Dynein axonemal heavy chain 10 deficiency causes primary ciliary dyskinesia in humans and mice

Abstract

Primary ciliary dyskinesia (PCD) is a congenital, motile ciliopathy with pleiotropic symptoms. Although nearly 50 causative genes have been identified, they only account for approximately 70% of definitive PCD cases. Dynein axonemal heavy chain 10 (DNAH10) encodes a subunit of the inner arm dynein heavy chain in motile cilia and sperm flagella. Based on the common axoneme structure of motile cilia and sperm flagella, DNAH10 variants are likely to cause PCD. Using exome sequencing, we identified a novel DNAH10 homozygous variant (c.589C > T, p.R197W) in a patient with PCD from a consanguineous family. The patient manifested sinusitis, bronchiectasis, situs inversus, and asthenoteratozoospermia. Immunostaining analysis showed the absence of DNAH10 and DNALI1 in the respiratory cilia, and transmission electron microscopy revealed strikingly disordered axoneme 9+2 architecture and inner dynein arm defects in the respiratory cilia and sperm flagella. Subsequently, animal models of Dnah10-knockin mice harboring missense variants and Dnah10-knockout mice recapitulated the phenotypes of PCD, including chronic respiratory infection, male infertility, and hydrocephalus. To the best of our knowledge, this study is the first to report DNAH10 deficiency related to PCD in human and mouse models, which suggests that DNAH10 recessive mutation is causative of PCD.

Keywords

DNAH10 / mice / motile cilia / mutation / primary ciliary dyskinesia

1 Introduction

Primary ciliary dyskinesia (PCD, (MIM: 244400)) is a clinically and genetically heterogeneous motile ciliopathy, mostly following autosomal recessive or, less frequently, X-linked recessive or autosomal dominant inheritance [1,2]. Approximately 50% of PCD cases present Kartagener syndrome, characterized by chronic sinusitis, bronchiectasis, and situs inversus [3]. Clinically, patients with PCD can manifest subfertility due to defective sperm flagella in men or oviduct cilia in women [2]. Occasionally, hydrocephalus arises from cerebrospinal fluid flow dysfunction due to ependymal cilia dysmotility [2]. Comprehensive examinations are currently being applied to PCD diagnosis owing to the variability of the underlying pathogenesis of PCD [1,4]. Although approximately 50 pathogenic genes have been identified, they only account for approximately 70% of PCD cases [2].

Motile cilia and sperm flagella are evolutionarily conserved organelles sharing similar microtubule-based axoneme architecture in various species, comprising nine outer doublet microtubules, which encircle a central pair (i.e., 9+2 structure), except nodal cilia (9+0 arrangement) [5]. The inner dynein arms (IDAs) and outer dynein arms (ODAs), which are attached to the peripheral microtubules, hydrolyze adenosine triphosphate (ATP) to provide ciliary beating power and modulate beat frequency and pattern by interacting with the axonemal complex [2,6]. Dynein axonemal heavy chain 10 (DNAH10) is a subunit dynein heavy chain of IDA that is expressed in the motile cilia of various tissues, including airway epithelial ciliary cells, ventricular epithelial cilia, and sperm flagella [7,8]. Emerging evidence suggests that DNAH family genes may cause complex phenotypes in motile ciliopathies. Several DNAH family members coding for ODA components in motile cilia, such as DNAH5, DNAH9, and DNAH11, were considered related to PCD [9–11]. Moreover, mutations in DNAH1 and DNAH6, which encode IDA components, have been reported in PCD or isolated infertility [12–15]. Previous studies have reported that DNAH10 mutations can cause isolated infertility characterized by asthenoteratozoospermia in humans and mice [16,17] and heterotaxy or scoliosis in zebrafish [18,19]. Based on its essential role in IDA and its widespread expression in motile cilia, we hypothesize that DNAH10 may also be a candidate pathogenic gene for PCD [20].

In the present study, we first identified a novel homozygous variant (c.589C > T) of DNAH10 in a patient diagnosed with PCD with microtubular disorganization and IDA defects from a Chinese consanguineous family. Moreover, Dnah10-knockout male mice and Dnah10-knockin male mice harboring a homozygous missense variant (Dnah10^M/M), equivalent to that observed in the patient, were constructed using clustered regularly interspaced short palindromic repeats (CRISPR) and CRISPR-associated protein 9 (CRISPR-Cas9) technology. Both animal models with Dnah10 deficiency exhibited airway cilia and sperm flagellar ultrastructures and mobility deficiency and recapitulated the complex PCD clinical phenotypes, including hydrocephalus, chronic respiratory infection, and male infertility. These findings strongly suggest that DNAH10 deficiency contributes to PCD in humans and mice.

2 Materials and methods

2.1 Human subjects

A proband with PCD and his family members were recruited from the Second Xiangya Hospital of Central South University (Changsha, China). Clinical diagnostic assessment of PCD, including medical imaging and nasal nitric oxide (nNO) measurement, in addition to electron and immunofluorescence microscopy to analyze the ciliary structure, was performed. PCD diagnosis was established based on the European Respiratory Society guidelines [4]. This study was approved by the Institutional Ethics Committee of the Second Xiangya Hospital of Central South University (Changsha, China). Written informed consent was obtained from all the patients and healthy controls.

2.2 Exome sequencing and bioinformatics analysis

Peripheral venous blood samples were collected from the patient and his family. Genomic DNA was extracted from the patient using the DNeasy Blood & Tissue Kit (51104, QIAGEN) as directed and used for subsequent exon sequencing. Library capture, sequencing, and data analysis were performed by Novogene Bioinformatics Institute, Beijing, China as described previously [21]. Briefly, the genomic DNA of the patient was captured using the Agilent SureSelect Human All Exon V6 Kit (G9706K, Agilent Technologies) and sequenced on an Illumina HiSeq 4000. After quality control, the sequencing reads were aligned to the National Center for Biotechnology Information human reference genome (GRCh37/hg19) using Burrows–Wheeler Aligner. ANNOVAR was used to annotate variant call format files.

Single-nucleotide polymorphisms (SNPs) and short insertion–deletions (indels) were filtered as follows: (1) variants with a minor allele frequency below 1% in the 1000 Genomes Project, Genome Aggregation Database (gnomAD), and in-house database of Novogene were assessed; (2) noncoding and intronic variants were excluded; (3) synonymous missense variants were filtered; (4) homozygous variants from the homozygous regions identified using the Automap algorithm were retained [22]; and (5) a PCD or PCD-candidate gene panel derived from the literature was used to identify disease-related variants [2,20].

2.3 Sanger sequencing

Sanger sequencing was performed in the patient to validate the variants. Primer sequences were designed using an online primer design tool. The primer sequences for the Sanger sequencing of humans and mice are displayed in Table S1.

2.4 Immunostaining

Bronchial epithelial tissues from the proband and a healthy control were obtained by fiber bronchoscopy and fixed in 4% paraformaldehyde overnight at 4 °C. Immunostaining on the slides was performed as described previously [23]. Briefly, the slides were incubated overnight at 4 °C with primary antibodies: DNAH10 (HPA039066, Sigma-Aldrich, 1:100), DNAH5 (HPA037470, Sigma-Aldrich, 1:100), DNALI1 (ab155490, 1:100, Abcam, UK), and anti-α-tubulin (T9026, Sigma-Aldrich, 1:500). Antibody binding was detected using Alexa Fluor 488 anti-mouse IgG (ab150113, Abcam, 1:500, UK) and Alexa Fluor 555 anti-rabbit IgG (A31572, Invitrogen, 1:500). The slides were incubated for 2 h at 37 °C and stained with 2-(4-amidinophenyl)-1H-indole-6-carboxamidine for 5 min at room temperature. Fluorescence signals were photographed using an Olympus BX53 fluorescence microscope and analyzed using the cellSens Dimension software (Olympus BX53).

2.5 Semen and sperm morphological analysis

Semen samples were obtained from the proband and a healthy control after 5–7 days of sexual abstinence and from Dnah10-knockin mice. Three independent semen analyses were performed according to the World Health Organization (WHO) guidelines [24]. Morphological defects of the flagella were classified as absent, short, bent, coiled, or irregular in width as determined by Papanicolaou staining [25]. The percentages of morphologically normal and abnormal spermatozoa were calculated according to the WHO guidelines [26].

2.6 Mouse models

Dnah10^M/M and Dnah10-knockout mice were constructed using the CRISPR-Cas9 system using previously published methods [27]. We designed a single-guide RNA (sgRNA-1: GATTCATTAAGAACTCATCGCGG) to target exon 6 of Dnah10 with a mutation (c.778CGC > TGG) equivalent to that observed in our patient (c.589C > T) to generate Dnah10^M/M mice. Dnah10-knockout mice against exons 2–5 were reported in our previous study [16].

We transcribed Cas9 mRNA and sgRNAs using T7 RNA polymerase in vitro and then mixed and co-microinjected them into fertilized oocytes of C57BL/6J mice. The founder animals and their offspring were genotyped via polymerase chain reaction (PCR) and Sanger sequencing of the tail genomic DNA using the specific primers listed in Table S1. All animal procedures were conducted according to the protocols established by the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health and the Institutional Animal Care and Use Committee of Central South University.

2.7 Transmission electron microscopy (TEM)

Respiratory cilia and spermatozoa from human and mice were treated as described previously [16,28]. Briefly, respiratory cilia or spermatozoa were fixed with glutaraldehyde (G1102, Servicebio) and osmium tetroxide, post-fixed with sucrose and osmium oxide, and dehydrated with gradient ethanol. Subsequently, 80 nm-thick sections were contrasted with lead citrate and uranyl acetate. Finally, digital images were captured using an HT7700 Hitachi electron microscope (HT7700, Hitachi) and a MegaView III digital camera (Olympus Optical).

2.8 High-speed video microscopy (HSVM) analysis

Tracheal ciliated epithelia from mice for HSVM analysis were photographed using an upright Olympus BX53 microscope (BX53, Olympus) with a 40× objective lens. Videos were recorded using a semiconductor camera (Prime BSI™, Photometrics) at a rate of 500 fps at 37 °C. Only intact ciliated edges ( > 50 μm) were used for functional analysis. Five separate ciliated epithelial strips from the mucus-free regions were measured. Ciliary beat frequency (CBF) was calculated using a validated automated open-source software (CiliarMove version 1, Microsoft).

2.9 Real-time quantitative polymerase chain reaction (RT-qPCR)

Total RNAs from the trachea and testes of adult Dnah10-knockin mice and Dnah10-knockout mice were extracted using an RNA extraction and purification kit (K0731, Thermo Fisher Scientific) for RT-qPCR. Approximately 1 µg RNA was reverse-transcribed into cDNA using EZscript. A reverse-transcription mix (EZB-RT2GQ, EZBioscience) was used according to the manufacturer’s instructions. The target mRNA expression levels were normalized to those of the mouse Gapdh primers using the primers listed in Table S2.

2.10 Statistical analyses

Statistical analyses were performed using the SPSS version 17.0 and GraphPad Prism version 8.4.0. Descriptive statistics of the variables are presented as mean ± standard deviation or counts (n) and proportions (%). Chi-square test was used for categorical data. Student’s t-test was used for continuous variables. The significance level was set at P < 0.05.

3 Results

3.1 Clinical assessment

A 28-year-old man from a consanguineous family presented with complaints of recurrent cough, purulent sputum, and nasal congestion since childhood (Fig.1). His early deceased sister was diagnosed with Kartagener syndrome (including chronic sinusitis, bronchiectasis, and situs inversus). The proband had no history of trauma, poliomyelitis, tuberculosis, or toxic exposure. High-resolution computed tomography of the paranasal sinuses, chest, and abdomen of the proband revealed chronic rhinosinusitis (Fig.1), bronchiectasis, dextrocardia (Fig.1), and abdominal situs inversus (Fig.1). The nNO production rate of the proband was 24 nL/min, which is far below the reference nNO cutoff value of PCD (77 nL/min) [29]. TEM analysis of respiratory cilia revealed disorganized 9+2 structure and IDA defects (Fig.1–1G). Routine semen analysis of the spermatozoa of the proband revealed severe asthenoteratozoospermia with completely immotile spermatozoa and a comparatively low sperm count. Papanicolaou staining of the proband revealed that most spermatozoa manifested absent, short, bent, and coiled flagella (Fig.1, Table S3). Similarly, TEM results showed a highly disorganized 9+2 structure and IDA defects in the mutant sperm flagella of the proband (Fig.1–1K). In summary, the proband presented complex PCD phenotypes, including sinusitis, bronchiectasis, situs inversus, and asthenoteratozoospermia.

3.2 Identification of a novel homozygous variant of DNAH10 in a patient with PCD

Whole exome sequencing was performed to identify the potential genetic source of the PCD. In accordance with our established protocol for investigating potential genes associated with PCD, we detected a homozygous missense variant of DNAH10 (NC_000012.12:g.123781230C > T, NM_207437.3:c.589C > T, NP_997320.2:p. R197W) in the proband (IV-4) (Fig.1). Sanger sequencing verified that the proband carried a homozygous variant, whereas the unaffected parents had a heterozygous variant (Fig.2). Homozygosity mapping confirmed that the homozygous region in chromosome 12 contained a variant of DNAH10 in the proband (Fig. S1). Homozygosity mapping showed that the largest homozygous fragment contained the PCD pathogenic genes DNAH5 and CCNO, but none of these two gene variants was found in the whole exome sequencing data. Homozygous fragments of other chromosomes were found to have known PCD causative genes, but no homozygous SNP or indel variants were found in any of them (Tables S4 and S5). The DNAH10 variant was rare in public human databases, including the 1000 Genomes Project and gnomAD (Table S6). The DNAH10 missense variant was also predicted to be disease causing using the Sorting Intolerant From Tolerant, PolyPhen-2, Protein Variation Effect Analyzer, and Combined Annotation Dependent Depletion tools (Table S6).

The p.R197W variant is located in the N-terminal of DNAH10 (Fig.2), which is responsible for dimerizing with other dynein proteins and connecting to the intermediated chain. Notably, the affected DNAH10 residues (R197) were conserved among different species (Fig.2). Modeling using SWISS-MODEL software suggested that R197W mutations in DNAH10 affected intramolecular hydrogen bond formation, which is predicted to affect the hydrophobicity and stability of the alpha helix (Fig.2). Collectively, these data suggest that DNAH10 is an excellent candidate to explain the PCD phenotype of the patient.

3.3 DNAH10 loss in airway cilia in the patient with DNAH10 homozygous variant

The presence and localization of several proteins belonging to different substructures of the axoneme, including DNAH10 and accepted markers of ODA (DNAH5) and IDA (DNALI1) were determined to further explore the ultrastructural defects of respiratory cilia revealed by TEM. Interestingly, compared with normal epithelial cells, in which DNAH10 staining was specifically localized along the ciliary axoneme, DNAH10 staining was absent in the respiratory cilia of the proband (Fig.3 and S2). DNAH5 was intact, whereas DNALI1 was absent in the respiratory epithelial cells of the proband (Fig.3 and 3C).

3.4 PCD phenotype in Dnah10^M/M mice

A Dnah10^M/M-knockin mouse model harboring a homozygous Dnah10 variant (c.778CGC > TGG, p.R260W) that corresponded to the PCD-affected proband (c.589C > T, p.R197W) was constructed using CRISPR-Cas9 technology to demonstrate whether the DNAH10 variant in the patient with PCD was pathogenic (Fig. S3A). The designed mutation site was confirmed in mouse genomic DNA using PCR amplification and Sanger sequencing (Fig. S3B). Dnah10 mRNA was also nearly absent in the trachea and testes of Dnah10^M/M mice as detected by RT-qPCR (Fig. S4A and S4B).

Approximately 7% (3/40) of Dnah10^M/M mice presented growth retardation and head enlargement (Fig.4). Evident hydrocephalus signs were suggested by the hematoxylin and eosin (H&E) staining of the brains of 5-week-old Dnah10^M/M mice (Fig.4).

We observed chronic lung infection in Dnah10^M/M mice by H&E staining, which manifested as lymphocyte and plasma cell aggregation in the lung stroma and pulmonary interstitial hyperplasia when compared with Dnah10^+/M mice (Fig.4). Ultrastructural examination of the airway epithelium cilia by TEM revealed microtubular disorganization and IDA defect of the ciliary axoneme, which was similar to that in the affected man (Fig.4–4F). Half of the visual fields of the cilia in the Dnah10^M/M mice were totally static or swing in residue pattern (Fig.4), whereas half of the other fields (10/20) remained in subtle reduced beat (CBF=11.0±0.1 Hz) when compared with those in the Dnah10^+/M mice (CBF=12.7±0.3 Hz, Fig.4).

Simultaneously, we observed infertility in Dnah10^M/M male mice (Fig.5), whereas Dnah10^M/M female mice gave birth normally. Similar testicular weights were observed in the Dnah10^M/M and Dnah10^+/M mice (Fig.5). Using computer-aided sperm analysis, we found that the sperm count in Dnah10^M/M mice was remarkably lower compared with that in Dnah10^+/M mice (Fig.5), and the sperm was completely immotile in Dnah10^M/M male mice (Fig.5). The sperm flagella of Dnah10^M/M male mice exhibited multiple morphological abnormalities, including absent, short, coiled, and irregular flagella (Fig.5, Table S7). TEM results showed a disorganized 9+2 axoneme structure and mitochondrial sheath and dense outer fibers in Dnah10^M/M male mice (Fig.5–5H). Additionally, laterality defects were not observed in the Dnah10^M/M mice. Together, the similar PCD profiles observed in Dnah10^M/M mice further indicated that the DNAH10 variant (c.589C > T) was pathogenic for PCD.

3.5 Dnah10-knockout mice recapitulates PCD phenotype in human

The methods used to construct the Dnah10-knockout mice were described in our previous report, and the Dnah10-knockout mice showed evident asthenoteratozoospermia similar to that in humans [16]. We expanded mouse breeding to perform follow-up observations of the phenotypes of Dnah10-knockout mice from our previous study. We found that approximately 10% (5/50) of Dnah10^−/− mice exhibited growth retardation and head enlargement (Fig.6). Evident hydrocephalus signs were suggested by the H&E staining of the brains of Dnah10^−/− mice aged 8 weeks (Fig.6). Laterality defects were not observed in Dnah10^−/− mice.

RT-qPCR of Dnah10 showed that Dnah10 expression was considerably abolished in the trachea and testes of Dnah10^−/− mice (Fig. S5A and S5B). H&E staining of lung tissues revealed that Dnah10^−/− mice showed signs of chronic lung inflammation characterized by lymphocyte and plasma cell-dominated chronic infiltration in the lung stroma and mucus accumulation in small airways (Fig.6). TEM results showed that the cross sections of the axoneme in Dnah10^−/− male mice revealed a disorganization of the 9+2 structure and IDA defects (Fig.6–6F). Compared with the normal CBF (12.3±0.3 Hz) of Dnah10^+/− mice via HSVM analysis (Fig.6), the cilia of Dnah10^−/− mice were totally static or with residual motility in 80% (16/20) of the visual fields and beat slowly (CBF=6±0.5 Hz, Fig.6) in a stiff manner in the other 20% (4/20) of the observed fields. Together, these data indicate that Dnah10 deficiency can cause a PCD phenotype in mice, which is similar to that of the affected man.

4 Discussion

To the best of our knowledge, this study is the first to report DNAH10 deficiency in PCD, characterized by recurrent respiratory infection, laterality defect, and male infertility. Dnah10-knockout mice and Dnah10-knockin mice resembled the features of PCD in the man in this study.

Motile cilia are widely distributed in the respiratory epithelium, ependyma, sperm flagella, oviduct, and embryonic nodes, which provide the driving force for epithelial fluid and nodal flow and sperm flagellar movement [2,5]. DNAH family members encode 13 macromolecular motor proteins of IDA (DNAH1, DNAH2, DNAH3, DNAH6, DNAH7, DNAH10, DNAH12, and DNAH14) and ODA (DNAH5, DNAH8, DNAH9, DNAH11, and DNAH17) that hydrolyze ATP and release energy to maintain ciliary beating in a regular pattern and specific frequency. The absence of only one of the many dynein isoforms in human cilia has been associated with pleiotropic symptoms due to defects in ciliary motility [5]. Previous studies have shown that DNAH5, DNAH9, and DNAH11 are PCD-associated genes [10,11,30]. DNAH1 and DNAH6 variants have been reported to result in asthenoteratozoospermia, with or without the manifestation of PCD symptoms [12–15]. These observations indicate that dynein isoforms play important roles in ciliary and flagellar functions.

Recently, DNAH10 has been recognized as an isolated asthenoteratozoospermia gene [16,17]. However, in our study, the patient carrying the DNAH10 homozygous variant and Dnah10-mutant animal models presented with typical PCD symptoms. These phenotypic differences may be due to population heterogeneity or differences in environmental exposure. Interestingly, similar differences between PCD and isolated infertility were observed for other PCD causative genes or isolated asthenoteratozoospermia genes, such as DNAH1 [12,13], DNAH6 [14,15], SPEF2 [31,32], and CFAP43 [33,34]. These different phenotypes may contribute to testis-specific isoforms or homologs between tissues, which may prevent the effect of specific mutations on spermatogenesis or ciliogenesis [35,36]. Therefore, the exact mechanisms involved in flagellar and ciliary functions require extensive investigation, which is a necessity in the counseling of patients about their PCD symptoms and men carrying the DNAH10 variant.

Dynein assembly involves complex processes, such as cytoplasmic preassembly through the aid of chaperoning complexes, transportation into the ciliary/flagellar compartment through interactions with intraflagellar transport (IFT) machinery, and anchoring onto doublets through the aid of proteins that form high-affinity docking sites [6,37]. In our study, we found an immotile or retarded beating frequency with a stiff pattern in the respiratory cilia of Dnah10-mutant mice. Accordingly, ultrastructure examinations revealed that DNAH10 deficiency led to IDA loss and axoneme disarrangement, and immunostaining revealed that DNALI1 (IDA subunit a–c markers) was also lost in the respiratory cilia, suggesting that DNAH10 may participate in multiple processes as a component of cytoplasmic preassembly, as a cargo of IFT, or as a cooperator with the docking complex. Similar ultrastructural deformities can be observed in mutations of ruler protein genes CCDC39 and CCDC40 [38–40], which display attachment sites for IDA and nexin–dynein regulatory complexes, and in other IDA-coding genes, such as DNAH1 [13] and DNAH6 [14]. Therefore, exploring the pathogenesis of DNAH10 in ciliogenesis and spermatogenesis is worthy of further studies.

Our study found that a small proportion of mice with Dnah10 deficiency developed hydrocephalus, a severe consequence of cerebrospinal fluid circulation disorders [41]. Several PCD pathogenic genes demonstrated hydrocephalus phenotypes in humans or mice, such as genes for ciliogenesis (FOXJ1, CCNO, MCIDAS), ODA components (DNAH5), and cilia- and flagella-associated protein (CFAP43) [30,34,42–44]. Hydrocephalus is more common in animal models of PCD than in humans. For example, hydrocephalus was found in homozygous mouse models with Dnah5 mutation, but the incidence rate of hydrocephalus in patients with PCD was only 2.5% (2/80), which may be related to the wider and shorter aqueduct of the midbrain in humans than in mice [45]. This difference in humans and mice was also observed in the present study. Our findings suggest that DNAH10 defects may affect ependymal motile cilia and consequently cause hydrocephalus, expanding the clinical spectrum of PCD.

Mice are the most common mammalian model because of their high similarity in genome, anatomy, and physiology with humans [46]. Dnah10-knockin and Dnah10-knockout mice recapitulated the manifestation of chronic respiratory infection and asthenoteratozoospermia, but they did not develop laterality defects, implying variable phenotypes in different species. Additionally, laterality defects in the mouse progeny was difficult to completely observe, which maybe due to the randomization of organ arrangement and the underlying compensation mechanism [47,48].

Our study has some limitations. First, we could not assess the ciliary motility of the proband owing to the patient’s refusal. We hope to analyze the CBF and ciliary beat pattern in a large cohort of patients with DNAH10 deficiency to describe its ciliary beating profile by HSVM analysis in the future. Second, we did not show the absence of DNAH10 at the protein level in mouse models, because a stable and ideal commercial anti-DNAH10 antibody for mice was unavailable. However, we tested Danh10 variants at the DNA and RNA levels, which could demonstrate the expression deficiency of DNAH10 to some extent. Moreover, PCD is a complex genetic disorder without a definite clinical cure strategy; thus, only symptomatic treatment is provided to patients with PCD. Gene editing through airway organoid construction may bring new treatment strategies for patients with PCD in the future [49,50].

In summary, we identified for the first time a novel DNAH10 homozygous variant in a patient with PCD. Genetic evidence from DNAH10-associated patient and Dnah10-knockin and Dnah10-knockout mice strongly suggests that biallelic DNAH10 variants can induce multisystem motile ciliopathies, such as PCD, which is characterized by chronic respiratory infection and infertility due to ultrastructural and functional abnormalities of the cilia and sperm flagella. Our clinical and genetic findings provide a candidate gene for the molecular diagnosis of PCD and extend our knowledge of the pathogenic effects of DNAH10 mutations.

References

[1] Lucas JS, Davis SD, Omran H, Shoemark A. Primary ciliary dyskinesia in the genomics age. Lancet Respir Med 2020; 8(2): 202–216

[2] Wallmeier J, Nielsen KG, Kuehni CE, Lucas JS, Leigh MW, Zariwala MA, Omran H. Motile ciliopathies. Nat Rev Dis Primers 2020; 6(1): 77

[3] Goutaki M, Meier AB, Halbeisen FS, Lucas JS, Dell SD, Maurer E, Casaulta C, Jurca M, Spycher BD, Kuehni CE. Clinical manifestations in primary ciliary dyskinesia: systematic review and meta-analysis. Eur Respir J 2016; 48(4): 1081–1095

[4] Lucas JS, Barbato A, Collins SA, Goutaki M, Behan L, Caudri D, Dell S, Eber E, Escudier E, Hirst RA, Hogg C, Jorissen M, Latzin P, Legendre M, Leigh MW, Midulla F, Nielsen KG, Omran H, Papon JF, Pohunek P, Redfern B, Rigau D, Rindlisbacher B, Santamaria F, Shoemark A, Snijders D, Tonia T, Titieni A, Walker WT, Werner C, Bush A, Kuehni CE. European Respiratory Society guidelines for the diagnosis of primary ciliary dyskinesia. Eur Respir J 2017; 49(1): 1601090

[5] Fliegauf M, Benzing T, Omran H. When cilia go bad: cilia defects and ciliopathies. Nat Rev Mol Cell Biol 2007; 8(11): 880–893

[6] KingSM. Dyneins: Dynein Mechanics, Dysfunction, and Disease. Saint Louis: Elsevier Science & Technology, 2017

[7] Maiti AK, Mattéi MG, Jorissen M, Volz A, Zeigler A, Bouvagnet P. Identification, tissue specific expression, and chromosomal localisation of several human dynein heavy chain genes. Eur J Hum Genet 2000; 8(12): 923–932

[8] Fagerberg L, Hallström BM, Oksvold P, Kampf C, Djureinovic D, Odeberg J, Habuka M, Tahmasebpoor S, Danielsson A, Edlund K, Asplund A, Sjöstedt E, Lundberg E, Szigyarto CA, Skogs M, Takanen JO, Berling H, Tegel H, Mulder J, Nilsson P, Schwenk JM, Lindskog C, Danielsson F, Mardinoglu A, Sivertsson A, von Feilitzen K, Forsberg M, Zwahlen M, Olsson I, Navani S, Huss M, Nielsen J, Ponten F, Uhlén M. Analysis of the human tissue-specific expression by genome-wide integration of transcriptomics and antibody-based proteomics. Mol Cell Proteomics 2014; 13(2): 397–406

[9] Olbrich H, Häffner K, Kispert A, Völkel A, Volz A, Sasmaz G, Reinhardt R, Hennig S, Lehrach H, Konietzko N, Zariwala M, Noone PG, Knowles M, Mitchison HM, Meeks M, Chung EM, Hildebrandt F, Sudbrak R, Omran H. Mutations in DNAH5 cause primary ciliary dyskinesia and randomization of left-right asymmetry. Nat Genet 2002; 30(2): 143–144

[10] Fassad MR, Shoemark A, Legendre M, Hirst RA, Koll F, le Borgne P, Louis B, Daudvohra F, Patel MP, Thomas L, Dixon M, Burgoyne T, Hayes J, Nicholson AG, Cullup T, Jenkins L, Carr SB, Aurora P, Lemullois M, Aubusson-Fleury A, Papon JF, O’Callaghan C, Amselem S, Hogg C, Escudier E, Tassin AM, Mitchison HM. Mutations in outer dynein arm heavy chain DNAH9 cause motile cilia defects and situs inversus. Am J Hum Genet 2018; 103(6): 984–994

[11] Pifferi M, Michelucci A, Conidi ME, Cangiotti AM, Simi P, Macchia P, Boner AL. New DNAH11 mutations in primary ciliary dyskinesia with normal axonemal ultrastructure. Eur Respir J 2010; 35(6): 1413–1416

[12] Ben Khelifa M, Coutton C, Zouari R, Karaouzène T, Rendu J, Bidart M, Yassine S, Pierre V, Delaroche J, Hennebicq S, Grunwald D, Escalier D, Pernet-Gallay K, Jouk PS, Thierry-Mieg N, Touré A, Arnoult C, Ray PF. Mutations in DNAH1, which encodes an inner arm heavy chain dynein, lead to male infertility from multiple morphological abnormalities of the sperm flagella. Am J Hum Genet 2014; 94(1): 95–104

[13] Imtiaz F, Allam R, Ramzan K, Al-Sayed M. Variation in DNAH1 may contribute to primary ciliary dyskinesia. BMC Med Genet 2015; 16(1): 14

[14] Li Y, Yagi H, Onuoha EO, Damerla RR, Francis R, Furutani Y, Tariq M, King SM, Hendricks G, Cui C, Saydmohammed M, Lee DM, Zahid M, Sami I, Leatherbury L, Pazour GJ, Ware SM, Nakanishi T, Goldmuntz E, Tsang M, Lo CW. DNAH6 and its interactions with PCD genes in heterotaxy and primary ciliary dyskinesia. PLoS Genet 2016; 12(2): e1005821

[15] Tu C, Nie H, Meng L, Yuan S, He W, Luo A, Li H, Li W, Du J, Lu G, Lin G, Tan YQ. Identification of DNAH6 mutations in infertile men with multiple morphological abnormalities of the sperm flagella. Sci Rep 2019; 9(1): 15864

[16] Tu C, Cong J, Zhang Q, He X, Zheng R, Yang X, Gao Y, Wu H, Lv M, Gu Y, Lu S, Liu C, Tian S, Meng L, Wang W, Tan C, Nie H, Li D, Zhang H, Gong F, Hu L, Lu G, Xu W, Lin G, Zhang F, Cao Y, Tan YQ. Bi-allelic mutations of DNAH10 cause primary male infertility with asthenoteratozoospermia in humans and mice. Am J Hum Genet 2021; 108(8): 1466–1477

[17] Li K, Wang G, Lv M, Wang J, Gao Y, Tang F, Xu C, Yang W, Yu H, Shao Z, Geng H, Tan Q, Shen Q, Tang D, Ni X, Wang T, Song B, Wu H, Huo R, Zhang Z, Xu Y, Zhou P, Tao F, Wei Z, He X, Cao Y. Bi-allelic variants in DNAH10 cause asthenoteratozoospermia and male infertility. J Assist Reprod Genet 2022; 39(1): 251–259

[18] Liu C, Cao R, Xu Y, Li T, Li F, Chen S, Xu R, Sun K. Rare copy number variants analysis identifies novel candidate genes in heterotaxy syndrome patients with congenital heart defects. Genome Med 2018; 10(1): 40

[19] Wang Y, Troutwine BR, Zhang H, Gray RS. The axonemal dynein heavy chain 10 gene is essential for monocilia motility and spine alignment in zebrafish. Dev Biol 2022; 482: 82–90

[20] Paff T, Kooi IE, Moutaouakil Y, Riesebos E, Sistermans EA, Daniels HJMA, Weiss JMM, Niessen HHWM, Haarman EG, Pals G, Micha D. Diagnostic yield of a targeted gene panel in primary ciliary dyskinesia patients. Hum Mutat 2018; 39(5): 653–665

[21] Guo T, Tu CF, Yang DH, Ding SZ, Lei C, Wang RC, Liu L, Kang X, Shen XQ, Yang YF, Tan ZP, Tan YQ, Luo H. Bi-allelic BRWD1 variants cause male infertility with asthenoteratozoospermia and likely primary ciliary dyskinesia. Hum Genet 2021; 140(5): 761–773

[22] Quinodoz M, Peter VG, Bedoni N, Royer Bertrand B, Cisarova K, Salmaninejad A, Sepahi N, Rodrigues R, Piran M, Mojarrad M, Pasdar A, Ghanbari Asad A, Sousa AB, Coutinho Santos L, Superti-Furga A, Rivolta C. AutoMap is a high performance homozygosity mapping tool using next-generation sequencing data. Nat Commun 2021; 12(1): 518

[23] Dai C, Hu L, Gong F, Tan Y, Cai S, Zhang S, Dai J, Lu C, Chen J, Chen Y, Lu G, Du J, Lin G. ZP2 pathogenic variants cause in vitro fertilization failure and female infertility. Genet Med 2019; 21(2): 431–440

[24] Cooper TG, Noonan E, von Eckardstein S, Auger J, Baker HW, Behre HM, Haugen TB, Kruger T, Wang C, Mbizvo MT, Vogelsong KM. World Health Organization reference values for human semen characteristics. Hum Reprod Update 2010; 16(3): 231–245

[25] Johanisson E, Campana A, Luthi R, de Agostini A. Evaluation of “round cells” in semen analysis: a comparative study. Hum Reprod Update 2000; 6(4): 404–412

[26] Sanchez-Alvarez J, Cano-Corres R, Fuentes-Arderiu X. A complement for the WHO Laboratory Manual for the Examination and Processing of Human Semen (First Edition, 2010). EJIFCC 2012; 23(3): 103–106

[27] Wang H, Yang H, Shivalila CS, Dawlaty MM, Cheng AW, Zhang F, Jaenisch R. One-step generation of mice carrying mutations in multiple genes by CRISPR/Cas-mediated genome engineering. Cell 2013; 153(4): 910–918

[28] Francis R, Lo C. Ex vivo method for high resolution imaging of cilia motility in rodent airway epithelia. J Vis Exp 2013; 8(78): 50343

[29] Leigh MW, Hazucha MJ, Chawla KK, Baker BR, Shapiro AJ, Brown DE, Lavange LM, Horton BJ, Qaqish B, Carson JL, Davis SD, Dell SD, Ferkol TW, Atkinson JJ, Olivier KN, Sagel SD, Rosenfeld M, Milla C, Lee HS, Krischer J, Zariwala MA, Knowles MR. Standardizing nasal nitric oxide measurement as a test for primary ciliary dyskinesia. Ann Am Thorac Soc 2013; 10(6): 574–581

[30] Ibañez-Tallon I, Gorokhova S, Heintz N. Loss of function of axonemal dynein Mdnah5 causes primary ciliary dyskinesia and hydrocephalus. Hum Mol Genet 2002; 11(6): 715–721

[31] Liu C, Lv M, He X, Zhu Y, Amiri-Yekta A, Li W, Wu H, Kherraf ZE, Liu W, Zhang J, Tan Q, Tang S, Zhu YJ, Zhong Y, Li C, Tian S, Zhang Z, Jin L, Ray P, Zhang F, Cao Y. Homozygous mutations in SPEF2 induce multiple morphological abnormalities of the sperm flagella and male infertility. J Med Genet 2020; 57(1): 31–37

[32] Tu C, Nie H, Meng L, Wang W, Li H, Yuan S, Cheng D, He W, Liu G, Du J, Gong F, Lu G, Lin G, Zhang Q, Tan YQ. Novel mutations in SPEF2 causing different defects between flagella and cilia bridge: the phenotypic link between MMAF and PCD. Hum Genet 2020; 139(2): 257–271

[33] Coutton C, Vargas AS, Amiri-Yekta A, Kherraf ZE, Ben Mustapha SF, Le Tanno P, Wambergue-Legrand C, Karaouzène T, Martinez G, Crouzy S, Daneshipour A, Hosseini SH, Mitchell V, Halouani L, Marrakchi O, Makni M, Latrous H, Kharouf M, Deleuze JF, Boland A, Hennebicq S, Satre V, Jouk PS, Thierry-Mieg N, Conne B, Dacheux D, Landrein N, Schmitt A, Stouvenel L, Lorès P, El Khouri E, Bottari SP, Fauré J, Wolf JP, Pernet-Gallay K, Escoffier J, Gourabi H, Robinson DR, Nef S, Dulioust E, Zouari R, Bonhivers M, Touré A, Arnoult C, Ray PF. Mutations in CFAP43 and CFAP44 cause male infertility and flagellum defects in Trypanosoma and human. Nat Commun 2018; 9(1): 686

[34] Morimoto Y, Yoshida S, Kinoshita A, Satoh C, Mishima H, Yamaguchi N, Matsuda K, Sakaguchi M, Tanaka T, Komohara Y, Imamura A, Ozawa H, Nakashima M, Kurotaki N, Kishino T, Yoshiura KI, Ono S. Nonsense mutation in CFAP43 causes normal-pressure hydrocephalus with ciliary abnormalities. Neurology 2019; 92(20): e2364–e2374

[35] Sironen A, Shoemark A, Patel M, Loebinger MR, Mitchison HM. Sperm defects in primary ciliary dyskinesia and related causes of male infertility. Cell Mol Life Sci 2020; 77(11): 2029–2048

[36] Onoufriadis A, Paff T, Antony D, Shoemark A, Micha D, Kuyt B, Schmidts M, Petridi S, Dankert-Roelse JE, Haarman EG, Daniels JM, Emes RD, Wilson R, Hogg C, Scambler PJ, Chung EM; UK10K; Pals G, Mitchison HM. Splice-site mutations in the axonemal outer dynein arm docking complex gene CCDC114 cause primary ciliary dyskinesia. Am J Hum Genet 2013; 92(1): 88–98

[37] Jain R, Pan J, Driscoll JA, Wisner JW, Huang T, Gunsten SP, You Y, Brody SL. Temporal relationship between primary and motile ciliogenesis in airway epithelial cells. Am J Respir Cell Mol Biol 2010; 43(6): 731–739

[38] Merveille AC, Davis EE, Becker-Heck A, Legendre M, Amirav I, Bataille G, Belmont J, Beydon N, Billen F, Clément A, Clercx C, Coste A, Crosbie R, de Blic J, Deleuze S, Duquesnoy P, Escalier D, Escudier E, Fliegauf M, Horvath J, Hill K, Jorissen M, Just J, Kispert A, Lathrop M, Loges NT, Marthin JK, Momozawa Y, Montantin G, Nielsen KG, Olbrich H, Papon JF, Rayet I, Roger G, Schmidts M, Tenreiro H, Towbin JA, Zelenika D, Zentgraf H, Georges M, Lequarré AS, Katsanis N, Omran H, Amselem S. CCDC39 is required for assembly of inner dynein arms and the dynein regulatory complex and for normal ciliary motility in humans and dogs. Nat Genet 2011; 43(1): 72–78

[39] Becker-Heck A, Zohn IE, Okabe N, Pollock A, Lenhart KB, Sullivan-Brown J, McSheene J, Loges NT, Olbrich H, Haeffner K, Fliegauf M, Horvath J, Reinhardt R, Nielsen KG, Marthin JK, Baktai G, Anderson KV, Geisler R, Niswander L, Omran H, Burdine RD. The coiled-coil domain containing protein CCDC40 is essential for motile cilia function and left-right axis formation. Nat Genet 2011; 43(1): 79–84

[40] Antony D, Becker-Heck A, Zariwala MA, Schmidts M, Onoufriadis A, Forouhan M, Wilson R, Taylor-Cox T, Dewar A, Jackson C, Goggin P, Loges NT, Olbrich H, Jaspers M, Jorissen M, Leigh MW, Wolf WE, Daniels ML, Noone PG, Ferkol TW, Sagel SD, Rosenfeld M, Rutman A, Dixit A, O’Callaghan C, Lucas JS, Hogg C, Scambler PJ, Emes RD; Uk10k; Chung EM, Shoemark A, Knowles MR, Omran H, Mitchison HM. Mutations in CCDC39 and CCDC40 are the major cause of primary ciliary dyskinesia with axonemal disorganization and absent inner dynein arms. Hum Mutat 2013; 34(3): 462–472

[41] Kumar V, Umair Z, Kumar S, Goutam RS, Park S, Kim J. The regulatory roles of motile cilia in CSF circulation and hydrocephalus. Fluids Barriers CNS 2021; 18(1): 31

[42] Wallmeier J, Frank D, Shoemark A, Nöthe-Menchen T, Cindric S, Olbrich H, Loges NT, Aprea I, Dougherty GW, Pennekamp P, Kaiser T, Mitchison HM, Hogg C, Carr SB, Zariwala MA, Ferkol T, Leigh MW, Davis SD, Atkinson J, Dutcher SK, Knowles MR, Thiele H, Altmüller J, Krenz H, Wöste M, Brentrup A, Ahrens F, Vogelberg C, Morris-Rosendahl DJ, Omran H. De novo mutations in FOXJ1 result in a motile ciliopathy with hydrocephalus and randomization of left/right body asymmetry. Am J Hum Genet 2019; 105(5): 1030–1039

[43] Núnez-Ollé M, Jung C, Terré B, Balsiger NA, Plata C, Roset R, Pardo-Pastor C, Garrido M, Rojas S, Alameda F, Lloreta J, Martín-Caballero J, Flores JM, Stracker TH, Valverde MA, Muñoz FJ, Gil-Gómez G. Constitutive Cyclin O deficiency results in penetrant hydrocephalus, impaired growth and infertility. Oncotarget 2017; 8(59): 99261–99273

[44] Robson EA, Dixon L, Causon L, Dawes W, Benenati M, Fassad M, Hirst RA, Kenia P, Moya EF, Patel M, Peckham D, Rutman A, Mitchison HM, Mankad K, O’Callaghan C. Hydrocephalus and diffuse choroid plexus hyperplasia in primary ciliary dyskinesia-related MCIDAS mutation. Neurol Genet 2020; 6(4): e482

[45] Ibañez-Tallon I, Pagenstecher A, Fliegauf M, Olbrich H, Kispert A, Ketelsen UP, North A, Heintz N, Omran H. Dysfunction of axonemal dynein heavy chain Mdnah5 inhibits ependymal flow and reveals a novel mechanism for hydrocephalus formation. Hum Mol Genet 2004; 13(18): 2133–2141

[46] Mouse Genome Sequencing Consortium. Initial sequencing and comparative analysis of the mouse genome. Nature 2002; 420(6915): 520–562

[47] Rossi A, Kontarakis Z, Gerri C, Nolte H, Hölper S, Krüger M, Stainier DY. Genetic compensation induced by deleterious mutations but not gene knockdowns. Nature 2015; 524(7564): 230–233

[48] El-Brolosy MA, Kontarakis Z, Rossi A, Kuenne C, Günther S, Fukuda N, Kikhi K, Boezio GLM, Takacs CM, Lai SL, Fukuda R, Gerri C, Giraldez AJ, Stainier DYR. Genetic compensation triggered by mutant mRNA degradation. Nature 2019; 568(7751): 193–197

[49] van der Vaart J, Böttinger L, Geurts MH, van de Wetering WJ, Knoops K, Sachs N, Begthel H, Korving J, Lopez-Iglesias C, Peters PJ, Eitan K, Gileles-Hillel A, Clevers H. Modelling of primary ciliary dyskinesia using patient-derived airway organoids. EMBO Rep 2021; 22(12): e52058

[50] Sachs N, Papaspyropoulos A, Zomer-van Ommen DD, Heo I, Böttinger L, Klay D, Weeber F, Huelsz-Prince G, Iakobachvili N, Amatngalim GD, de Ligt J, van Hoeck A, Proost N, Viveen MC, Lyubimova A, Teeven L, Derakhshan S, Korving J, Begthel H, Dekkers JF, Kumawat K, Ramos E, van Oosterhout MF, Offerhaus GJ, Wiener DJ, Olimpio EP, Dijkstra KK, Smit EF, van der Linden M, Jaksani S, van de Ven M, Jonkers J, Rios AC, Voest EE, van Moorsel CH, van der Ent CK, Cuppen E, van Oudenaarden A, Coenjaerts FE, Meyaard L, Bont LJ, Peters PJ, Tans SJ, van Zon JS, Boj SF, Vries RG, Beekman JM, Clevers H. Long-term expanding human airway organoids for disease modeling. EMBO J 2019; 38(4): e100300

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