Skip to main content
NCBI home page
Wiley Open Access Collection logoLink to Wiley Open Access Collection
. 2026 Oct 4;61(10):e71866. doi: 10.1002/ppul.71866

Variants in CEP135 Cause Congenital Microcephaly and Primary Ciliary Dyskinesia

Abigail Bergman‐Sieger 1, Marc‐André Turcot 2, Lawrence E Ostrowski 3,4, Patrick R Sears 4, Maimoona A Zariwala 3,4, Fateh Bechkir 5, Lisa R Young 1,6, Adam J Shapiro 5,✉, Maureen B Parenti 1,6
PMCID: PMC13634309  PMID: 42829883

ABSTRACT

Introduction

Primary ciliary dyskinesia (PCD), a disorder of motile ciliary dysfunction causing chronic respiratory infections, may also rarely present with aspects of non‐motile ciliary dysfunction, including retinitis, central nervous system malformations, skeletal dysplasia, and sensorineural hearing loss. However, congenital microcephaly has not been previously associated with PCD.

Methods

We identified three children with congenital microcephaly and chronic respiratory disease consistent with PCD. Comprehensive PCD diagnostic testing and whole exome sequencing were performed. Characterization of nasal epithelial cells after expansion and regrowth included reverse transcription polymerase chain reaction, high speed videomicroscopy, and transcript analysis of novel genetic variants.

Results

Three pediatric cases from two families had primary microcephaly from autosomal recessive variants in CEP135. All cases displayed chronic suppurative respiratory symptoms, recurrent otitis media, and bronchiectasis with low nasal nitric oxide. One case had pathogenic, compound heterozygous, loss of function variants, while the other two cases had homozygous variants of uncertain significance. Transcript analyses of CEP135 variants of uncertain significance supported a splicing defect as disease‐causing. Regrown nasal epithelial cells demonstrated overall normal ciliary ultrastructure with decreased numbers of full‐length axonemes. Ciliary beat pattern was grossly abnormal, and distal axoneme swelling with bulbous‐tip structures were identified pre‐ and post‐cellular regrowth.

Conclusion

These cases establish variants in CEP135 as a novel cause of PCD with microcephaly. The TEM findings of bulbous ciliary tips may be a reliable marker of overlapping ciliopathies in patients with aspects of motile and non‐motile ciliary dysfunction.

Keywords: bronchiectasis, ciliopathy, microcephaly, primary ciliary dyskinesia

Abbreviations

CT

computed tomography

nNO

nasal nitric oxide

PCD

Primary Ciliary Dyskinesia

RT‐PCR

(reverse transcription polymerase chain reaction)

TEM

transmission electron microscopy

1. Introduction

Cilia are microtubule‐based organelles extending from the surface of most eukaryotic cells. Motile cilia typically exhibit a 9 + 2 microtubule arrangement and generate directional fluid flow, whereas non‐motile, primary cilia usually have a 9 + 0 microtubule configuration and serve sensory and signaling roles. Motile cilia are present in the respiratory epithelium, the central nervous system, the developing embryo, and the reproductive tract, where they enable mucociliary clearance, cerebrospinal fluid circulation, normal embryonic laterality, and propulsion of gametes [1]. Non‐motile cilia play critical roles in developmental signaling pathways and may cause malformations in the brain, retina, kidney, liver, and skeleton [2].

Primary ciliary dyskinesia (PCD, OMIM #244000) is a rare, genetically heterogeneous disorder of motile cilia leading to impaired mucociliary clearance and chronic oto‐sino‐pulmonary disease with progressive bronchiectasis. Diagnostic confirmation requires either a disease‐causing ciliary ultrastructural defect identified on transmission electron microscopy (TEM) or pathogenic variants in 1 of 60 PCD‐associated genes [3, 4]. Ancillary PCD screening and diagnostic tests include nasal nitric oxide (nNO) measurement, high‐speed videomicroscopy analysis of ciliary beat pattern, and ciliary protein immunofluorescence.

CEP135 is a gene encoding a scaffold protein essential for centriole duplication and basal body anchoring in both motile and non‐motile cilia [5]. Pathogenic variants in CEP135 are a well‐established cause of autosomal recessive primary microcephaly but are not associated with a motile ciliopathy [6]. We report novel cases with CEP135 variants that exhibit aspects of an overlapping motile and non‐motile ciliopathy with PCD and microcephaly. We further characterize the epithelial cellular properties and changes in these cases and expand upon an emerging TEM phenotype, which may be representative of overlapping, non‐motile ciliopathies with PCD.

2. Methods

All participants consented to approved protocols at Children's Hospital of Philadelphia (CHOP, IRB #014568) or the McGill University Health Centre (MUHC, IRB #2023‐9523). For Case 1 at CHOP (CH‐1), trio exome sequencing (ES) was performed at GeneDx, and TEM was analyzed at Mayo Clinic Laboratories. For MUHC cases (MC‐1, MC‐2), ES was performed by Blueprint Genetics, which consists of analysis of all protein coding genes coupled with their development disorder panel, while TEM analysis was done in‐house. Nasal nitric oxide (nNO) was measured by chemiluminescence technology using CLD88 devices from Eco Physics (Duernten, Switzerland) and per PCD Foundation protocol [7].

For Cases MC‐1 and MC‐2, nasal epithelial cells were expanded as conditionally reprogrammed cells (CRC) by the Marsico Lung Institute Tissue Procurement and Cell Culture Core at the University of North Carolina (UNC). Expanded cells were cultured at the air/liquid interface on collagen coated MilliCell inserts in Pneumacult media until differentiated, as previously described [8]. Differentiated nasal epithelial cells were analyzed for ciliary beat frequency (CBF) and waveform by high speed videomicroscopy. For all video recordings, a Nikon Eclipse inverted microscope was used with an OkoLab microscope incubator set to 37°C, 5% CO2, and humidified air. Videos were captured using SAVA software (Ammons Engineering, Clio, MI). For CBF, two MC‐1 and two MC‐2 cultures were measured. Each culture was first given a 5‐min Phosphate‐Buffered Saline (PBS) wash. Then 15 µL of PBS was added to the surface and the cultures were transferred to the microscope stage incubator. The cultures were allowed to equilibrate for 5 min. Then videos were taken at places where cilia could be seen beating with a measurable frequency. A 20× phase objective was used, and for each video, 512 frames were captured at 120 frame/second. Post‐capture, videos were manually searched for areas that showed a high‐quality frequency.

For waveform analysis, one MC‐1 and one MC‐2 culture were used. For each culture, the membrane was first detached from the insert. Cells were then scraped off using a glass coverslip and transferred to a flow chamber. The chamber was placed on the microscope stage and allowed to equilibrate for 5 min. Cells were observed using a 60× oil objective (DIC, NA 1.4) with 2× post‐objective optical magnification. For each video, 512 frames were captured at 300 frames/second. Videos were captured of cilia in profile and in top‐down views.

To evaluate the effect of intron 8 splice region VUS (c.1044+3A>G), cultured human nasal epithelial cells (HNECs) were generated from the probands MC‐1 and MC‐2 using a previously described protocol [8]. Reverse transcription polymerase chain reaction (RT‐PCR) for CEP135 exons 6–9 was performed on RNA from HNECs isolated using a RNeasy Kit (Qiagen, Germantown, MD, USA) as per manufacturer's instructions. First strand cDNA was synthesized using SuperScript II Reverse Transcriptase kit (Thermo Fisher Scientific). Gene‐specific primers 5′‐GGCTGATAACAGGATTCAAGAAC‐3′ (sense) and 5′‐GGCATAAGTTCAATTCCAGCTGAAG‐3′ (antisense) were used for amplification to generate 542‐bp product for wild type (reference sequence) and Sanger sequencing. PCR was carried out using 2 µL cDNA, 400 pmol each primer, 1X PCR buffer I, containing 1.5 mM MgCl2 (Applied Biosystem), 0.2 mM total dNTP (Promega), and 0.05 units of AmpliTaq polymerase (Sigma‐Aldrich) in 20 µL total volume. Reactions were cycle sequenced (ABI 2720 Thermal Cycler, Applied Biosystem) for initial denaturation at 94°C for 5 min, followed by 35 cycles of denaturation at 94°C for 30 s, annealing at 60°C for 45 s, and extension at 72°C for 45 s, followed by a final extension at 72°C for 10 min. Amplified fragments were visualized by 2% agarose gel electrophoresis, and treated with ExoSAP‐IT (Applied Biosystem) prior to carrying out Sanger sequencing. To confirm the findings, RT‐PCR was repeated using an additional primer set (sequences not shown).

3. Results

3.1. Case One (CH‐1)

A 33‐month‐old white male with congenital microcephaly and growth delay presented for recurrent respiratory tract infections and chronic cough. He was born full‐term without neonatal respiratory distress. At 7 weeks of age, he developed chronic rhinorrhea and wet cough, requiring hospitalization for pneumonia. By 15 months, he had multiple hospitalizations for respiratory infections. Chest imaging revealed normal organ placement but repeatedly demonstrated persistent areas of opacification. He also experienced recurrent otitis media requiring tympanostomy tube placement by 6 months of age.

Growth history revealed both congenital and postnatal microcephaly with growth restriction. At birth, head circumference was 29.7 cm (0.01 percentile), while weight and length were at the 25th percentile. At 36 months of age, head circumference was 43 cm (0.01 percentile; equivalent to the 50th percentile for a 6‐month‐old) and weight and length decreased to the 3rd and 1st percentiles, respectively. Chest computed tomography (CT) demonstrated right middle lobe and lingula collapse, mucus plugging, and cylindrical bronchiectasis (Figure 1). Flexible bronchoscopy showed purulent secretions, and cultures grew Pseudomonas aeruginosa, Streptococcus pneumoniae, Moraxella catarrhalis, and Haemophilus influenzae. Additional studies, including a videofluoroscopic swallow study, echocardiogram, sweat chloride testing, and a thorough immunologic evaluation, were all normal. At 5 and 6 years of age, nNO values, measured by exhalation against resistance, were low at 13 and 69 nL/min. TEM of airway epithelial cells showed intermittent central microtubule defects and some basal body misplacement with basal foot misalignment, but was overall non‐diagnostic (Figure 2).

Figure 1.

Figure 1

Open in a new tab

Radiology imaging in CEP135‐affected cases. (A, B) Coronal and axial computed tomography (CT) in case CH‐1 showing right middle lobe and lingula collapse with cylindrical bronchiectasis at 4 years of age (white arrowheads). (C, D) Coronal and axial CT images in case MC‐1 showing lobar atelectasis with bronchiectasis in right middle lobe at 13 years of age. (E, F) Coronal and axial CT images in case MC‐2 showing severe, saccular bronchiectasis affecting all lobes at 12 years of age. (G, H) Axial and sagittal magnetic resonance imaging of the brain in case MC‐2 showing foci of gray matter heterotopia in the lateral ventricles (white arrow) and microcephaly with underdeveloped frontal lobes and a simplified gyral pattern.

Figure 2.

Figure 2

Open in a new tab

Transmission ciliary electron microscopy (TEM) images from CEP‐135 affected cases. (A–D) TEM images from CH‐1, showing basal body misalignment and misplacement within the cell cytoplasm (A); Non‐diagnostic ultrastructural changes with 8 outer microtubule doublets plus a central single microtubule (8 + 1) (B), 9 outer microtubule doublets with 2 central pair (9 + 4) (C), and 8 outer microtubule doublets and a single central pair (8 + 2) (D). Dynein arms appear normal. (E, F) TEM images from MC‐1 showing decreased ciliary numbers but normally aligned basal bodies at the apical cell surface (E), and normal, 9 + 2 ciliary ultrastructure on axonemal cross‐sections (F). (G) TEM image from MC‐2 showing normal, 9 + 2 ciliary ultrastructure on axonemal cross‐sections.

Trio exome sequencing with both parents did not detect variants in established PCD‐causing genes but did reveal biallelic, pathogenic, loss of function variants in CEP135, previously reported in ClinVar (variation ID #982087, NM_025009.5, c.2722C>T, (p.Arg908*) and #977828, c.3211A>T, (p.Lys1071*)). These are consistent with primary autosomal recessive microcephaly Type 8 (OMIM #611423) associated with severe microcephaly, short stature, and intellectual disability (though this patient did not exhibit developmental or cognitive delays).

3.2. Cases 2 (MC‐1) and 3 (MC‐2)

Two brothers, ages 13 (MC‐1) and 11 (MC‐2) years, presented for chronic cough. They were refugees from Afghanistan, and family history revealed parental consanguinity (second cousins). Respiratory symptoms in both included daily wet cough since birth, recurrent pneumonias, year‐round nasal congestion since birth, and chronic otitis media. Neither had neonatal respiratory distress nor organ laterality defects. Both had microcephaly, short stature (Z‐score for height MC‐1, −1.72; MC‐2, −1.51), and severe cognitive dysfunction. Magnetic resonance imaging of their brains showed foci of gray matter heterotopia in the lateral ventricles, microcephaly with underdeveloped frontal lobes and a simplified gyral pattern, and dysmorphic/shortened corpus callosum (Figure 1).

Computed tomography chest scans revealed atelectasis with bronchiectasis in the right middle lobe and lingula in MC‐1, and saccular bronchiectasis affecting all lobes in MC‐2 (Figure 1). Videofluoroscopic swallowing studies were negative for aspiration. Both had normal immune function and negative cystic fibrosis transmembrane conductance regulator (CFTR) sequencing. Expectorated sputum cultures grew Haemophilus influenzae in both, with mucoid Pseudomonas aeruginosa in MC‐2. They were unable to complete pulmonary function testing due to developmental delay. Nasal nitric oxide, measured by tidal breathing [9], was repeatedly low in both siblings (MC‐1, 41, 27, 28 nL/min; MC‐2, 26, 41 nL/min).

Analysis of nasal cells by TEM showed normal 9 + 2 ciliary ultrastructure. However, the overall number of full‐length ciliary axonemes appeared decreased per epithelial cell, with normal basal body alignment (Figure 2). Bulbous ciliary tips with eccentric microtubules were seen in limited images from MC‐2 (Figure 3). Nasal epithelial cells from MC‐1 and MC‐2 were successfully expanded and differentiated in air/liquid interface culture. Histological sections revealed a multi‐layered epithelium with a non‐uniform ciliary surface (Figure 3). While some cells appeared to have patches of cilia of uniform length, other cells appeared to have few cilia, and many cilia had swollen bulb‐like structures at their tips. Examination of intact nasal epithelial cell cultures by high‐speed videomicroscopy did not reveal consistent ciliary beating, with standard procedures unable to report any consistent CBF. High‐speed videomicroscopy of isolated cell clumps from MC‐1 and MC‐2 nasal epithelial cells culture showed an uncoordinated and irregular beat pattern. Most waveforms appeared quite abnormal and desynchronized from adjacent cilia (Figure S1). In addition, bulbous ciliary tips could be seen on many cilia (Videos [Link], [Link], [Link], [Link], [Link]).

Figure 3.

Figure 3

Open in a new tab

Transmission ciliary electron microscopy (TEM) and light microscopy images from case MC‐2. (A, B) TEM images showing swollen, bulbous ciliary tips with eccentric microtubule structure. (C) Light microscopy (60X magnification) of hematoxylin and eosin‐stained epithelial cells after air/liquid interface culture of nasal epithelial cells from MC‐2, showing numerous cilia with swollen tips (arrows) while some cells appear to have reduced cilia numbers.

Exome sequencing of both brothers revealed an apparent homozygous variant of uncertain significance (VUS) in CEP135 (NM_005029.5; c.1044+3 A>G, p.(splice)), but parental specimens were not available (Figure 4). No other disease associated pathogenic variants in either PCD‐related, or other phenotype‐relevant genes, were found. The brothers also carried a hemizygous variant in DLG3 (NM_021120.4; c.1748G>A, p.(Arg583Lys), which is implicated in an X‐linked form of intellectual disability (OMIM# 300189). This variant is intriguing as it is rare in large population database (gnomAD) and has a CADD score of 25.5, but without functional studies, it is currently considered a VUS (Clinvar variation ID #4083349).

Figure 4.

Figure 4

Open in a new tab

Transcript analysis for CEP135 (NM_005025.5) variant c.1044+3A>G in Intron 8: (A) Exome sequencing by Blueprint Genetics (https://blueprintgentics.com/) identified an apparent homozygous variant c.1044+3A>G in affected sibs. (B) Transcript analysis of CEP135 (NM_005025.5) showing effect of c.1044+3A>G variant by qualitative RT‐PCR followed by agarose gel electrophoresis. Control samples revealed a single 524‐bp fragment corresponding to the wild type (reference) transcript. In the affected siblings, a major longer fragment of 579‐bp and two minor products of 179‐bp and 524‐bp were observed. Following primers were used for CEP135 RT‐PCR and sequencing: 5′‐GGCTGATAACAGGATTCAAGAAC3′ (Exon 5/6 Sense) and 5′‐GGCATAAGTTCAATTCCAGCTGAAG3′ (Exon 9/10 Antisense). (C, D) Representative electropherograms showing the transcripts corresponding to exons 7 and 8, and exons 8 and 9 junctions in control. (E) Representative electropherograms showing the two minor products, one corresponding to an aberrant transcript of 179‐bp due to an in‐frame deletion of exons 7 and 8 (r.700_1044del), p.(Ile234_Lys348del) and other a wildtype transcript due to leaky splicing in affected siblings (MC‐1 and MC‐2). (D–F) Representative electropherograms showing the location of the pathogenic variant (r.1044+3A>G) in a major longer transcript, with the inclusion of 55‐bp pseudo‐exon (r.1044+1_1044+55ins). This is predicted to lead to the frameshift [p.(Lys349Valfs*9)], likely affecting protein function. Base sequences, amino acid sequences and codon numbers are indicated. Location of the pathogenic variant in the major longer transcript is shown by a red diamond and variant and the pseudo‐exon are shown with gray highlights. Exon/exon junctions are indicated by red vertical broken line. [Color figure can be viewed at wileyonlinelibrary.com]

The VUS c.1044+3A>G in CEP135 is not listed in Clinvar, but present in the dbSNP database (https://www.ncbi.nlm.nih.gov/snp/) as rs752733205 [10], and present at a very low allele frequency (0.000495%) in a population cohort (https://gnomad.broadinstitute.org/) [11]. To evaluate the functional consequences of this VUS, qualitative transcript analysis was performed (Figure 4). Cultured nasal cells from both patients underwent reverse transcription polymerase chain reaction (RT‐PCR) analysis and were compared to cells from a healthy control. Primers encompassing exons 6–9 revealed a single major fragment of expected size (524‐bp) in RNA prepared from normal airway epithelia (control). In contrast, RNA from the affected siblings homozygous for c.1044+3A>G produced a longer major transcript and two minor transcripts. The longer transcript (579‐bp) revealed a frameshift insertion of 55‐bp pseudo‐exon (r.1044+1_1044+55ins) which is predicted to lead to premature translation termination signal [p.(Lys349Valfs*9)] presumably affecting protein function. Additionally, the r.1044+3A>G variant was visible in the longer transcript. One of the minor products represented aberrant transcript of 179‐bp due to an in‐frame deletion of exons 7 and 8 [(r.700_1044del), p. (Ile234_Lys348del)], while the other minor product was of expected size representing leaky splicing. The transcript analysis was validated with an independent primer set (data not shown). This provides evidence that c.1044+3A>G is disease‐causing due to splicing disruption.

4. Discussion

We present the first cases of variants in CEP135 resulting in an overlapping syndrome with aspects of a non‐motile ciliopathy and PCD. CEP135 encodes a centrosome scaffold protein essential for centriole duplication and basal body anchoring in both motile and non‐motile cilia [5]. Variants in CEP135 cases are linked to autosomal recessive primary microcephaly with growth restriction and developmental delay [6]. While CEP135 has not been previously associated with respiratory ciliary dysfunction, disruption of centriole architecture impairs basal body docking and ciliary assembly, providing a mechanistic basis for the motile ciliopathy phenotype observed here. Male infertility via effects on sperm flagellar structure and dysmotility have also been reported with CEP135 variants in mice and humans, supporting the likely involvement of CEP135 in motile ciliary function [12, 13].

The phenotypes we present are consistent with PCD and include the key clinical features of year‐round wet cough and rhinosinusitis from early infancy, plus common features of recurrent pneumonia, chronic otitis media, and bronchiectasis [14]. In addition, cultures of nasal epithelial cells display abnormal cilia and a clearly defective waveform. However, our patients lack other key PCD symptoms of neonatal respiratory distress and organ laterality defects. While the pathophysiology underlying neonatal respiratory distress in PCD is unknown, it is well recognized that other PCD genotypes affecting the ciliary basal body (GAS2L2, CCNO) similarly lack organ laterality defects as these structures are not required for normal function of embryonic nodal cilia, which direct prenatal organ placement [15, 16]. The motile ciliary phenotype that we report here may not have been previously recognized in patients with CEP135 variants, as caregivers may mistakenly attribute chronic respiratory symptoms to more common mechanisms, including pulmonary aspiration or poor cough clearance, which are often seen in neurologically delayed patients.

PCD is difficult to diagnose, as no single test detects all forms of this disease. Diagnostic confirmation requires either a class 1 ultrastructural defect on TEM analysis [17] or biallelic, disease‐causing variants in a gene associated with autosomal recessive PCD (or a single variant in the few X‐linked or autosomal dominant forms of PCD). Ancillary PCD tests with nNO measurement, high speed videomicroscopy analysis, or ciliary protein immunofluorescence, are difficult to access outside of expert centers. Repeatedly low nNO levels strongly suggest PCD, yet 10% of known PCD cases have normal nNO values [18]. As in some of our cases, TEM analysis often shows non‐specific changes which can be misleading. However, several rare PCD genotypes show TEM findings of bulbous ciliary tips with eccentric microtubules and decreased full‐length axoneme number per epithelial cell. Each of these past, bulbous ciliary defect cases report an overlapping phenotype of PCD with aspects of a non‐motile ciliopathy, including variants in TUBB4B causing PCD with sensorineural hearing loss, Leber congenital amaurosis, skeletal and renal malformations [19], and variants in KIAA0586/TALPID causing Joubert Syndrome [20]. Thus, in suspected cases of PCD accompanied by aspects of a non‐motile ciliopathy phenotype, these characteristic bulbous tips on TEM may strongly support a PCD diagnosis. Notably, detection of these bulbous ciliary changes may not be achieved on the customary mid‐axonemal cuts required to assess the 9 + 2 ciliary ultrastructure on TEM. Instead, detection of these bulbous changes may require imaging of strips of ciliated epithelium at lower power magnification, assessing the distal axoneme tips. Moreover, our TEM images showed scattered and limited bulbous changes, which were much less apparent compared to their presence on regrown cells. This difference may stem from sample processing and/or selection bias which occurs with TEM fixation and analysis without regrowth.

An increasing number of ciliopathies have syndromic manifestations accompanied by the chronic respiratory symptoms of PCD. These include retinitis pigmentosa associated with RPGR variants [21]; neurodevelopmental impairment and macrocephaly reported with OFD1 variants [22]; Joubert‐spectrum phenotypes associated with variants in KIAA0586 (TALPID3) or CBY1 [20, 23]; and skeletal ciliopathies such as Jeune/short‐rib thoracic dysplasia associated with IFT74 or DYNC2H1 variants [24, 25]. Hydrocephalus and other neurological manifestations have also been described in PCD with variants in FOXJ1, TUBB4B, MCIDAS, or TP73, the latter also being associated with brain malformations including lissencephaly [19, 26, 27, 28]. Recognition of PCD in individuals with microcephaly and cognitive dysfunction, as observed in our patients with CEP135 variants, therefore expands the clinical spectrum of syndromic ciliopathies and supports multidisciplinary evaluation by respiratory, neurologic, developmental, ophthalmologic, and genetic specialists.

These cases also highlight the importance of broadening the genetic framework for investigating rare lung diseases. Next‐generation sequencing genetic panels analyzing variants in 40–50 known PCD genes are often first‐line investigations in suspected PCD, yet these do not detect cases involving genes that are not yet definitively linked to PCD. In our three cases, exome sequencing, prompted by syndromic features of microcephaly and growth impairment, enabled identification of CEP135 variants, which are not analyzed on standard PCD genetic panels. In approximately 6% of PCD cases, non‐coding genetic variants can also be disease‐causing, and whole genome sequencing analysis increases diagnostic success in PCD [29, 30]. The benefits of whole genome sequencing have also been repeatedly demonstrated in diagnosing non‐motile ciliopathies [31, 32, 33]. These results underscore the need to pursue exome and genome analysis when suppurative respiratory disease or bronchiectasis coexist with extra‐pulmonary anomalies seen in non‐motile ciliopathies. As knowledge of the intricate protein interactions that drive motile cilia formation advances, additional genes essential to airway structure and function are also likely to be discovered through genomic analysis.

5. Conclusion

This case series expands the phenotypic spectrum of CEP135 variants and establishes a previously unrecognized genetic cause of PCD. It underscores the importance of comprehensive genetic testing in children with unexplained bronchiectasis accompanied by syndromic features, including microcephaly or other findings common to non‐motile ciliopathies. A confirmed PCD diagnosis in individuals with CEP135 variants supports timely specialist care, including regular airway clearance, prompt treatment of respiratory exacerbations, and consideration of prophylactic azithromycin [34]. It may also guide surveillance for bronchiectasis which may facilitate access to emerging disease‐modifying treatments such as brensocatib [35]. Distal cilia with bulbous malformations may be a reliable indicator of an overlapping ciliopathy with both motile and non‐motile ciliary dysfunction, but further research on this finding is required for validation.

Author Contributions

Abigail Bergman‐Sieger: conceptualization, methodology, investigation, writing – review and editing, writing – original draft, formal analysis, data curation. Marc‐André Turcot: investigation, writing – original draft, writing – review and editing, data curation. Lawrence E. Ostrowski: conceptualization, investigation, methodology, validation, formal analysis, data curation, resources, writing – review and editing. Patrick R. Sears: methodology, software, data curation, formal analysis, investigation, writing – review and editing, resources. Maimoona A. Zariwala: methodology, software, data curation, validation, investigation, formal analysis, visualization, writing – review and editing, resources. Fateh Bechkir: software, data curation, formal analysis, writing – review and editing, visualization, resources. Lisa R. Young: conceptualization, methodology, investigation, writing – original draft, writing – review and editing, supervision, data curation, project administration. Adam J. Shapiro: conceptualization, investigation, methodology, validation, data curation, supervision, writing – original draft, writing – review and editing. Maureen B. Parenti: conceptualization, investigation, writing – original draft, writing – review and editing, methodology, validation, data curation, supervision, resources, project administration.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File 1

PPUL-61-0-s006.png (426.9KB, png)

Supporting File 2

Download video file (8.6MB, mp4)

Supporting File 3

Download video file (9MB, mp4)

Supporting File 4

Download video file (8.9MB, mp4)

Supporting File 5

Download video file (10.7MB, mp4)

Supporting File 6

Download video file (9.9MB, mp4)

Acknowledgments

We thank the research participants, Research coordinators, as well as investigators and coordinators of the Genetic Disorders of Mucociliary Clearance Consortium, part of the Rare Disease Clinical Research Network. Funding support for AJS, MAZ was provided by US NIH/ORDR/NACTS/NHLBI grant U54HL096458, R01HL117836. The Genetic Disorders of Mucociliary Clearance Consortium (U54HL096458) is part of the National Center for Advancing Translational Sciences (NCATS) Rare Diseases Clinical Research Network (RDCRN) and is supported by the RDCRN Data Management and Coordinating Center (DMCC) (U2CTR002818). RDCRN is an initiative of the Office of Rare Diseases Research (ORDR) funded through a collaboration between NCATS and NHLBI. Funding support for LEO, PRS provided by US NIH R01HL1178336. Funding support for ABS is supported by the Cystic Fibrosis Foundation through a Clinical Fellowship–Education and Training Grant. The funding organizations had no involvement in the conduct of the research or the decision to submit the manuscript.

Bergman‐Sieger A., Turcot M.‐A., Ostrowski L. E., et al., “Variants in CEP135 Cause Congenital Microcephaly and Primary Ciliary Dyskinesia,” Pediatric Pulmonology 68 (2026): e71866. 10.1002/ppul.71866.

Adam J. Shapiro and Maureen B. Parenti are co‐senior authors.

Data Availability Statement

The data that support the findings of this study are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.

Males and females are designated by the squares and circles, respectively. Filled symbols show affected individuals. Proband designated with an arrow next to the filled symbol, homozygous (Hom), heterozygous (Het) and wild type (WT) are labeled.

References

  • 1. Fliegauf M., Benzing T., and Omran H., “When Cilia Go Bad: Cilia Defects and Ciliopathies,” Nature Reviews Molecular Cell Biology 8, no. 11 (2007): 880–893, 10.1038/nrm2278. [DOI] [PubMed] [Google Scholar]
  • 2. Reiter J. F. and Leroux M. R., “Genes and Molecular Pathways Underpinning Ciliopathies,” Nature Reviews Molecular Cell Biology 18, no. 9 (2017): 533–547, 10.1038/nrm.2017.60. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3. Shoemark A., Goutaki M., Kinghorn B., et al., “European Respiratory Society and American Thoracic Society Guidelines for the Diagnosis of Primary Ciliary Dyskinesia,” European Respiratory Journal 66, no. 6 (2025): 2500745, 10.1183/13993003.00745-2025. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4. Zariwala M. A., Despotes K. A., and Davis S. D., “Primary Ciliary Dyskinesia,” in GeneReviews(®), eds. Adam M. P., Bick S., Mirzaa G. M., Pagon R. A., Wallace S. E., and Amemiya A.. University of Washington, 1993. [PubMed] [Google Scholar]
  • 5. Lin Y. C., Chang C. W., Hsu W. B., et al., “Human Microcephaly Protein CEP135 Binds to hSAS‐6 and CPAP, and Is Required for Centriole Assembly,” EMBO Journal 32, no. 8 (2013): 1141–1154, 10.1038/emboj.2013.56. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6. Hussain M. S., Baig S. M., Neumann S., et al., “A Truncating Mutation of CEP135 Causes Primary Microcephaly and Disturbed Centrosomal Function,” American Journal of Human Genetics 90, no. 5 (2012): 871–878. 10.1016/j.ajhg.2012.03.016. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 7. Shapiro A. J., Dell S. D., Gaston B., et al., “Nasal Nitric Oxide Measurement in Primary Ciliary Dyskinesia. A Technical Paper on Standardized Testing Protocols,” Annals of the American Thoracic Society 17, no. 2 (2020): e1–e12, 10.1513/AnnalsATS.201904-347OT. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8. Smith A. J., Bustamante‐Marin X. M., Yin W., et al., “The Role of SPAG1 in the Assembly of Axonemal Dyneins in Human Airway Epithelia,” Journal of Cell Science 135, no. 6 (2022): jcs259512, 10.1242/jcs.259512. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9. Leigh M. W., Shapiro A. J., Chawla K. K., et al., “Tidal Breathing Nasal Nitric Oxide Measurement as a Test for Primary Ciliary Dyskinesia in Young Children,” European Respiratory Journal 67 (2026): 2501540, 10.1183/13993003.01540-2025.. [DOI] [PubMed] [Google Scholar]
  • 10. Phan L., Zhang H., Wang Q., et al., “The Evolution of Dbsnp: 25 Years of Impact in Genomic Research,” Nucleic Acids Research 53, no. D1 (2025): D925–D931, 10.1093/nar/gkae977. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11. Karczewski K. J., Francioli L. C., Tiao G., et al., “The Mutational Constraint Spectrum Quantified from Variation in 141,456 Humans,” Nature 581, no. 7809 (2020): 434–443, 10.1038/s41586-020-2308-7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12. Liu H., Zhang H., Qin G., et al., “Loss of Cep135 Causes Oligoasthenoteratozoospermia and Male Infertility in Mice,” Cellular and Molecular Life Sciences 82, no. 1 (2025): 117, 10.1007/s00018-025-05616-w. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13. Sha Y. W., Xu X., Mei L. B., et al., “A Homozygous CEP135 Mutation Is Associated With Multiple Morphological Abnormalities of the Sperm Flagella (MMAF),” Gene 633 (2017): 48–53, 10.1016/j.gene.2017.08.033. [DOI] [PubMed] [Google Scholar]
  • 14. Leigh M. W., Ferkol T. W., Davis S. D., et al., “Clinical Features and Associated Likelihood of Primary Ciliary Dyskinesia in Children and Adolescents,” Annals of the American Thoracic Society 13, no. 8 (2016): 1305–1313, 10.1513/AnnalsATS.201511-748OC. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15. Bustamante‐Marin X. Z. M., Yin W., Sears P. R., Knowles M. R., and Ostrowski L. E., Lack of GAS2L2 Induces Uncoordinated and Hyperactive Ciliary Beat in Human and Mouse Ciliated Cells. PCD Foundation Annual Meeting, 2017. [Google Scholar]
  • 16. Wallmeier J., Al‐Mutairi D. A., Chen C. T., et al., “Mutations in CCNO Result in Congenital Mucociliary Clearance Disorder With Reduced Generation of Multiple Motile Cilia,” Nature Genetics 46, no. 6 (2014): 646–651, 10.1038/ng.2961. [DOI] [PubMed] [Google Scholar]
  • 17. Shoemark A., Boon M., Brochhausen C., et al., “International Consensus Guideline for Reporting Transmission Electron Microscopy Results in the Diagnosis of Primary Ciliary Dyskinesia (BEAT PCD TEM Criteria),” European Respiratory Journal 55, no. 4 (2020): 1900725, 10.1183/13993003.00725-2019. [DOI] [PubMed] [Google Scholar]
  • 18. Raidt J., Krenz H., Tebbe J., et al., “Limitations of Nasal Nitric Oxide Measurement for Diagnosis of Primary Ciliary Dyskinesia With Normal Ultrastructure,” Annals of the American Thoracic Society 19, no. 8 (2022): 1275–1284, 10.1513/AnnalsATS.202106-728OC. [DOI] [PubMed] [Google Scholar]
  • 19. Dodd D. O., Mechaussier S., Yeyati P. L., et al., “Ciliopathy Patient Variants Reveal Organelle‐Specific Functions for TUBB4B in Axonemal Microtubules,” Science 384, no. 6694 (2024): eadf5489, 10.1126/science.adf5489. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20. Taudien J. E., Bracht D., Olbrich H., et al., “Pathogenic KIAA0586/TALPID3 Variants Are Associated with Defects in Primary and Motile Cilia,” iScience 28, no. 2 (2025): 111670, 10.1016/j.isci.2024.111670. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21. Moore A., Escudier E., Roger G., et al., “RPGR Is Mutated in Patients With a Complex X Linked Phenotype Combining Primary Ciliary Dyskinesia and Retinitis Pigmentosa,” Journal of Medical Genetics 43, no. 4 (2006): 326–333, 10.1136/jmg.2005.034868. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22. Budny B., Chen W., Omran H., et al., “A Novel X‐Linked Recessive Mental Retardation Syndrome Comprising Macrocephaly and Ciliary Dysfunction Is Allelic to Oral‐Facial‐Digital Type I Syndrome,” Human Genetics 120, no. 2 (2006): 171–178, 10.1007/s00439-006-0210-5. [DOI] [PubMed] [Google Scholar]
  • 23. Epting D., Senaratne L. D. S., Ott E., et al., “Loss of CBY1 Results in a Ciliopathy Characterized by Features of Joubert Syndrome,” Human Mutation 41, no. 12 (2020): 2179–2194, 10.1002/humu.24127. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24. Fassad M. R., Rumman N., Junger K., et al., “Defective Airway Intraflagellar Transport Underlies a Combined Motile and Primary Ciliopathy Syndrome Caused by IFT74 Mutations,” Human Molecular Genetics 32, no. 21 (2023): 3090–3104, 10.1093/hmg/ddad132. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25. Emiralioglu N., Wallmeier J., Olbrich H., Omran H., and Ozcelik U., “DYNC2H1 Mutation Causes Jeune Syndrome and Recurrent Lung Infections Associated With Ciliopathy,” Clinical Respiratory Journal 12, no. 3 (2018): 1017–1020, 10.1111/crj.12620. [DOI] [PubMed] [Google Scholar]
  • 26. Wallmeier J., Frank D., Shoemark A., et al., “De Novo Mutations in FOXJ1 Result in a Motile Ciliopathy With Hydrocephalus and Randomization of Left/Right Body Asymmetry,” American Journal of Human Genetics 105, no. 5 (2019): 1030–1039, 10.1016/j.ajhg.2019.09.022. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27. Robson E. A., Dixon L., Causon L., et al., “Hydrocephalus and Diffuse Choroid Plexus Hyperplasia in Primary Ciliary Dyskinesia‐Related MCIDAS Mutation,” Neurology: Genetics 6, no. 4 (2020): e482, 10.1212/nxg.0000000000000482. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28. Wallmeier J., Bracht D., Alsaif H. S., et al., “Mutations in TP73 Cause Impaired Mucociliary Clearance and Lissencephaly,” American Journal of Human Genetics 108, no. 7 (2021): 1318–1329, 10.1016/j.ajhg.2021.05.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29. Briggs L., Brandão C., Fleming A., et al., “Noncoding DNA Variants Increase the Genetic Diagnostic Yield in Primary Ciliary Dyskinesia,” American Journal of Respiratory and Critical Care Medicine 211, no. 11 (2025): 2043–2052, 10.1164/rccm.202501-0186OC. [DOI] [PubMed] [Google Scholar]
  • 30. Black H. A., de Proce S. M., Campos J. L., et al., “Whole Genome Sequencing Enhances Molecular Diagnosis of Primary Ciliary Dyskinesia,” Pediatric Pulmonology 59, no. 12 (2024): 3322–3332, 10.1002/ppul.27200. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31. Mallawaarachchi A. C., Lundie B., Hort Y., et al., “Genomic Diagnostics in Polycystic Kidney Disease: An Assessment of Real‐World Use of Whole‐Genome Sequencing,” European Journal of Human Genetics 29, no. 5 (2021): 760–770, 10.1038/s41431-020-00796-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32. Geoffroy V., Stoetzel C., Scheidecker S., et al., “Whole Genome Sequencing in Patients with Ciliopathies Uncovers a Novel Recurrent Tandem Duplication in IFT140,” Human Mutation 39, no. 7 (2018): 983–992, 10.1002/humu.23539. [DOI] [PubMed] [Google Scholar]
  • 33. Stellacci E., Niceta M., Bruselles A., et al., “Whole Genome Sequencing Solves an Atypical Form of Bardet‐Biedl Syndrome: Identification of Novel Pathogenic Variants of BBS9,” International Journal of Molecular Sciences 25, no. 15 (2024): 8313. 10.3390/ijms25158313. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34. Kobbernagel H. E., Buchvald F. F., Haarman E. G., et al., “Efficacy and Safety of Azithromycin Maintenance Therapy in Primary Ciliary Dyskinesia (BESTCILIA): A Multicentre, Double‐Blind, Randomised, Placebo‐Controlled Phase 3 Trial,” Lancet Respiratory Medicine 8, no. 5 (2020): 493–505, 10.1016/s2213-2600(20)30058-8. [DOI] [PubMed] [Google Scholar]
  • 35. Chalmers J. D., Burgel P. R., Daley C. L., et al., “Phase 3 Trial of the DPP‐1 Inhibitor Brensocatib in Bronchiectasis,” New England Journal of Medicine 392, no. 16 (2025): 1569–1581, 10.1056/NEJMoa2411664. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supporting File 1

PPUL-61-0-s006.png (426.9KB, png)

Supporting File 2

Download video file (8.6MB, mp4)

Supporting File 3

Download video file (9MB, mp4)

Supporting File 4

Download video file (8.9MB, mp4)

Supporting File 5

Download video file (10.7MB, mp4)

Supporting File 6

Download video file (9.9MB, mp4)

Data Availability Statement

The data that support the findings of this study are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.

Males and females are designated by the squares and circles, respectively. Filled symbols show affected individuals. Proband designated with an arrow next to the filled symbol, homozygous (Hom), heterozygous (Het) and wild type (WT) are labeled.

Articles from Pediatric Pulmonology are provided here courtesy of Wiley

RESOURCES