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Primary ciliary dyskinesia (PCD) is a genetically heterogeneous autosomal recessive disorder characterized by recurrent infections of the respiratory tract associated with abnormal function of motile cilia. Approximately half of PCD patients also have alterations in the left-right organization of internal organ positioning including situs inversus and situs ambiguous (Kartagener’s Syndrome, KS). Here we identify an uncharacterized coiled-coil domain containing protein (CCDC40) essential for correct left-right patterning in mouse, zebrafish and humans. Ccdc40 is expressed in tissues that contain motile cilia and mutation of Ccdc40 results in cilia with reduced ranges of motility. Importantly, we demonstrate that CCDC40 deficiency causes a novel PCD variant characterized by misplacement of central pair microtubules and defective axonemal assembly of inner dynein arms (IDAs) and dynein regulator complexes (DRCs). CCDC40 localizes to motile cilia and the apical cytoplasm and is responsible for axonemal recruitment of CCDC39, which is also mutated in a similar PCD variant.
Underlying defects in cilia ultrastructure are responsible for altered ciliary beat in PCD patients. The core structure of the cilium is the axoneme: nine peripheral microtubule doublets with or without a central pair of microtubles (9+2 or 9+0), interconnected by outer and inner dynein arms (ODAs and IDAs), radial spokes, nexin links and a central sheath 1. Coordinated activation of the ODAs and IDAs generates the ciliary beat. Most of the characterized PCD variants exhibit mutations in genes that encode dynein arm components such as DNAI1, DNAI2, DNAH5, DNAH11, and TXNDC3 2. Mutations in genes encoding cytoplasmic proteins such as C14orf104 (KTU) and LRRC50 also affect assembly of dynein arm complexes in the cytoplasm in a poorly understood process 3–6.
In our forward genetic screens to identify genes required for normal development of the mouse embryo 7–8, we isolated a mutant which exhibits left-right patterning defects. Greater than one-third of homozygous links (lnks) mutant embryos (39%, n=172) display laterality defects at E11.5–15.5 (Fig. 1a–d) including situs inversus (8%) or left isomerism (19%) based on lung lobation patterns. The majority of homozygous lnks mutant pups die before weaning due to unknown causes. In two homozygous lnks mutant pups that were examined, no kidney cysts were detected but hydrocephalus was noted (Supplementary Fig. 1). These observations resemble findings obtained in Mdnah5 deficient mice, a mouse model for PCD where ependymal cilia motility is important to prevent hydrocephalus 9. The lnks mutation was mapped to a 0.3 MB region of mouse chromosome 11 (Fig. 1e) that included the uncharacterized Coiled-coil domain containing 40 (Ccdc40) gene. Coiled-coil domains typically function in homodimerization and are present in a number of proteins involved in intracellular transport 10. Ccdc40 is specifically expressed in the embryonic node and midline tissues (Fig. 1f–i), key tissues that control left-right patterning. Upon sequencing the 3378 base pair Ccdc40 transcript from lnks mutant mice, a C to A transversion was identified (Fig. 1j). This nonsense mutation converts Valine792 to a stop codon in the middle of the coiled-coil domain, truncating the predicted 1125 amino acid protein (Fig.1j,k).
In zebrafish embryos, ccdc40 is expressed in tissues that contain motile cilia including Kupffer’s vesicle, floorplate, pronephric tubules and otic vesicle (Fig. 2; and data not shown). To explore the evolutionary conserved role of ccdc40 in left-right patterning, we designed two different antisense morpholino oligonucleotides (MOs) against zebrafish ccdc40. Both MOs disrupt splicing of the ccdc40 transcript (Supplementary Fig. 2) and produce similar phenotypes upon injection (Fig. 2e,g,i). Injection of MO resulted in a curly-tail down phenotype characteristic of other zebrafish mutants with laterality defects. Uninjected control embryos exhibited predominantly situs solitus (SS) at the 48 hpf stage with normal rightward looping of the heart, liver on the left and pancreas on the right side of the midline. By contrast, injection of either MO resulted in laterality defects: either reversed organ patterning, situs inversus (SI; 15–24%), or randomized organ patterning, heterotaxia (HTX; 13–19%). Both laterality and curly-tail down phenotypes could be rescued by co-injection of ccdc40 mRNA (Fig. 2h,j).
ccdc40 maps to zebrafish chromosome 6 in a region associated with the zebrafish mutant locke (lok), previously described as having a strong curly-tail down phenotype, laterality defects and pronephric cysts, without defects in sensory cilia or presence of hydrocephalus 11–12 (and J. S-B and R.D.B. unpublished). The locke phenotype is indistinguishable from knockdown of Ccdc40 in zebrafish (Fig. 2f,g,i). We sequenced genomic DNA from lokto237b mutants and found a C to T transition within the 3370 base pair transcript that changes Glutamine778 to a stop codon (Fig. 2d).
The laterality defects observed in mouse zebrafish mutants combined with expression of the transcript in the node/Kupffer’s vesicle suggest that Ccdc40 may act to regulate cilia function (Fig. 3). Indeed, scanning electron microscopy (SEM) revealed that the length of the cilia projecting from the nodal pit cells in lnks mutants is drastically reduced (Fig. 3a,b,e,f). Similarly, cilia were shorter in Kupffer’s vesicle and the pronephric tubules of ccdc40 morphants compared to uninjected controls (Fig. 3c,d,g,h). These results indicate that Ccdc40 is required for proper formation or maintenance of cilia.
Based on the cilia and laterality phenotypes in mouse and zebrafish ccdc40 mutants, we considered CCDC40, a strong candidate gene for human PCD. All coding CCDC40 exons and the adjacent intron-exon boundaries were amplified by PCR in a cohort of 26 PCD patients displaying a similar axonemal defect (see below). Sequence analyses revealed CCDC40 loss-of-function mutations in 17 PCD patients (Supplementary Fig. 3 and Table 1). Segregation analyses in all PCD families with CCDC40 mutations were consistent with autosomal recessive inheritance (Supplementary Fig. 4). Furthermore, in 15 affected individuals originating from 13 families, sequence analyses identified mutations on both CCDC40 alleles. However, in two families a mutation on the second allele was not identified by this approach. We addressed whether large deletions involving CCDC40 might be present on the other allele in these patients. Indeed segregation analysis of single nucleotide polymorphisms (SNPs) identified by sequence analysis of PCR products provided evidence for parental non-contribution suggestive of heterozygous CCDC40 deletion in family OP-43. SNP segregation was consistent with the interpretation that the three affected individuals inherited a large genomic deletion involving at least exon 1 from the mother and the point mutation (c.C1366T; p.R449X) from the father (Supplementary Fig. 3 and 5). In the affected individual of the one remaining family that carried point mutations solely on a single allele, we might have missed larger genomic mutations due to limitations of SNP analyses. Alternatively, mutations may reside in the non-coding regulatory or intronic regions.
The clinical phenotype of PCD patients harboring CCDC40 mutations is consistent with a severe defect of cilia beating, because patients suffered from recurrent upper and lower airway infections. To examine this directly, high-speed videomicroscopy analyses of respiratory cilia obtained by nasal brushing biopsies revealed a severely altered beating pattern in all analyzed samples. Respiratory cilia from affected patients exhibited markedly reduced beating amplitudes and the cilia appeared rigid with fast flickery movements (Supplementary Fig. 6; Supplementary videos 1–4). These motility defects are similar to those reported for pronephric cilia in lok mutants 11 and those observed in ccdc40 morphants (data not shown). No significant difference between cilia length was found in analyses of respiratory cilia from seven PCD patients carrying recessive CCDC40 loss of function mutations compared with normal controls (Supplementary Fig. 7), implying that ciliary movement can be disrupted in the absence of gross structural defects.
Consistent with a conserved functional role of CCDC40 for nodal cilia function, five patients displayed situs solitus (32%) and 11 patients situs inversus (68%). Together, these findings provide compelling evidence that recessive loss-of-function mutations within CCDC40 are responsible for a novel PCD variant characterized by altered mucociliary clearance of the airways and randomization of left/right body asymmetry.
We hypothesize that CCDC40 affects axonemal assembly of protein complexes leading to abnormal cilia morphology and/or motility. Axonemal structure was examined by Transmission electron microscopy (TEM) of cilia in zebrafish embryos and human cells. Motile cilia display a typical 9+2 microtubule configuration whereas lok mutants showed misplaced and/or duplicated central tubules and misplaced peripheral doublets (Fig. 3i–l; see similar axonemal defects in 12). Intriguingly, outer dynein arm morphology appeared normal. Similarly, TEM analyses of CCDC40-mutant respiratory cilia from PCD patients revealed defects in several axonemal structures including occasional absent or eccentric central pairs, displacement of outer doublets, reductions in the mean number of inner dynein arms and abnormal radial spokes and nexin links (Fig. 4) yet outer dynein arms appeared normal. Interestingly, in a parallel work Merveille et al. 13 show that recessive CCDC39 mutations cause a PCD variant indistinguishable from that caused by CCDC40 mutations.
To characterize further the molecular defect in CCDC40 mutant respiratory cells, we performed high-resolution immunofluorescence analyses on control and CCDC40 patient samples. In confirmation of the TEM analysis, we found normal composure of axonemal outer dynein arm motor proteins DNAH5, DNAH9 and DNAI2 (data only shown for DNAH5, Fig. 4a). Moreover, we confirmed an absence of the IDA component DNALI1 (Fig. 4c) from respiratory ciliary axonemes in all analyzed samples of affected patients. In most mutant respiratory cells DNALI1 accumulated in the apical cytoplasm (Fig. 4c). The severely reduced beating amplitude of respiratory cilia prompted us to investigate whether the axonemal assembly of the dynein regulatory complex (DRC) is also affected by CCDC40 deficiency. Thus, we examined expression of the mammalian DRC protein GAS11 (orthologous to Chlamydomonas DRC protein PF2 14) in all affected patients carrying CCDC40 mutations (see Table 1). Control respiratory cells showed strong GAS11 localization throughout all ciliary axonemes; however, in CCDC40 mutant respiratory cells, GAS11 was undetectable in ciliary axonemes (Fig. 4b). Similar to DNALI1, GAS11 accumulated in the apical cytoplasm of most mutant respiratory cells (Fig. 4b). Thus, we provide evidence that CCDC40 is necessary for correct assembly of at least two distinct axonemal complexes regulating ciliary beat: DNALI1-containing IDAs and GAS11-containing DRC. Furthermore, based on TEM findings, radial spokes are also altered in CCDC40 deficient respiratory cilia.
We generated polyclonal antibodies to determine the intracellular localization of Ccdc40 in sections of the mouse node (Supplementary Fig. 7). Wildtype embryos at E8.0 showed a punctuate pattern of Ccdc40 localization throughout the cytoplasm of node cells with significant overlap of expression with tubulin in the apical cytoplasmic regions of nodal cells (Fig. 3m–o). Ccdc40 antibody staining in E8.0 lnks mutant embryos confirmed antibody specificity and showed that truncation of the coiled-coil domain of Ccdc40 results in markedly decreased antibody staining in the node of lnks mutant embryos (Fig. 3p–r). Interestingly, we did not observe Ccdc40 protein localized to the 9+0 cilium in the mouse node (white arrow Fig. 3o); however, we do see axonemal localization of Ccdc40 in 9+2 respiratory (tracheal) cells (white arrow Fig. 3t), which is lost in lnks-mutant respiratory cells (3w). Ccdc40 may be at too low a level in the node cilium to detect in this assay, or this may reflect a difference in localization between monociliated 9+0 and 9+2 multiciliated cells. Together, these results suggest that Ccdc40 is required for cilia function by acting in the cytoplasm and possibly in the cilium itself. Because CCDC39 mutations cause a remarkably similar PCD phenotype 13, we analyzed whether CCDC40 deficiency affects axonemal localization of CCDC39. Interestingly, in all analyzed CCDC40-mutant respiratory cells, CCDC39 is absent from the cilium and is instead enriched in the apical cytoplasm at the ciliary base (Fig. 5). Thus, CCDC40 appears to be responsible for axonemal recruitment of CCDC39.
Our findings suggest that CCDC40 may physically interact with the other axonemal components and serve as a part of the axoneme structural scaffold, possibly as a new DRC component. This conclusion is consistent with findings that mutations in genes encoding DRC components in Chlamydomonas cause a similar ultrastructural phenotype in flagella including IDA defects15–17. Alternatively, it is possible that CCDC40 is important for cytoplasmic pre-assembly, axonemal targeting, and/or transport of the axonemal components CCDC39, GAS11 and DNALI1. Mutations in genes responsible for cytoplasmic pre-assembly and/or and axonemal targeting of DNALI1-containing IDA complexes, such as KTU and LRRC50, have thus far only been reported when ODA complexes are also affected 4,5. Nothing is yet known of the process of cytoplasmic pre-assembly and axonemal targeting/delivery of DRC complexes. Based on our functional data we propose that CCDC40 belongs to a group of novel evolutionarily conserved coiled-coil domain-containing proteins (including CCDC39) that govern the assembly of DRC and IDA complexes responsible for cilia beat regulation but not ODA complexes. Identification and molecular characterization of this process greatly aids diagnosis of PCD and will help direct research for novel therapeutics.
Signed and informed consent was obtained from patients fulfilling diagnostic criteria of PCD 19 and family members using protocols approved by the Institutional Ethics Review Board at the University of Freiburg and collaborating institutions. We studied DNA from a total of 26 PCD patients originating from 24 unrelated families. These patients exhibited axonemal defects documented ether by electron microscopy analyses and/or with immunofluorescence analyses as described in 13. 22 patients without evidence of CCDC39 mutations as well as four new patients displaying the same phenotype were analyzed for presence of CCDC40 mutations.
Nasal brush biopsies were taken from the middle turbinate and fixed in 2.5% glutaraldehyde in 0.1M sodium cacodylate buffer at 4°C, washed overnight and postfixed in 1% osmium tetroxide. After dehydration, samples were embedded in a propylene oxide / epoxy resin mixture. After polymerisation several resin sections were cut using an ultra-microtome. The sections were picked up onto copper grids and stained with Reynold's lead citrate. Transmission electron microscopy was performed with a Philips CM10.
Respiratory epithelial cells were obtained by nasal brush biopsy (Engelbrecht Medicine and Laboratory Technology, Germany) and suspended in cell culture medium. Samples were spread onto glass slides, air dried and stored at −80°C until use. Cells were treated with 4% paraformaldehyde, 0.2% Triton-X 100 and 1% skim milk prior to incubation with primary (at least 3 hours at room temperature or over night at 4°C) and secondary (30 minutes at room temperature) antibodies. Appropriate controls were performed omitting the primary antibodies. Monoclonal anti-DNALI1 antibodies, monoclonal anti-Gas11 antibodies and polyclonal anti-DNAH5 were reported previously 5,20 (and parallel work Merveille 13). Polyclonal rabbit anti-α/β-tubulin was from Cell Signaling Technology (USA); monoclonal mouse anti-acetylated-α-tubulin antibody and polyclonal CCDC39 antibodies were from Sigma (Germany). Highly cross adsorbed secondary antibodies (Alexa Fluor 488, Alexa Fluor 546) were obtained from Molecular Probes (Invitrogen). DNA was stained with Hoechst 33342 (Sigma). Confocal images were taken on a Zeiss LSM 510 i-UV.
Ciliary beat was assessed with the SAVA system 21. Nasal brush biopsies were rinsed in cell culture medium and immediately viewed with an Olympus IMT-2 inverted phase-contrast microscope equipped with a Redlake ES-310 Turbo monochrome high-speed video camera (Redlake, San Diego, USA) and a 40× objective. Digital image sampling was performed at 125 frames per second and 640×480 pixel resolution. The ciliary beat pattern was evaluated on slow motion playbacks.
The lnks mouse line was identified in a screen for recessive ENU-induced mutations that cause laterality defects at E11.5 and E12.5. The lnks mutation was generated on a C57BL/6J genetic background and backcrossed to C3H. In a mapping cross of 124 opportunities for recombination, the lnks mutation was mapped between the Massachusetts Institute of Technology (MIT) simple sequence length polymorphism (SSLP) markers D11mit48 and D11mit104. For high-resolution mapping, additional polymorphic DNA markers were generated based on nucleotide repeat sequences and include: D11ski2, D11ski10 and D11ski16 (see http://mouse.ski.mskcc.org/ for sequence of primers). The entire lnks transcript was sequenced by RT-PCR (Superscript One-Step RT-PCR, Invitrogen) using RNA isolated from lnks/lnks and C57BL/6 control embryos.
Whole-mount and section RNA in situs in mouse were preformed as described 22–23. The expression pattern of Ccdc40 was determined using an anti-sense RNA probe synthesized from IMAGE clones: 1362516 and 5702143 with identical results. The affinity purified polyclonal anti-Ccdc40 antibody was generated by immunization of rabbits using the peptide AYPPKKAKHRKVRPQAEV (Bio-Synthesis Inc). Antibody specificity was initially tested by immunoblotting. Briefly, protein extracts prepared from wildtype and lnks mutant mouse respiratory epithelial cells were resolved on a NuPAGE 4–12% bis-tris gel (Invitrogen, Karlsruhe, Germany) and blotted onto a PVDF membrane (Amersham). The blot was processed for ECL plus (GE Healthcare, UK) detection using anti-Ccdc40 (1:100) and anti-rabbit-HRP (1:3000) antibodies (GE Healthcare, UK). Immunofluorescence experiments were performed as described 24 using anti-acetylated tubulin (Sigma; 1:1000) and Hoechst (Sigma; 10 µg/ml). For analysis of Ccdc40 expression in the mouse node, in addition to examination of staining in 20µm sections, protein expression was examined in whole mount of 4 wildtype embryos and the Ccdc40 protein was absent in all 56 nodal cilia examined.
Morpholino antisense oligonucleotides targeting ccdc40 (ccdc40MO) were purchased from GENE Tools, LLC (Philomath, OR, USA). MO1 was designed against the splice-donor sites of exon 12 and intron 12 of ccdc40; 5‘-TGTTAACTGTGGTACATACTC TCTC-3’ (e12i12). MO2 was designed against the splice-acceptor sites of intron 10 and exon 11 of ccdc40; 5’-GCCTCCTGAAAAATCAAATATACAC-3’ (i10E11). Both MOs were conjugated to fluorescein isothiocyanate (FITC). 3ng – 6ng of ccdc40MO per 500 pl were injected per embryo and both MOs produced similar results. For mRNA rescue, plasmid lockeT7TS was linearized with XbaI and transcribed using T7 RNA polymerase. 500pg of mRNA per embryo was co-injected with 3 ng per embryo of morpholino (e4i4). To assess splicing defects, total RNA was isolated from morpholino injected embryos or uninjected controls and used to synthesize cDNA libraries for PCR analysis with the SuperScript First-Strand Synthesis System (Invitrogen). Primer sequences used are available upon request.
The ccdc40 antisense probe was prepared from EcoRI linearized BL283 using T3 RNA polymerase. RNA in situ hybridization to analyze organ laterality was performed as described 25 using standard procedures 26. Ccdc40MO injected embryos were collected at 11–13 somites, fixed in 4% paraformaldehyde for 45 min. at room temperature, and processed for immunohistochemistry using standard methods. Antibodies used include: anti-acetylated tubulin monoclonal antibody at 1:400 (IgG2b isotype, clone: 6-11B-1; Sigma St. Louis, MO, USA) and goat anti-mouse IgG2b, FITC-conjugated antibody at 1:400 (Southern Biotech, Birmingham, AL, USA). Nuclei were visualized with Hoechst or Draq5 and F-actin was stained with Rhodamine-phalloidin. Embryos were mounted in 50% glycerol/PBS and analyzed using the Zeiss LSM510. Zebrafish TEM samples were prepared as described 27 and analyzed on a Zeiss 921AB.
We thank the German patient support group "Kartagener Syndrom und Primaere Ciliaere Dyskinesie e.V.", Stefanie Glaser for the initial genomic mapping of locke, Dr. Judy Liu for help with imaging Ccdc40 protein expression in the mouse node, Lori Bulwith, Angelina Heer, Carmen Kopp, Denise Nergenau, and Karin Sutter for excellent technical assistance, and Derrick Bosco for zebrafish facility maintenance. We also thank M. Griese (Munich), E. v. Mutius (Munich), T. Nuesslein (Koblenz), N. Schwerk (Hannover), S. Reithmayr (Vienna), H. Seithe (Nuernberg) and M. Stern (Tuebingen) for supporting the study. The lnks mutant mouse line was established as part of the Sloan-Kettering Institute Mouse Project (R37-HD035455). This work was supported by: Basal O’Conner Award from the March of Dimes, Young Investigator Award from the Spina Bifida Association, and R01-HD058629 to I.E.Z.; the German Human Genome Project DHGP grant 01 KW9919 to R.G.; Howard Hughes Medical Institute to L.N.; “Deutsche Forschungsgemeinschaft” DFG Om 6/4, GRK1104, and the SFB592 to H.O.; and NICHD - R01-HD048584 to R.D.B.
Author contributions. Studies in mice were conducted by I.Z., A.P., A.B-H., H.O., K.V.A. and L.N. Studies in zebrafish were conducted by N.O., K.B.L., J.S-B., J.M., R.G. and R.D.B. Studies with patient samples were conducted by A.B-H., N.T.L., H.O., K.H., M.F., J.H., R.R., K.G.N., J.K.M. G.B. and H.O. The manuscript was prepared by A.B-H, I.E.Z., L.N., H.O. and R.D.B.
The authors have no competing financial interests.