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Dyx1c1 has been associated with dyslexia and neuronal migration in the developing neocortex. Unexpectedly, we found that deletion of Dyx1c1 exons 2–4 in mice caused a phenotype resembling primary ciliary dyskinesia (PCD), a genetically heterogeneous disorder characterized by chronic airway disease, laterality defects, and male infertility. This phenotype was confirmed independently in mice with a Dyx1c1c.T2A start codon mutation recovered from an ENU mutagenesis screen. Morpholinos targeting dyx1c1 in zebrafish also created laterality and ciliary motility defects. In humans, recessive loss-of-function DYX1C1 mutations were identified in twelve PCD individuals. Ultrastructural and immunofluorescence analyses of DYX1C1-mutant motile cilia in mice and humans revealed disruptions of outer and inner dynein arms (ODA/IDA). DYX1C1 localizes to the cytoplasm of respiratory epithelial cells, its interactome is enriched for molecular chaperones, and it interacts with the cytoplasmic ODA/IDA assembly factor DNAAF2/KTU. Thus, we propose that DYX1C1 is a newly identified dynein axonemal assembly factor (DNAAF4).
Cilia, hair-like organelles projecting from the surface of nearly all polarized cell types, serve essential roles in cellular signalling and motility1. The basic structure of motile cilia, and the related organelle flagellum is remarkably conserved throughout evolution. In most motile cilia, a ring of nine peripheral microtubule doublets surrounds a central pair apparatus of single microtubules that connect to the nine peripheral doublets by radial spokes (9+2 structure). Motile monocilia present at the mouse node during early embryogenesis are an exception, lacking the central pair apparatus (9+0 structure). Distinct multi-protein dynein complexes attached at regular intervals to the peripheral microtubule doublets contain molecular motors that drive and regulate ciliary motility. Specifically, outer dynein arms (ODA) are responsible for beat generation, while both the inner dynein arms (IDA) and the nexin link-dynein regulatory complexes (N-DRCs) regulate ciliary and flagellar beating pattern and frequency. Identifying the proteins responsible for correct assembly of this molecular machinery is critical to understanding the causes of motile cilia-related diseases.
Primary ciliary dyskinesia (PCD) (MIM 242650), a rare genetic disorder affecting approximately 1 in 20,000 individuals, is caused by immotile or dyskinetic cilia. Loss of ciliary function in upper and lower airways causes defective mucociliary airway clearance and subsequently, chronic inflammation that regularly progresses to destructive airway disease (bronchiectasis). Organ laterality defects are also observed with approximately half of PCD patients exhibiting situs inversus, and more rarely situs ambiguus, which can associate with complex congenital heart disease2. Dysfunctional sperm tails (flagella) frequently cause male infertility in PCD individuals, warranting assisted reproductive technologies. Another consequence of ciliary dysfunction, particularly evident in mouse models, is hydrocephalus caused by disrupted flow of cerebrospinal fluid through the cerebral aqueduct connecting the third and fourth brain ventricles3. Although ciliary dysmotility is not sufficient for hydrocephalus formation in humans due to morphological differences between the mouse and human brain, the incidence of hydrocephalus, secondary to aqueduct closure, is increased in PCD individuals3.
Genetic analyses of PCD patients have now revealed several autosomal recessive mutations in genes encoding axonemal subunits of the ODA complexes and related components4–12. In addition, recessive mutations in CCDC39 (MIM 613798) and CCDC40 (MIM 613799) have been linked to PCD with severe tubular disorganisation and defective nexin links13,14. Mutations in the radial spoke head genes RSPH4A and RSPH9 as well as in HYDIN can cause intermittent or complete loss of the central apparatus microtubules15–17. Two X-linked PCD variants associated with syndromic cognitive dysfunction and retinal degeneration are caused by mutations in OFD1 (MIM 311200) and RPGR (MIM 312610), respectively18, 19. Another functional class of proteins emerging from identification of PCD causing mutations are proteins involved in cytoplasmic pre-assembly of both ODA and IDA: DNAAF2 (KTU, MIM 612517)20, DNAAF1 (LRRC50, MIM 613190)21, 22, DNAAF3 (C19orf51, MIM 614566)23 and the recently identified LRRC6 (MIM 614930)24.
DYX1C1 (MIM 608706), dyslexia susceptibility 1 candidate 1 gene, was initially identified as a candidate dyslexia gene due to both a single balanced translocation t(2;15)(q11;q21) coincidentally segregating with dyslexia in a family25, and subsequent single nucleotide polymorphism (SNP) association studies. Follow-up gene association studies have provided both positive26–28 and negative29–31 support for association with dyslexia. Molecular and cellular analyses of DYX1C1 have indicated potential functional roles with chaperonins32,33, estrogen receptor trafficking34, and neuronal migration35,36, while recent proteomic and gene expression studies have suggested a possible role in cilia37,38.
In order to reveal the required biological functions of Dyx1c1, we performed a forward genetic experiment by producing an allele of Dyx1c1 in mice in which exons 2–4 were deleted (Fig. 1a). Homozygous mutant mice (Dyx1c1Δ/Δ) expressed no detectable Dyx1c1 protein by western blot analysis of all tissues tested, including brain and lung (Fig. 1b). Mice heterozygous for the deleted allele were viable, fertile, and not noticeably different from wild-type littermates. Homozygous mutant mice were recovered after birth from heterozygous breeding pairs at rate deviating from the expected Mendelian ratio (295:570:87, (Dyx1c+/+:Dyx1c1Δ/+:Dyx1c1Δ/Δ), Chi=128.017, p<0.0001) but were recovered at early embryonic times (gestational days E6.5–E12) at the expected Mendelian ratio (22:43:20; Chi=0.105, p=0.85) suggesting embryonic lethality of approximately 2/3 of homozygous mutants. Homozygous mutants that survived after birth developed severe hydrocephalus by postnatal (P) day 16 (Fig. 1c), and died by P21, similar to what has been described for other mouse mutants with defective motile cilia3,39.
In addition to hydrocephaly, postnatal homozygous mutant mice displayed laterality defects with 59% of mutants (51/87) showing situs inversus totalis, a complete inversion of left right asymmetry (Fig. 1d), 17% (15/87) displaying situs ambiguous with inverted heart and lung position relative to stomach and spleen, or inverted position of stomach and spleen relative to heart and lung position, and 24% (21/87) displaying situs solitus, normal left-right asymmetry. Mutations that cause disruptions in left-right asymmetry in mice40 are known to result from defective function of motile nodal monocilia, and more specifically, the loss of cilia-generated leftward flow across the node in early embryogenesis40. The typical phenotype for laterality mutants is a 1:1 ratio for situs inversus totalis and situs solitus indicating a randomization of L-R patterning, although the inv mutant mouse shows a 100% rate of defects in L-R patterning. Even if the ratio of situs inversus to situs solitus seen in surviving Dyx1c1 mutant mice deviates significantly from what would be expected for a 1:1 ratio of situs inversus to situs solitus (51:21, Chi: 12.5, p=0.0004), this ratio was already observed in other L-R patterning mutants like the Dnahc5 mutant mouse41. Consistent with a role for Dyx1c1 function in the embryonic node where L-R patterning is established in the mouse, we found by whole mount in situ hybridization that Dyx1c1 expression in the early embryo (E7.5) is restricted to pit cells of the embryonic node (Fig. 1e).
In an independent mouse ENU mutagenesis screen for congenital heart defects (CHD) of the NHLBI Bench to Bassinet Program, a mutant named Sharpei was recovered with a Dyx1c1 missense mutation (c.T2A) that resulted in an altered AUG start codon, and expression of an aberrant N-terminal truncated Dyx1c1 protein product of approximately 31 kDa (Supplementary Figure 1a). This mutant was recovered based on the finding of complex congenital heart and therefore carefully histologically phenotyped. Some mutants with apparent situs inversus comprising dextrocardia with inverted lung lobation and right sided stomach also had subtle visceral organ situs defects, such as discordant left sided pancreas and spleen despite right sided stomach positioning (Supplementary Figure 1b). Sharpei mutants with complex CHD died prenatally or at birth with a spectrum of complex CHD, such as transposition of the great arteries with ventricular septal defect and coronary fistula (Supplementary Fig. 1b and Supplementary Video 1) or double outlet right ventricle with right atrial isomerism, muscular VSD, and atrioventricular septal defects (Supplementary Fig. 1b and Supplementary Videos 2 and 3). All these findings resemble congenital heart defects observed in other mouse models for PCD such as Dnahc5 mutant mice41.
Recently, Chandrasekar et al. showed by knockdown of dyx1c1 results in phenotypes characteristically associated with cilia defects such as body curvature, hydrocephalus, cystic kidneys and situs inversus42. To test whether Dyx1c1 has an evolutionarily conserved role in establishing left-right asymmetry in vertebrates, we also performed morpholino-mediated knockdown experiments in zebrafish embryos. Zebrafish dyx1c1 is expressed in embryonic tissues that contain motile cilia42, including Kupffer's vesicle, which is known to play an important role in establishing the left-right axis. Morpholino knockdown of zebrafish dyx1c1 resulted in hydrocephalus, kidney cysts and body axis curvature; phenotypes consistent with ciliary dysfunction in zebrafish. Morpholino knockdown of dyx1c1 also produced laterality defects (Fig. 2a,b) as assessed by the position of the heart (cmlc2), liver (fkd2) and pancreas (ins) in dyx1c1 morphant embryos at 48 hours post-fertilization (hpf). In a wild-type zebrafish embryo, the ventricle of the heart loops towards the right and the atrium loops towards the left and the liver is positioned to the left of the midline, and the pancreas lies to the right of the midline (Fig. 2a). This wild-type pattern was observed in 36.5% of the morphants, whereas 37.9% of the morphants showed a complete reversal of the placement of the organs (Fig. 2b). A heterotaxic phenotype was seen in 25.6% of the embryos (Fig. 2b). To investigate when dyx1c1 affects early left-right patterning, the expression of the zebrafish nodal gene southpaw (spaw) was studied in a timecourse of morpholino injected embryos. The expression of zebrafish spaw is restricted to the left LPM during somitogenesis prior to asymmetric organ positioning at 48 hpf43. At the 12- to 14-somite stage, 82.8% of the control embryos had left LPM expression of spaw whereas all of the morphant embryos lacked any expression of spaw, indicating that spaw initiation is delayed in the absence of dyx1c1 (Fig. 2b). The delay in spaw expression upon loss of ciliary motility in KV has not been reported previously but has been observed in other ciliary motility mutants that do not affect KV structure (R.D.B. unpublished). At the 16- to 18-somite and 20- to 22-somite stage, most of the control embryos (97.7% and 98.8% respectively) had spaw expression in the left LPM; but the morphants displayed a randomized pattern; bilateral (30% and 30.6% resp.), left-sided (20% and 29.4% resp.), right-sided (26.3% and 30.6% resp.) or absent (23.8% and 9.8% resp.) (Fig. 2b). Thus, in the zebrafish embryo, dyx1c1 is necessary for left-sided expression of spaw in the left LPM which is a crucial step for normal left-right axis development. Overall our results confirm the findings from Chandrasekar et al42. Furthermore, we demonstrate that knockdown of dyx1c1 affects the left-sided expression of spaw consistent with an important role in the left-right axis development.
Hydrocephalus and organ laterality defects are hallmarks of mutations that cause defects in ciliary motility in mice and zebrafish1,3. We therefore used light and electron microscopy as well as videomicroscopy to determine whether Dyx1c1 deficiency caused loss of motile cilia formation or a loss of cilia. Cilia extending from mouse ependymal cells of the cerebral ventricles, visualized by light microscopy, appeared similar in length and distribution in Dyx1c1Δ/Δ and wild-type littermates (Fig. 3a). Although cilia were abundant on respiratory epithelial cells in mutants and wild-type, cilia in mutants lacked immunofluorescence signal for both the outer dynein arm heavy chain Mdnah5, and the inner dynein arm light chain, Dnali1 (Fig. 3b,c). Loss of these dyneins from motile cilia would predict a loss of motility. We used live-cell imaging to directly assess cilia-mediated fluid flow and ciliary motility on the ependymal surface in Dyx1c1Δ/Δ mice. Explants or slices of the lateral ventricular surfaces were prepared from Dyx1c1Δ/Δ and wild-type mice at P6. Ependymal cilia in wild-type mice continued to beat vigorously in these preparations, and created a directional fluid flow across the surface that could be visualized by the displacement of a small volume of India ink pressure injected onto the ependymal surface. This flow was present in all tested wild-type mice tested (n=6; Fig. 3d and Supplementary Video 4) but was completely missing in tissue obtained from all Dyx1c1Δ/Δ mice tested (n=4; Fig. 3d and Supplementary Video 5). We next examined the motility of ependymal cilia in coronal brain slices of wild-type and Dyx1c1Δ/Δ mice by infrared-DIC videomicroscopy. Cilia at the ependymal surface in wild-type and heterozygous mice were found to beat at a frequency of approximately 9 beats/sec (n=8; 34°C; Supplementary Video 6), while cilia on ependymal cells from all Dyx1c1Δ/Δ examined (n=4) lacked ciliary beating (Supplementary Video 7). Videomicroscopy of tissue slice from the third brain ventricle in newborn homozygous Sharpei mutants also showed completely immotile cilia (n=7, Supplementary Video 8). Beads added to the solution above the brain slice exhibited only random motion, while in wild-type littermate controls, the beads showed ependymal cilia generated flow (Supplementary Video 8). Similarly, videomicroscopy of the tracheal airway epithelia in newborn homozygous Sharpei mutants showed completely immotile cilia, consistent with PCD, while littermate controls showed normal rapid synchronous ciliary beat (Supplementary Video 9). We also assessed the motility of cilia in Kupffer's vesicle in dyx1c1 morphant zebrafish with videomicroscopy. Dyx1c1 morphants had cilia in Kupffer’s vesicles that lacked motility compared to uninjected embryos (Supplementary Video 10 and 11).
To assess the ultrastructure of respiratory cilia in mutants we obtained transmission electron micrographs of tracheal cilia. As on the ependymal surface, tracheal cilia were abundant in both wild-type and Dyx1c1Δ/Δ cells (Fig. 3e), but unlike tracheal cilia in wild-type mice, cilia in Dyx1c1Δ/Δ trachea were surrounded by cellular debris and mucus (Fig. 3e). We further examined the ultrastructure of tracheal cilia in cross sections by TEM. As shown in Fig. 3f, tracheal cilia in wild-type and mutant mice had typical 9+2 microtubular structure with intact radial spokes. In contrast, and consistent with the absence of heavy and light chain dynein subunits (Fig. 3b), cilia in Dyx1c1Δ/Δ mice lacked both ODA and IDA structures in respiratory cilia (Fig. 3f). Thus, the phenotype of Dyx1c1 loss of function is a severe ciliary motility defect associated with absent axonemal ODA and IDA structures.
The apparent conservation of function of mouse and zebrafish Dyx1c1/dyx1c1 encouraged us to search for mutations in DYX1C1 in patients with PCD, the human disorder connected to defective ciliary motility. Because of the observed ultrastructural phenotype and severe ciliary beating defect in Dyx1c1Δ/Δ mice, we considered DYX1C1 an excellent candidate gene for PCD with abnormal axonemal ODA and IDA assembly. DYX1C1, located on chromosome 15q21.3, comprises 10 exons (translation starts in exon 2) and encompasses 77.93 kb of genomic DNA. In one highly inbred Irish family (UCL-200), a CNV analysis of whole exome sequence data (using ExomeDepth)44 identified a homozygous DYX1C1 deletion in two affected siblings (UCL-200 II:1 and II:2; Supplementary Figure 2a,b). This finding was confirmed by Sanger sequencing of the deletion breakpoints and was present in heterozygous state in their carrier mother (Supplementary Figure 2c). The 3.5 kb deletion leads to the loss of exon 7 of DYX1C1 (Fig. 4a). Interestingly, the same 3.5 kb deletion was also identified in individuals with PCD in five American/Australian families. Five of these individuals were heterozygous for the deletion along with a mutation in the splice donor site of exon 6, stop mutations and a frame shift mutation in the other allele (UNC-158, UNC-159, UNC-1669, UNC-1839, UNC-1171, Fig. 4a and Supplementary Fig. 3e–h) and one individual was homozygous for the deletion (UNC-663, Supplementary Figure 3i). In addition we screened all DYX1C1 exons and adjacent intronic sequences by PCR amplification and subsequent Sanger DNA sequencing in 105 PCD individuals with combined ODA and IDA defects. Mutations were identified in ten affected individuals from nine unrelated families that predicted premature termination of translation (Fig. 4a,b and Table 1). In all analysed families, the mutations segregated with the disease status consistent with an autosomal recessive inheritance pattern (Supplementary Fig. 2 and 3). In three families, recessively inherited homozygous mutations were confirmed by sequencing of other family members (F648 II1, OP-359 II1, OP-556 II1). In another family, the PCD-affected individual, OP-86 II2, was compound heterozygous for two different mutations, with each mutation tracking uniquely to one of the parents (Supplementary Fig. 3a). A total of 9 human DYX1C1 mutations were therefore defined (Supplementary Table 1). Apart from the 3.5 kb deletion and a splice site mutation, the other seven detected mutations predicted premature protein termination and clustered towards the middle of the 420 amino acid sequence between amino acids 128 and 195 (Fig. 4b). Thus, remarkably, seven of the nine identified mutations predict a truncated DYX1C1 protein that would lack more than half of the protein, including the carboxyl-terminal tetratricopeptide repeat domains (TPR). The TPR domains in DYX1C1 have been shown previously to be functionally important domains required for DYX1C1 in neuronal motility, cellular localization, and interaction with molecular chaperones33,35.
All twelve PCD patients with recessive DYX1C1 mutations suffer from classical symptoms of PCD, including recurrent upper and lower airway disease and bronchiectasis. Seven patients had neonatal respiratory distress syndrome. Four patients exhibited reduced fertility (Supplementary Table 1) and one male patient was treated for infertility, having three children by assisted reproduction, suggesting a probable DYX1C1 function also in sperm tails. Five of the twelve affected individuals display situs inversus totalis (42%), two have situs ambiguous (16%), one with dextrocardia and polysplenia and one with left atrial isomerism and polysplenia, and five have situs solitus (42%). Thus, DYX1C1 deficiency in PCD patients causes disruption of left-right body asymmetry, similar to findings observed in mouse and zebrafish. Interestingly, no patient was diagnosed with dyslexia or with hydrocephaly. Hydrocephalus is a common phenotype in mouse mutants with immotile cilia but rare in human patients3,39. Two patients had a learning disability, but these can likely be attributed to other causes including microcephaly (OP-86 II2, UCL-200 II:1).
Respiratory cilia isolated from patients with biallelic DYX1C1 mutations displayed severe ultrastructural defects (Fig. 4c–f) resembling those defects observed in mouse Dyx1c1 mutants (Fig. 3b,c and e). Specifically, both ODA and IDA defects were present in TEM analyses in all ten assessed probands (OP-86 II2, OP-556 II1, UNC1669, UNC1839, UNC1171, UNC158, UNC159, UNC663, UCL-200 II:1 and II:2) (Fig. 4c and Supplementary Table 1). To further understand the defect on the molecular level, we performed immunofluorescence microscopy of cilia using antibodies targeting components of the axonemal ODAs, IDAs and N-DRC. Immunofluorescence analysis revealed an absence or marked reductions of proteins normally present in type-1 and type-2 ODA complexes (DNAH5, DNAH9, and DNAI2; Fig. 4d,e and Supplementary Fig. 4–6) as well as IDA subtype complexes (DNALI1; Fig. 4f and Supplementary Fig. 7). These findings are similar to cytoplasmic pre-assembly (DNAAF) defects reported in DNAAF2 (KTU)20, DNAAF1 (LRRC50)21,22, and DNAAF3 (C19orf51)23 mutant ciliary axonemes. Interestingly, we found that the extent of axonemal ODA defects varied among the respiratory cells tested, and in some cases the ODA proteins DNAH5 and DNAI2 could be detected in the axonemes (Supplementary Fig. 4 and 6). Assembly of proximal type-1 ODA complexes (DNAH9 negative and DNAH5 positive) appeared to be better preserved than distally localized type-2 ODA complexes (DNAH5 and DNAH9 positive; Supplementary Fig. 4–6). To further understand the functional consequences of our observations, we performed nasal brush biopsy in patients (F-648 II1, OP-86 II2, UNC1669, UNC1839, UNC1171, UNC663, UCL-200 II:1 and II:2) and analysed respiratory cilia beating by high-speed videomicroscopy. Videomicroscopy revealed that most respiratory cells had immotile cilia (Supplementary Video 12 and 13); however, cilia were found to beat in some respiratory cells, albeit with reduced frequency (Supplementary Video 14 and 15) when compared to control cilia (Supplementary Video 16). These variable motility defects in DYX1C1 mutant cells are consistent with our previous observations of variable degrees of axonemal ODA defects. Interestingly, only mutations in DNAAF2 (KTU) in human patients have revealed respiratory cells with similarly variable ODA defects and associated IDA abnormalities20.
The phenotypes described above and previously reported interactions between exogenously expressed DYX1C1 protein and molecular chaperones suggested to us that DYX1C1 may function as a newly identified cytoplasmic axonemal dynein assembly factor. Consistent with this possibility, we found by immunofluorescence that Dyx1c1 protein is located in the cytoplasm of respiratory epithelia (Fig. 5a). We confirmed this finding by immunoblot by showing that similar to DNAAF2, DYX1C1 is also present in the cytoplasmic protein fraction and almost undetectable in the axonemal protein fraction of human respiratory cells. (Fig. 5b). Considering the similarities in phenotype caused by DYX1C1 and DNAAF2 mutations with regard to variable ODA defects, we tested for possible interactions between these proteins. Using myc- and FLAG-tagged proteins that were coexpressed in HEK293 cells, we found by co-immunoprecipitation that DYX1C1 interacted with DNAAF2/KTU (Fig. 5c), but not the cytoplasmic pre-assembly factors DNAAF1 and DNAAF3 or the PCD-associated protein CCDC103 that localizes in both the cytoplasm and axoneme and plays a role in dynein arm attachment (Supplementary Fig. 8), as well as the newly identified dynein arm assembly protein LRRC6 (data not shown). Furthermore, we demonstrate that DYX1C1 and DNAAF2 interact directly by yeast two-hybrid assay (Fig. 5d). Based on these findings, we hypothesize that DYX1C1 represents a novel cytoplasmic axonemal dynein assembly factor possibly acting together with DNAAF2/KTU at an early step of cytoplasmic ODA and IDA assembly.
To categorize the molecular function of Dyx1c1 more completely in respiratory tissue we defined the protein interactome of Dyx1c1 in mouse trachea by co-immunoprecipitation and tandem mass spectroscopy (MS/MS). Input extracts for co-immunoprecipitation were prepared from trachea tissue of either wild-type or mutant mice and these appeared similar in protein composition as evaluated by coomassie staining after SDS-PAGE (Supplementary Fig. 8a). Following co-immunoprecipitation with anti-Dyx1c1 antibodies coomassie stained protein bands were apparent in preparations from wild-type extracts but not in mutant extracts (Supplementary Fig. 8a). Fourteen matched pairs of gel pieces covering the range of approximately 20 to 200 kDa, with many pieces containing mutiple bands, were cut from wild-type and mutant co-immunoprecipitations and subjected to tryptic digest and tandem mass spectrometry analysis for protein identification. In all, 702 proteins were positively identified in immunoprecipitates from wild-type trachea, while a total of 29 proteins were identified in gel slices from the mutant immunoprecipitates (Supplementary Table 2). To determine whether the identified Dyx1c1 protein interactome was enriched for particular molecular or biological functions we used DAVID to determine the presence of enriched Panther Gene Ontology terms in the Dyx1c1 interactome and proteins identified in the knockout control. Using a mouse lung proteome as a background we found several molecular functional categories enriched in the Dyx1c1 interactome and several of these were in the categories containing molecular chaperones (MF00077: Chaperone, p<0.001, BP00062:Protein folding, p<0.01, BP00072:Protein complex assembly, p<0.05, and MF00078:Chaperonin, p=0.05, Supplementary Table 3). All four of these chaperone containing categories were not significantly enriched in the protein set identified in the mutant immunoprecipitates (Supplementary Table 3). The Dyx1c1 interactome contains a total of 28 proteins in the chaperone or chaperonin category including 6 of 8 subunits in the T-compex chaperonin complex, and multiple heat shock proteins. We confirmed by co-immunoprecipitation and western blot analysis six chaperones interacting with Dyx1c1 including Cct3, Cct4, Cct5, Cct8, Hsp70 and Hsp90 (Supplementary Fig. 8b).
The protein interactions we found in tracheal tissue with endogenous Dyx1c1 are in agreement with results from a recent study using neuroblastoma cell lines and exogenously expressed Dyx1c145. Our results provide the first evidence that may link the T-complex of chaperones, known primarily for their role in cytoskeletal protein assembly, to the folding and assembly of axonemal dynein complexes. Interestingly, evidence for a role of Ccts in cytoplasmic assembly of protein complexes required for motile cilia has been reported for Tetrahymena46. Although the Dyx1c1 interactome defined with tandem mass spectrometry did not include any of the known DNAAF proteins absence in an MS-MS screen does not necessarily mean absence of potential interaction and highlights to screen for potential interactors by several means.
This study is the first to show the effects of DYX1C1/Dyx1c1/dyx1c1 deficiency in human, mouse and zebrafish. The phenotypes in all species are consistent in showing a selective defect in motile cilia reflecting deficient dynein arm transport or assembly. Furthermore, since the human patients carrying mutations in DYX1C1 showed no evidence of dyslexia, we propose that the loss-of-function of DYX1C1 may not be a highly penetrant cause of dyslexia. In conclusion, based on the pattern of cilia defects, its cellular localization, a protein interactome enriched for chaperones, and genetic-physical interaction with DNAAF2, we propose that DYX1C1 represents a novel axonemal dynein assembly factor (DNAAF4).
Sense and antisense probes for Dyx1c1 were made from a 554 bp pCRII-TOPO construct (nt 1098 – 1651; RefSeq NM_026314.3, Mus musculus Dyx1c1 transcript variant 1) made by TOPO TA cloning (Invitrogen) after amplification from complementary DNA. Primers used for amplification were 5´-aaacctacacaaggccatcg-3´ (Dyx1c1-F) and 5´-atcctggcaatttcaacagc-3´(Dyx1c1-R). Probes were synthesized with digoxigenin NTPs (Roche) after template linearization with Hind III (antisense) or Not I (sense) before RNA synthesis with T7 or SP6 RNA polymerases, respectively. For whole mount in situ hybridization (WISH) staged embryos were fixed overnight at 4°C in 4% paraformaldehyde in 1× PBS. WISH was then performed according to standard procedures with minor modifications47. Stained samples were transferred into 80% glycerol, and images were captured using a Scion CFW-1310C color digital camera mounted on an Axioskop2 plus microscope (Zeiss) and Image-Pro Express.
Mice were perfused transcardially with 0.9% saline followed by 4% paraformaldehyde (Electron Microscopy Science, Hatfield, PA) in 1× PBS. Brains were removed, fixed overnight in the same fixative at 4°C, and washed in 1× PBS three times for 40 min the next day before cutting into 50 µm sections with a vibratome (VT-1000S; Leica, Wetzlar, Germany). Nuclei were counterstained with Hoechst33342 (2µg/ml PBS, Sigma). Stained sections were washed for 10 min in PBS, coverslipped using prolong gold anti-fade (Invitrogen), and imaged on a Carl Zeiss Axiovert 200M Inverted microscope with an ApoTome attachment and Axiovision 4.6 software (Carl Zeiss). Whole mounts of the entire lateral wall of the lateral ventricles were prepared as described previously48 and fixed overnight in 4% paraformaldehyde in PBS at 4°C, and washed in 1× PBS three times for 40 min the next day. They were then permeabilized with 0.1% Triton X-100 (Sigma, St. Louis, MO) in PBS for 10 min, blocked in 10% goat serum (Invitrogen, Carlsbad, CA) in PBS/0.1% Triton X-100 for 1 h, and incubated with the following primary antibodies: mouse anti-acetylated tubulin (1:500; Sigma, T6793); rabbit anti-β- catenin (1:100; 9562, Cell Signaling Technology, Beverly, MA) and mouse anti-γ-tubulin (1:500; T6557, Sigma). After washing three times in PBS, tissues were incubated with appropriate Alexa Fluor dye-conjugated secondary antibodies (Goat anti-rabbit 488 [A11008], Goat anti-mouse 488 [A11001], Goat anti-rabbit 568 [A11011], Goat anti-mouse 568 [A11031] Invitrogen) at a dilution of 1:400 for 1 h. Tissues were washed in PBS and incubated for 5 min in 2 µg/ml Hoechst33342 (Sigma) for counterstaining of nuclei. Secondary antibodies alone were used as a control. Whole mounts were placed onto depressed glass slides and coverslipped with CoverWell imaging chambers (Grace Bio-Labs, Bend, OR). Samples were imaged either on a Carl Zeiss Axiovert 200M Inverted microscope with an ApoTome attachment and Axiovision 4.6 software (Carl Zeiss) or on a Leica TCS SP2 confocal laser-scan microscope.
Nasal epithelial cells were harvested from mouse septa by acute dissociation on a 4-well slide in 1:1 PBS/HBSS. The slides were dried to let the cells stick to the wells and then immediately fixed with 4% PFA for 2 min at RT. The cells were then washed with 1× PBS for 2 min each twice. Blocking buffer (10% Goat serum, 0.1% Triton in 1X PBS) was added for 15 mins at RT. The slides were incubate with rabbit polyclonal anti-DYX1C1 antibody (Sigma SAB4200128) at a dilution of 1:500 and mouse monoclonal anti-acetylated tubulin (Sigma, T7451) at 1:1000 in the same blocking buffer for 30 mins at RT. After 3 washes with 1X PBS for 5 min each, the slides were incubated with goat anti-rabbit 488 (Invitrogen, A11034 dilution 1:5000) and goat anti-mouse 568 (invitrogen, A11031 dilution 1: 1000) for 20 min at RT. The slides were washed 2 times with 1X PBS for 5 min each and nuclear stain Hoechst33342 (Invitrogen, H3570) at 1:2500 was added for 15 min at RT. Slides were dried and coverslipped with Prolong Gold antifade (Invitrogen, P36930). The cells were imaged on Leica TCS Sp2 laser scan microscope.
For harvesting the brains, mice (12 days old) were perfused transcardially with 0.9% saline followed by 2% paraformaldehyde/2.5% glutaraldehyde in 0.1 mM phosphate buffer (PB). Brain samples were further fixed by immersion overnight in 2% paraformaldehyde/2.5% glutaraldehyde in 0.1 mM PB; and washed in PB three times for 40 min. Sections were postfixed with 2% OsO4 in 0.1 mM PB for 1.5 h and dehydrated through a graded ethanol (EtOH) series. Following dehydration, tissues were twice washed in acetone and embedded in epoxy resin in capped inverted Beem capsules. Thin sections were cut with a diamond knife, placed onto Formvar-coated slot grids, and heavy metal stained with uranyl acetate and lead citrate. Trachea tissues were directly dissected without perfusion and fixed by immersion overnight in 2.5% glutaraldehyde/2% paraformaldehde in 0.12 mM PB. They were washed in PB three times (60 min in total), postfixed in 1% OsO4 and 0.8% potassium ferricyanide in 0.12M PB for 1 hour, dehydrated through graded EtOH series, rinsed twice in acetone and embedded in epoxy resin. Thin sections were cut with diamond knife, placed on copper grids, and heavy metal stained with ethanolic uranyl acetate and Sato's lead citrate. Electron micrographs were captured using an FEI Tecnai 12 Biotwin TEM equipped with a side mounted AMT XR-40 CCD Camera and Epson Expression 1680 flatbed scanners.
Wildtype and mutant mice embryos (E7.5 days) were harvested and fixed in 1.5% paraformaldehyde/1.5% glutaraldehyde (Electron Microscopy Science) in 0.10 M sodium cacodylate containing 0.05 M NaCl overnight at 4°C. Samples were postfixed with 2% OsO4 in the same buffer overnight followed by graded ethanol dehydration. Specimens were dried in a Polaron (Hertfordshire, UK) E3000 Critical Point Dryer and mounted onto aluminum specimen mounts (Ted Pella) using carbon tape and silver paint (Ernest F. Fullam, Clifton Park, NY). Each mount was sputter coated with gold palladium (60% gold, 40% palladium) using a Polaron E5100 Sputter Coater. Samples were examined and photographed using a LEO DSM982 field emission SEM.
Wild-type and mutant animal brains and lungs were harvested and lysed in RIPA Buffer (Sigma) supplemented with 1× Protease Inhibitor Cocktail (Sigma). For tracheal and lung preparation, tissues were dissected and carefully separated from the surrounding tissues. The samples were homogenized using a tissue homogenizer and cleared by centrifugation at 10,000×g for 10 min. Proteins were separated on 10% SDS-PAGE minigels and then transferred to Immobilon (Millipore Inc., Billerica, MA, USA) membrane for Western blotting. For detecting Dyx1c1 protein, the N-terminal DYX1C1 (Sigma, SAB4200128) antibody was used at a dilution of 1:200, anti-Gapdh antibody (Sigma, G8795) was used at 1:1500 as a loading control. Licor Odyssey infrared secondary antibodies were used at a dilution of 1:10,000 (goat anti-mouse 680 [926–32220]; goat anti-mouse 800 [926–32210]; goat anti-mouse 680 [926–32221]; goat anti-mouse 800 [926–32211]) were used at dilution of 1:10,000. All blots were imaged and analysed by Licor Odyssey Scanner and Software.
Immunoprecipitation assay was performed using Dynabeads Protein G Immunoprecipitation Kit (Invitrogen). Briefly, Dynabeads were resuspended in the vial and separated on a magnet from the solution. N-terminal Dyx1c1 antibody (5µg) was diluted in 200 µL of Washing and Binding Solution and incubated with rotation for 60 min at room temperature. The beads-antibody complexes (beads-Ab) were separated on the magnet, washed by gentle pipetting and separated. Protein lysates as described earlier from the wildtype and mutant mice brains were incubated with the beads-Ab overnight at 4C. The beads-Ab-antigen complex was then washed using the washing solution 3 times. The complex was then incubated with elution buffer for 10–15 mins to dissociate the complex. The beads were separated on a magnet and the supernatant containing the proteins was separated by SDS–PAGE and analysed by western blotting using anti-DNAI2 monoclonal antibody (M01, clone 1C8, Abnova, 1:500), anti-Hsp70 (BD Biosciences, 610607, 1:1000) antibody, anti-Hsp90 (BD Biosciences, 610418, 1:1000) antibody, anti-CCT4 (Aviva Systems Biology, ARP34271_P050, 1:500), anti-CCT3 antibody (Proteintech, 10571-1-AP, 1:500), anti-CCT5 antibody (Proteintech, 11603-1-AP, 1:500), anti-CCT8 antibody (Proteintech, 12263-1-AP, 1:500) and anti-IC74 monoclonal antibody (gift from Dr. Stephen King, 1:750). Rabbit IgG was used as a control.
P6–P10 wildtype and mutant mice were deeply anesthetized with isoflurane and then decapitated. Brains were rapidly removed and immersed in ice-cold oxygenated (95% O2 and 5% CO2) dissection buffer containing (in mM): 83 NaCl, 2.5 KCl, 1 NaH2PO4, 26.2 NaHCO3, 22 glucose, 72 sucrose, 0.5 CaCl2, and 3.3 MgCl2. The lateral wall of the lateral ventricle was dissected using a fine scalpel and forceps and immediately observed in a chamber containing 37°C buffer. For visualization of flow, a small amount of Indian ink was placed on the surface of the lateral wall of the dissected ventricle. Movements of Indian ink were observed and recorded with an IR differential interference microscopy (DIC) (E600FN, Nikon) and a CCD camera (QICAM, QImaging, 120fps). For direct observation of cilia movement, mice brains were harvested as above and coronal slices (400 µm) were cut using a vibratome (VT1200S, Leica). Slices from the third ventricle through the fourth ventricle were visualized using IR differential interference microscopy (DIC) (E600FN, Nikon) and a CCD camera (QICAM, QImaging, 120fps). The images were analysed with ImageJ software (NIH).
DIG-labelled RNA probes were transcribed from linearized DNA templates and used in RNA in situ hybridization by standard methods. Antisense probes included cardiac myosin light chain (cmlc2; myl7 – ZFIN)49, forkhead 2 (fkd2; foxa3 – ZFIN)50, preproinsulin (ins)51 and southpaw (spaw)43.
Images of live zebrafish embryos were taken using the ProgressC14 digital camera (Jenoptik) mounted on a Leica MZFL III microscope. Embryos processed for in situ hybridization analysis were mounted in modified GMM52 [100 ml Canada Balsam (C-1795, Sigma), 10 ml methylsalicylate (M0387-100G, Sigma)], visualized using a Leica DMRA microscope at 10× magnification, and photographed with the ProgressC14 digital camera.
Signed and informed consent was obtained from patients fulfilling diagnostic criteria of PCD53 and family members using protocols approved by the Institutional Ethics Review Board at the University of Muenster and University College London Hospital NHS Trust. Genomic DNA was isolated by standard methods directly from blood samples or from lymphocyte cultures after Epstein-Barr virus transformation. Exome analysis of family UCL200 was performed as part of the UK10K Project, as previously described15. Amplification of 10 genomic fragments comprising all 10 exons of DYX1C1 was performed for each exon and patient in a volume of 50 µl containing 30 ng DNA, 50 pmol of each primer, 2 mM dNTPs, and 1.0 U GoTaq DNA polymerase (Promega Corporation, Wisconsin, USA). PCR amplifications were carried out by means of an initial denaturation step at 94°C for 3 min, and 33 cycles as follows: 94°C for 30 sec, 58–60°C for 30 sec, and 72°C for 70 sec., with a final extension at 72°C for 10 min. PCR-products were verified by agarose gel electrophoresis, purified by PCR product pre-sequencing kit (USB, Ohio, USA) and sequenced bi-directionally using BigDye Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems, California, USA). Samples were separated and analysed on an Applied Biosystems 3730×l DNA Analyzer. Sequence data were evaluated using the Codoncode software (CodonCode Corporation, Dedham, USA).
Transmission electron microscopy of human respiratory cilia was performed as previously described17.
Ciliary beat was assessed with the SAVA system54. Respiratory epithelial cells were viewed with an Olympus IMT-2 microscope (40 phase contrast objective) equipped with a Redlake ES-310Turbo monochrome high-speed video camera (Redlake, San Diego, CA) set at 125 frames. The ciliary beating pattern was evaluated on slow-motion playbacks.
Respiratory epithelial cells were obtained by nasal brush biopsy (Engelbrecht Medicine and Laboratory Technology) 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 (all percentages are v/v) before incubation with primary (2–3h at room temperature or overnight at 4 °C) and secondary (25 min at room temperature) antibodies. Appropriate controls were performed omitting the primary antibodies. Mouse monoclonal anti-DNAI2 (1:200; H00064446-M01 (clone1C8)) was obtained from Abnova. Mouse monoclonal anti-acetylated tubulin (1:10000; T7451-200UL) and rabbit polyclonal anti-CCDC39 (1:300; HPA035364) were obtained from Sigma. Rabbit polyclonal anti-DNAH5 and anti-DNALI1 antibodies were generated as reported55,56. Highly cross-adsorbed secondary antibodies goat anti-mouse Alexa Fluor 488 (1:1000; A11029) goat anti-rabbit Alexa Fluor 546 (1:1000; A11035) were from Molecular Probes (Invitrogen). DNA was stained with Hoechst33342 (1:1000; 14533-100MG, Sigma) or DAPI (1:1000; 32670-25MG-F, Sigma). Immunofluorescence images were taken with a Zeiss Apotome Axiovert 200 and processed with AxioVision 4.8 and Adobe Creative Suite 4.
The following clones were purchased from Origene (Rockville, USA): DYX1C1 (cat no. SC313387), DNAAF3/C19orf51 (cat no. SC126165), and CCDC103 (cat no. RC208345). cDNA clones for DNAAF2/KTU and DNAAF1/LRRC50 were amplified in a two-step (nested) PCR reaction from human bronchial epithelial cell cDNA (ScienCell, cat no. 3214) for Gateway cloning. All PCR products were amplified using KOD polymerase according to manufacturer´s directions, recombined with the pDONR201 Gateway vector via BP Clonase II reaction, and subcloned into Gateway entry vectors for myc and 3×FLAG via LR Clonase reaction.
Human respiratory cells were obtained either by brushing or cell culture (spheroids)57. Cells were incubated in 800–1000 µl NP40 or RIPA lysis buffer containing protease inhibitor cocktail (P8340, Sigma-Aldrich) on ice for 30 min and occasionally vortexed. Lysates were spun at 14,000 rpm at 4°C for 10 min. Supernatants were removed into a new tube (cytoplasmic fraction). Pellets were resuspended and incubated in 100–150 µl modified Reeds High salt extraction buffer58 (30 mM HEPES pH 7.4, 5 mM MgSO4, 0.1 mM EDTA, 625 mM NaCl, 2 mM DTT, 70 mM β-Mercaptoethanol, 0.1% Triton-X 100) lysis buffer containing protease inhibitor cocktail (P8340, Sigma-Aldrich) on ice for 30–60 min and frequently vortexed. Lysates were spun down at 14,000 rpm at 4°C for 10 min. Supernatants were removed into a new tube (axonemal fraction). Lysates were controlled by silver staining using the ProteoSilver silver staining kit (PROTSIL1-1KT, Sigma-Aldrich) and stored at −20°C or −80°C until use. Using this method we obtained two protein fractions, the first enriched for cytoplasmic proteins (cytoplasmic fraction) and the second enriched for axonemal proteins (axonemal fraction). We confirmed enrichment of cytoplasmic proteins with anti-DNAAF2 antibodies (Fig. 5b) and axonemal proteins with anti-LRRC48 antibodies (Fig. 5b).
HEK293 cells were transfected with plasmids encoding myc- and FLAG-tagged cDNA constructs using Gene Juice (Novagen) at approximately 0.1 µg DNA per ml of media. Within 24 hrs, cells were collected in 1× PBS and lysed in 1 ml of the following buffer: 50 mM Tris-Cl, pH 8.0, 150 mM NaCl, 1% IGEPAL, 0.5 mM EDTA, and 10% glycerol supplemented with protease (Roche Complete) and phosphatase inhibitors (Cocktails 2 and 3, Sigma Aldrich). Lysates were centrifuged at 16,000 × g for 30 min. at 4°C. Approximately 2 mg of each lysate was precleared with 4 µg of rabbit control IgG antibody for 2 hrs. at 4°C, and then incubated with MagSi/protein A beads (MagnaMedics, Germany) for 1 hr. Lysates were then incubated with 4 µg of rabbit anti-FLAG or anti-myc antibody overnight at 4°C, and then incubated with MagSi/protein A beads for 1 hr. to capture immunoprecipitates. Bead complexes were washed four times in lysis buffer and then resuspended in 1× LDS buffer supplemented with DTT (1/8 lysis volume) and heated for 10 min. at 90°C. Lysates were electrophoresed in NuPAGE 4–12% Bis-Tris gels, transferred to PVDF filters, and subsequently immunoblotted with either anti-myc (A7) or anti-FLAG (M2) mouse monoclonal antibodies. PVDF filters were washed three times in TBS-T (10 minutes each) before blocking in 5% BSA for 2 hours at room temperature. Filters were then washed three times (10 minutes each) before incubation with primary antibody (diluted in TBS-T) overnight at 4°C. Filters were washed three times (10 minutes each) and then incubated with secondary antibody for 1 hour at room temperature. Filters were then washed four times and developed by ECL using Prime Western Blotting Detection Reagent (Amersham). Images were digitally acquired using a FUSION-SL Advance Imager (PeqLab) and modified for contrast using Adobe Photoshop v. CS4. All wash and incubation steps were performed with gentle shaking. The following antibodies were used: Rabbit polyclonal anti-DNAAF2 (1:1000; Atlas Antibodies; HPA004113), rabbit polyclonal anti-LRRC48 (1:500; Atlas Antibodies; HPA036040), rabbit polyclonal anti-DYX1C1 (1:1000; ProteinTech; 14522-1-AP); Rabbit polyclonal anti-myc (1:25; clone A-14, Santa Cruz), mouse monoclonal anti-myc (1:2000; clone A.7, Abcam), rabbit polyclonal control IgG (1:25; sc-2027, Santa Cruz), rabbit polyclonal anti-FLAG (1:250; clone F7425, Sigma Aldrich), mouse monoclonal anti-FLAG (1:2000; clone M2, Sigma Aldrich), goat anti-mouse HRP antibody (1:5000; NA931V, GE Healthcare) and goat anti-rabbit HRP antibody (1:3000; NA934, GE Healthcare).
To analyze the binding capacity between DYX1C1 and DNAAF2, plasmids expressing full-length DYX1C1 fused to a DNA-binding domain (GAL4-BD) and full-length DNAAF2 fused to an activation domain (GAL4-AD) were transformed in yeast strains PJ69-4A and PJ69-4α respectively, and subsequently combined by yeast mating and diploids containing both plasmids were selected on media lacking leucine and tryptophane. Interactions were analyzed by assessment of reporter gene activation via growth on media additionally lacking histidine and adenine to detect HIS3 and ADE2 reporter gene activation, α-galactosidase colorimetric plate assays (MEL1 reporter gene, not shown), and β-galactosidase colorimetric filter lift assays (LacZ reporter gene). As a positive control, the binding capacity of the known interactors BD-USH2A_icd and AD-NINL_isoB was assessed, and as a negative control the inability of BD-USH2A_icd to bind to only the GAL4 domain (AD-GAL4). Detailed protocols for evaluation protein-protein interactions are available from the authors upon request.
We thank the patients and families for participation in this study, the German patient support group "Kartagener Syndrom und Primaere Ciliaere Dyskinesie e.V." and the US PCD foundation.
This work was funded by the “Deutsche Forschungsgemeinschaft” (DFG Om 6/4), the IZKF Muenster (HO), by the European Community's Seventh Framework Programme FP7/2009, under grant agreement no: 241955, SYSCILIA (RR, HO), BESTCILIA under grant agreement no: 305404 (HO) and by the Netherlands Organization for Scientific research (NWO Vidi-91786396 and Vici-016.130.664) (RR) and by grants from the NIH, R01 HD055655, R01 MH056524; P01 HD057853 (JJL); research grant 5 U54 HL096458-06 (MRK, MWL, JLC, MAZ); NHLBI grant 5 R01HL071798 (MRK, MAZ); NICHD grant 2 R01HD048584 (CES, JVT, SC and RDB); NIH grant U01HL098180 (CWL); AHA fellowship (YL). Newlife Foundation grant 10/11/15 (HMM), Action Medical Research grant RTF1411 (MS). UK10K was funded by the Wellcome Trust (award WT091310). This work was supported partially US NIH grants UL1 TR000083 from the NCATS.
ZFIN Direct Data Submission: http://zfin.org (2004); For further information about the UK10K group: http://uk10k.org.uk; Complex congenital heart disease (CCHD) database: http://www.informatics.jax.org/javawi2/servlet/WIFetch?page=alleleDetail&id=MGI:531 1375
Database accession numbers
All cDNA clones were confirmed by sequence analysis and matched the following gene accession numbers: NM_130810.3 (DYX1C1), NM_178452.4 (DNAAF1/LRRC50), BC016843 (DNAAF3/C19orf51), NM_213607.1 (CCDC103), and NM_018139.2 (DNAAF2/KTU).
Authors Contributions J.L. and H.O. designed the study. Authors contributing to human DYX1C1: N.T.L., C.J., H.Ol., R.H., G.W.D., P.P., M.A., D.A.M., S.J.F.L., R.R., K.B. and J.R. performed the experiments and analysed the human data, G.K., C.W., J.R., M.G., M.J., H.O. provided clinical OP patient data; M.Z., L.C.M., A.J. S., J.L.C., M.W.L., W.E.W. and M.R.K. performed mutational analyses and provided the clinical data from UNC patients; UCL patient clinical ascertainment: J.S.L.; UCL mutation analysis: A.O., M.S., H.M.M.; deletion algorithm for UCL patients: V.P., UK’s exome sequencing: UK10K. Authors contributing to mouse Dyx1c1: A.T., B.S., M.C., N.T.L., P.P., M.A. performed the experiments and analysed the mouse data. Authors contributing to mouse Sharpei: R.F., Y.L., K.L., N.K., X.L., G.G., K.T. performed the experiments and analysed the mouse data. Authors contributing to zebrafish Dyx1c1: C.E.S., J.V.T., S.C., R.D.B. performed the experiments and analysed the zebrafish data. Authors contributing to Y2H experiments: D.A.M., S.J.F.L., R.R.
Author information The authors have no competing interests as defined by Nature Publishing Group, or other interests that might be perceived to influence the results and/or discussion reported in this article.