis in the kidneys. The ciliary defects seem to be most severe in the cortical proximal tubules of the Pkhd1–/– kidneys. In addition, confocal images also displayed similar ciliary changes in corresponding regions of littermate kidneys (Figure 6, C versus D). Scanning electron microscopy confirmed these results in 3-mo-old littermates with or without the targeted Pkhd1 mutation (Figure 6, E versus F). For further validation of these findings, primary culture of renal epithelial cells derived from 2-mo-old Pkhd1–/– and WT littermates was used to determine whether the ciliary malformation could be observed in vitro. Compared with the cultured WT cells, a shortened ciliary structure and decreased ciliary staining were seen in the Pkhd1–/– cells (Figure 6, G versus H). The cilia stained in approximately 94% of WT cells and in fewer than 34% of Pkhd1–/– cells (P < 0.001; Figure 6I). The mean length of primary cilia was 10 µm in cultured WT cells and was <5 µm in Pkhd1–/– littermate cells (P < 0.001; Figure 6J). That short or absent cilia were observed in Pkhd1–/– cells but not in WT cells suggests that lack of FPC causes defects in ciliogenesis in renal epithelial cells in vivo. 
-tubulin, anti–β-actin, anti-Flag, and anti-HA mAb (Sigma, St. Louis, MO); anti-GFP polyclonal antibodies (ab6556; Abcam, Cambridge, MA); fluorescein lotus tetragonolobus lectin (Vector Laboratories, Burlingame, CA); anti–cytokeratin 7 polyclonal antibody (Santa Cruz Biotechnology, Santa Cruz, CA); and the nucleic acid dyes To-pro and Yo-pro (Molecular Probes, Eugene, OR). 
