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Cell Biology |
* Department of Pediatrics and Adolescent Medicine and Renal Division, University Hospital Freiburg, Freiburg, Department of Pediatric Nephrology, Charite, Berlin, and || Deutsches Krebsforschungszentrum, Heidelberg, Germany; and Program in Molecular Medicine, University of Massachusetts Medical School, Worcester, Massachusetts
Address correspondence to: Dr. Heymut Omran, Department of Pediatrics and Adolescent Medicine, Mathildenstrasse 1, 79106 Freiburg, Germany. Phone: +49-761-270-4301; Fax: +49-761-270-4344; E-mail: heymut.omran{at}uniklinik-freiburg.de
Received for publication December 20, 2005. Accepted for publication June 26, 2006.
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Abstract |
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Introduction |
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NPHP type 1 (NPHP1; OMIM #256100) accounts for 62% of NPHP cases (1,11) and is one of the most frequent genetic causes of ESRD in children and young adults. In the majority (94%) of patients with NPHP1, large homozygous deletions of approximately 290 kb involving the NPHP1 locus (on chromosome 2q12-q13) can be detected, whereas only some patients carry point mutations in combination with a heterozygous deletion (3,4,11).
NPHP1 encodes nephrocystin, a 733–amino acid protein with an N-terminal coiled-coil domain, an adjacent Src homology 3 domain flanked by two highly acidic E-rich domains, and a conserved nephrocystin homology domain that encompasses the C-terminal two thirds of the protein (3,4). A number of protein interaction partners, including p130CAS, proline-rich tyrosine kinase 2, and tensin, that are supposed to function in focal adhesion complexes or at sites of cell–cell contact in polarized MDCK cells have been identified (12,13). In addition, the proteins that are involved in the NPHP2, NPHP3, and NPHP4 have been shown to associate with nephrocystin, suggesting assembly into a large, multiprotein complex (2,6,7,14).
Various cystic kidney disorders are associated with dysfunction of renal monocilia (15), which are localized on the epithelial surface of nephron segments, where they extend into the lumen of the kidney tubules and possibly act as fluid flow or chemosensors (16). Proteins that are involved in renal monocilia function include the ciliary proteins polycystin-1 and -2 and fibrocystin (mutated in autosomal dominant and autosomal recessive polycystic kidney disease, respectively) as well as the BBS1 through BBS8 proteins (mutated in Bardet-Biedl syndrome [BBS]), which localize to the basal bodies of cilia (16–22). The localization of the newly identified BBS9 through BBS11 proteins has not yet been investigated. Furthermore, nephrocystin (NPHP1), inversin (NPHP2), and nephrocystin-4 (NPHP4) localize to primary cilia predominantly at the ciliary base in renal epithelial cells (2,14).
Most of our knowledge about cilia structure and function originates from studies of the biflagellate unicellular alga Chlamydomonas rheinhardtii. The axonemal architecture of the motile Chlamydomonas flagella, composed of nine peripheral doublet microtubules that surround two central single microtubules, is highly reminiscent of motile respiratory cilia lining the upper and lower airways. Each flagellum and cilium extends from a specialized centriole, the basal body. The centriolar triplet microtubular structure converts within the transition zone into the axonemal doublet microtubular structure of the cilium. We recently showed that nephrocystin is present at the ciliary base of human respiratory cilia, where it co-localizes with the retinitis pigmentosa GTPase regulator (RPGR) and the phosphoacidic cluster sorting protein-1 (PACS-1) (23).
In this study, we analyze the dynamic expression of nephrocystin during ciliogenesis and evaluate its functional role. We show that nephrocystin is absent in undifferentiated respiratory cells, becomes cytoplasmically expressed during early phases of in vitro ciliogenesis, and is localized predominantly within the apical cytoplasmic area when the cell starts to polarize. With the onset of axoneme budding, nephrocystin becomes tightly localized at the ciliary base and exclusively localizes to the ciliary transition zone of mature respiratory epithelial cells. We further show that nephrocystin co-localizes with p130CAS and tensin in fully differentiated respiratory cells and thus possibly participates in a multimeric protein complex at the transition zone. In nephrocystin-deficient respiratory cells of patients with homozygous NPHP1 deletions, we analyzed the effect on ciliogenesis and motile cilia function. In addition, we demonstrate localization of nephrocystin within the photoreceptor connecting cilium, which is consistent with retinitis pigmentosa that is observed in a subset of patients with NPHP1 (24).
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Materials and Methods |
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Cell Culture
MDCK and HEK293T cells were cultured in DMEM/10% FCS, murine inner medullary collecting duct (mIMCD3) and LLC-PK1 (porcine kidney) were cultured as recommended by American Type Culture Collection (Manassas, VA). All cell culture reagents were from Life Technologies/Invitrogen (Karlsruhe, Germany). Cells were grown on coverslips for 8 d past confluence to allow for epithelial cell polarization and cilia formation, washed with PBS, and subjected to immunofluorescence staining as described below.
Immunoblotting
Protein extracts from Epstein-Barr virus–2-transformed B-lymphocytes, HEK293T, MDCK, and respiratory epithelial cells were prepared by standard procedures using NP-40 or RIPA lysis buffers. Axonemal high-salt protein extracts were obtained from a fresh pig trachea as described previously (25–27). Samples were separated on NuPAGE 4 to 12% bis-tris gels (Invitrogen) and blotted onto polyvinylidene difluoride membranes (Amersham). Blots were processed for ECL plus (Amersham/GE Healthcare, Freiburg, Germany) detection using rabbit anti-nephrocystin (1:2500) and anti-rabbit–horseradish peroxidase (1:2500) antibodies (Santa Cruz, Heidelberg, Germany).
Immunofluorescence Analysis
Respiratory epithelial cells were obtained by transnasal brush biopsy (Cytobrush Plus, Medscand, Malmö, Sweden) and suspended in RPMI 1640 medium without supplements. Cells were spread onto glass slides, air-dried, and stored at –80°C until use. A pig eye was obtained from a local butchery, and the retina was removed carefully using a scalpel. Cryosections (10 µm) were prepared according to standard methods. Samples were treated with 4% paraformaldehyde, 0.2% Triton-X 100, and 5% skim milk (all in PBS) before incubation with primary (at least 2 h) and secondary (30 min) antibodies at room temperature. Slides were washed with PBS after each step. Appropriate controls were performed omitting the primary antibodies. Antibodies were mouse anti–acetylated-
-tubulin and mouse anti–
-tubulin (Sigma, Taufkirchen, Germany), mouse anti–
-tubulin (Abcam, Cambridge, UK), mouse anti–PACS-1, mouse anti-tensin, and mouse anti-p130cas (Transduction Laboratories, BD Biosciences, Heidelberg, Germany). Polyclonal rabbit antibodies against DNAH5, nephrocystin, IFT88, and IFT20 as well as mouse anti-nephrocystin antibodies have been described previously (13,23,27–29). Secondary antibodies (Alexa Fluor 488, Alexa Fluor 546) were from Molecular Probes (Invitrogen). DNA was stained with Hoechst 33342 (Sigma). Confocal images were taken on a Zeiss laser scanning microscope (Axiovert 200 LSM510 META) using a 63 x 1.2 numerical aperture water immersion or a 100 x 1.3 numerical aperture oil immersion objective. A four-channel, eight-bit multitracking scan mode was used with a 1024 x 1024 frame size and four-fold average line scan settings. Images were processed with the Zeiss LSM510 software.
High-Speed Video Analysis for Ciliary Beat Assessment
Ciliary beat frequency was assessed with the SAVA system (30). Transnasal brush biopsies were viewed immediately with an Olympus IMT-2 microscope (x40 phase contrast objective) equipped with a Redlake ES-310Turbo monochrome high-speed video camera (Redlake, San Diego, CA) set at 125 frames per second. The ciliary beating pattern was evaluated on slow-motion playbacks.
Electron Microscopy
A mouse eye was cut into slices using a scalpel. Ultrathin retina sections were prepared according to standard methods and subjected to transmission electron microscopy using a Zeiss EM 900.
Ciliogenesis
Respiratory epithelial cells from nasal conchae or polyps were obtained from patients who underwent ear, nose, and throat surgery or after nasal brushing biopsy. Primary cell culture was performed essentially as described previously (31). Briefly, cells were isolated from tissue samples with pronase (Sigma) and grown to confluent monolayers on collagen-coated tissue flasks in F12/DMEM/2% Ultroser G (Pall Life Sciences, Cergy-Saint-Christophe, France). Cell layers were treated with collagenase, cut into pieces, and cultured in Ham's F12/DMEM/10% NU-serum (BD Biosciences) on a rotary shaker. After 10 d, most of the cells were organized in spheroids covered with motile cilia.
Ca2+-Dependent Deciliation
Respiratory epithelial cells from brush biopsies were collected by centrifugation (300 x g, 5 min) and resuspended in deciliation buffer that contained 15.7 mM Tris-Cl (pH 7.5), 79.1 mM NaCl, 1.56 mM EDTA, 0.1% Triton-X 100, 15.8 mM CaCl2, and protease inhibitors (32). Deciliation (30 to 60 min at room temperature with occasional shaking) was monitored under a microscope. Aliquots were removed, cells were pelleted, and cilia were collected from the supernatant by centrifugation (16,000 x g, 5 min). Samples were spread onto glass slides, air-dried, and used for immunofluorescence staining as described above.
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Results |
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-tubulin, a marker of the microtubule organizing centers (MTOC), which are located adjacent to the basal bodies, demonstrates that nephrocystin localizes in renal monocilia of mIMCD-3 cells distal to the MTOC within the transition zone (Figure 1B).
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On the basis of interaction with tensin and p130cas, it has been speculated that nephrocystin functions at focal adhesions (12,13). In contrast, we find that in human ciliated respiratory epithelial cells, tensin and p130cas localize at the apical cytoplasmic area, predominantly at the MTOC region, where their signals overlap at the proximal end of the transition zone with nephrocystin (Figure 1, E and F). Because the transition zone is thought to act as the docking site for the intraflagellar transport (IFT) particles, we next tested whether nephrocystin co-localizes with the IFT proteins IFT88 and IFT20. Similarly, as in Chlamydomonas (33), both IFT proteins are localized predominantly at the ciliary base and with a speckled pattern along the ciliary axonemes in respiratory epithelial cells (Figure 1, G and H). Overlapping staining of nephrocystin with IFT proteins is observed only at the proximal part of the transition zone but not within the ciliary axoneme. In addition, antibodies directed against -tubulin stained the entire axonemes. At the proximal axonemal end, -tubulin localizes in close vicinity of nephrocystin (Figure 1I).
Because retinitis pigmentosa can be found in patients with NPHP1, we tested whether nephrocystin localizes to the connecting cilium that bridges the outer and inner photoreceptor segments (Figure 2A). Co-staining of retinal cryosections with antibodies against nephrocystin and either acetylated -tubulin or -tubulin demonstrates that nephrocystin exclusively localizes to the connecting cilium in close proximity to the basal bodies (Figure 2, B and C).
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At an early stage of in vitro ciliogenesis, cell morphology of spheroids is characterized by a symmetric cell body and a central nucleus in most spheroids (data not shown). These cells do not express nephrocystin and do not carry cilia as evidenced by the absence of acetylated -tubulin staining. A few cells within each spheroid then start to express nephrocystin with a diffuse cytoplasmic localization (Figure 4, A and B), which precedes cilia formation (4 to 8 d). In the next period of respiratory cell differentiation (6 to 8 d), most cells in each spheroid show diffuse cytoplasmic nephrocystin expression (Figure 4B) that becomes apically enhanced in those cells that exhibit cell polarization, recognizable by a more prolonged cell body and downward placement of the nucleus. The concentration of nephrocystin at the apical cell region coincides with expression of acetylated -tubulin that specifically localizes to well-defined spots beneath the apical plasma membrane in these cells, indicating the beginning of axoneme budding and cilia formation (Figure 4, B and C). During later stages of ciliogenesis (8 to 12 d), the punctual localization of nephrocystin at the base of each growing and mature axoneme and proximally to the acetylated -tubulin remains unchanged (Figure 4, D and E). Staining of nephrocystin at the basolateral side of respiratory cells never was observed.
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To test whether nephrocystin localization also is altered in patients with other forms of NPHP, we analyzed respiratory epithelial cells from two patients who had adolescent NPHP and carried compound heterozygous NPHP3 mutations (patients F23II1 and F23II2) and one patient who had NPHP type 5 and carried compound heterozygous NPHP5 mutations (patient A19), which were reported previously (7,8). In these patients, cell morphology and nephrocystin localization were normal (data not shown), demonstrating that NPHP3 and NPHP5 mutations in the analyzed patients do not alter nephrocystin localization. In addition, we found normal nephrocystin localization in two patients with BBS, five patients with autosomal recessive polycystic kidney disease, and three patients with NPHP (for which NPHP1 deletions have been excluded) and unknown mutations (data not shown). These results demonstrate that absence of nephrocystin from the transition zone is a specific finding in patients who have NPHP with NPHP1 deletions rather than a common alteration in cystic kidney diseases.
Next, we asked whether nephrocystin is essential for correct targeting of ciliary/basal body proteins. However, staining of nephrocystin-deficient respiratory cells with antibodies against PACS-1 (Figure 5, C and D); -tubulin, a component of the pericentriolar material around the basal bodies; and IFT88 (Figure 5, E and F) revealed normal localization of all three proteins at the ciliary base. These results demonstrate that nephrocystin is not required for targeting of these proteins to the ciliary base and that the structural integrity of the ciliary base remains unaffected in nephrocystin-deficient cells. Furthermore, because the localization IFT88 along the ciliary axonemes also remains unaffected in nephrocystin-deficient cells, intraflagellar transport per se obviously is not impaired by the absence of nephrocystin from the transition zone.
We also tested whether the loss of nephrocystin impairs the function of respiratory cilia because three of the patients with NPHP1 deletions (ON-21, ON-23, and ON-50) reported mild respiratory symptoms with chronic sinusitis and rhinitis suggestive of a cilia dysmotility defect. High-speed video-microscopic analyses of respiratory epithelial cells from these patients showed normal ciliary beat frequencies (5 to 9 Hz at room temperature) and normal beat amplitudes. However, evaluation of slow-motion playbacks revealed that ciliary motility is slightly irregular (Supplementary Videos 1 and 2).
To examine whether the observed dysmotility is caused by secondary effects, we grew respiratory epithelial cells that were obtained by transnasal brush biopsies from patient ON-50 to heavily ciliated spheroids in vitro, which bypasses secondary ciliary dyskinesia (35). Nephrocystin-deficient spheroids (Figure 5G) had normal morphology and normal ciliogenesis when compared with control cells (Figure 4) as well as normal ciliary beat frequencies and amplitudes. Although we noted a slightly irregular beating pattern, the degree of the observed abnormality was not as severe as usually observed in primary ciliary dyskinesia (Supplementary Videos 3 and 4).
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Discussion |
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In Caenorhabditis elegans, the homologs of nephrocystin and nephrocystin-4 also localize to the cilia transition zone of sensory neurons but were not detected within the ciliary axoneme (37), which indicates evolutionary conservation of subcellular localization from C. elegans to human. In addition, we demonstrate for the first time that nephrocystin localizes to the photoreceptor-connecting cilium (Figure 2), which might explain why a subset of patients with NPHP1 exhibit retinal degeneration (24). It is interesting that the photoreceptor-connecting cilium is the analogous structure of the ciliary transition zone, where the interaction partners retinitis pigmentosa GTPase regulator (RPGR) and NPHP5 also are localized (8,38). In addition, NPHP4 and NPHP6 have been localized to the ciliary base (9,14,39). This indicates that nephrocystin possibly participates in the function of these proteins at the ciliary base.
To increase our understanding of nephrocystin function, we analyzed primary human respiratory cells, which are readily accessible, carry multiple instead of single cilia on their surface, and allow in vitro ciliogenesis. Furthermore, nephrocystin-deficient cells from patients with juvenile NPHP can be obtained, obviating analyses of genetically manipulated cell systems.
Because nephrocystin physically interacts with a number of proteins, including p130cas, tensin, PACS-1, -tubulin, and nephrocystin-2, -3, and -4, previous data suggested that these proteins form a complex at cell junctions, at focal adhesions, or within the ciliary axoneme (2,12,13,19). Our data indicate, however, that these proteins probably assemble into a functional protein complex at the ciliary base, which is supported by the subcellular localization of p130cas, tensin, -tubulin (Figure 1, E, F, and I) and PACS-1 (Figure 5, C and D) with overlapping staining patterns or localization in close proximity to nephrocystin in respiratory epithelial cells.
We show that nephrocystin localization extends proximally and distally to the ciliary autotomy site (Figure 3), within the transition zone where Ca2+-dependent microtubule severing occurs (34), where it is tightly attached to the axonemal structure. This robust, detergent-resistant attachment possibly is mediated by interaction with the axonemal structural component -tubulin (2) and indicates that nephrocystin is neither a component of the ciliary membrane nor a transiently bound molecule at the ciliary base. During in vitro ciliogenesis (Figure 4), we found that nephrocystin is not detectable in undifferentiated respiratory cells but is expressed with a diffuse cytoplasmic localization as soon as establishment of cellular polarity becomes evident. Simultaneously with axoneme budding and the appearance of respiratory cilia, nephrocystin completely translocalizes to the ciliary bases. Although reminiscent of a possible role for establishment or maintenance of cell polarity, we did not find evidence for obviously disrupted respiratory epithelial cell polarization in nephrocystin-deficient cells (Figure 5). In contrast, such a role was demonstrated previously for BBS proteins and inversin (NPHP2), which are involved in the pathogenesis of BBS and infantile NPHP, respectively (40,41).
In nephrocystin-deficient respiratory cells from patients with homozygous NPHP1 deletions (Figure 5), expression and localization of PACS-1, a binding partner of nephrocystin, was normal, indicating that nephrocystin is not essential for the localization of PACS-1 to the ciliary base. In addition, because the localization of -tubulin and IFT88 at the ciliary base and along the ciliary axoneme, respectively, is normal in nephrocystin-deficient cells, the structural integrity of the cilia and the intraflagellar transport per se obviously is unaffected by the absence of nephrocystin from the transition zone. Furthermore, in vitro ciliogenesis of nephrocystin-deficient respiratory cells did not reveal any obvious abnormalities, indicating that nephrocystin is essential neither for establishment of cell polarity nor for cilia formation in respiratory epithelial cells. Similar results have been obtained in C. elegans, where nphp-1 and nphp-4 mutants did not exhibit abnormalities of cilia morphology in sensory neuronal cells (37).
Defects of the intraflagellar transport cause shortened flagella in Chlamydomonas and shortened renal monocilia and cystic kidney disease in tg737orpk mice (15,42). The observation of normal cilia morphology in nephrocystin-deficient cells argues against a function of nephrocystin as an intraflagellar transport protein that is essential for transport of ciliary proteins across the compartment border and along the axonemes (43). In addition, the subcellular localization of the intraflagellar transport proteins IFT88 and IFT20, typically within the MTOC and with a punctuate pattern along the ciliary axonemes, differs from the localization of nephrocystin, which is confined to the transition zone (Figure 1). It is interesting that IFT88 and IFT20 localize in close proximity to nephrocystin with a narrow overlapping localization at the proximal end of the transition zone. Thus, nephrocystin exactly localizes to the ciliary substructure, where protein transfer to and from the ciliary compartment occurs. Nephrocystin might be a component of a supramolecular structure called the ciliary (flagellar) pore complex, which is marked by transition fibers in Chlamydomonas (43).
Three studied patients with homozygous NPHP1 deletions reported symptoms that also occur in primary ciliary dyskinesia (44,45), suggesting a possible cilia motility defect. Although the beating patterns of respiratory cilia were slightly irregular in these patients, we do not consider the observed findings to be diagnostic for primary ciliary dyskinesia, in absence of other typical symptoms such as bronchiectasis.
Our finding that nephrocystin deficiency can be readily identified by immunofluorescence microscopy in nasal respiratory cells that are obtained by noninvasive brush biopsy might contribute to the development of novel diagnostic tools in cystic kidney disorders. This suggests that DNA diagnosis by molecular genetic approaches may be directed depending on a characteristic aberrant staining pattern.
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Acknowledgments |
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We are grateful to the patients for participation in this study and for the support of the Arbeitsgemeinschaft fuer Paediatrische Nephrologie. We thank Roland Nitschke and Sabine Haxelmans, Life Imaging Center, Institute for Biology I, University Freiburg, for excellent support with confocal imaging and Julia Kalnitski, Karin Sutter, Carmen Kopp, and Myriam Krome for technical assistance.
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