Ccdc57 regulates cilia and left-right patterning in Xenopus
Binyi Yang, Emily K. Mis, Xianglin Zhou, Faiza Aslam, Jie He, Xiangyang Lu, Hui Fan, Ting Guo, Engin Deniz, Hong Luo, Mustafa K. Khokha
TL;DR
This study shows that CCDC57 is important for cilia function and left-right body asymmetry in frogs and may be linked to human disorders like situs inversus.
Contribution
The study identifies CCDC57 as a novel regulator of ciliary function and left-right patterning in Xenopus and links it to human laterality disorders.
Findings
Depletion of ccdc57 in Xenopus leads to abnormal cilia and disrupted left-right patterning.
Human CCDC57 variants from a patient with situs inversus fail to rescue ciliary defects in Xenopus.
CCDC57 is essential for normal ciliary structure and function during embryonic development.
Abstract
During embryogenesis, the establishment of left–right (LR) asymmetry depends on directional fluid flow generated by motile cilia within the left–right organizer (LRO). Disruption of this process can lead to laterality disorders such as situs inversus, heterotaxy, and congenital heart defects. Here, we identify CCDC57 as a regulator of ciliary function and LR patterning. Depletion of ccdc57 via morpholino oligonucleotides (MOs) led to abnormal cilia in the multiciliated cells of the embryonic epidermis of Xenopus. Additionally, LR markers, dand5 and pitx2c were misexpressed resulting in defects in normal rightward cardiac looping. Finally, we identified a patient with situs inversus carrying compound heterozygous CCDC57 missense variants. We tested these variants in Xenopus depleted of ccdc57. Wild-type human CCDC57 mRNA, but not the patient variants, rescued ciliary structure and…
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Fig. 5- —National Natural Science Foundation of Chinahttp://dx.doi.org/10.13039/501100001809
- —Yale Universityhttp://dx.doi.org/10.13039/100005326
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TopicsGenetic and Kidney Cyst Diseases · Developmental Biology and Gene Regulation · Congenital heart defects research
INTRODUCTION
The establishment of left–right (LR) asymmetry during embryogenesis is critical for proper development and function of internal organs (Hamada et al., 2002). For example, cardiac anatomy is asymmetric across the LR axis, which is critical for oxygenation of blood and delivery to tissues. Additionally, the stomach and the spleen are on the left side while the liver is located on the right. Abnormalities in laterality (known as heterotaxy) can lead to severe congenital heart disease, malrotation of the gut, and defects in splenic function. Surgical intervention remains challenging leading to significant morbidity and mortality (Cohen et al., 2007).
In early embryonic development, the function of the left–right organizer (LRO) is critical for robust LR development (Forrest et al., 2022). The LRO is conserved across multiple vertebrate species, including mice (Basu and Brueckner, 2008), frogs (Schweickert et al., 2007), and fish (Essner et al., 2005), although, interestingly, not in all vertebrates (Szenker-Ravi et al., 2025, 2022; Gros et al., 2009; Blum et al., 2009) The LRO is a transient, midline structure formed at the end of gastrulation posteriorly, encompassing two sets of monociliated cells. There, motile cilia generate leftward extracellular fluid flow, which is sensed by non-motile cilia at the lateral margins of the LRO (Boskovski et al., 2013; Tavares et al., 2017; Djenoune et al., 2023; Katoh et al., 2023). This leftward fluid flow leads to asymmetric expression of dand5 at the margins of the LRO (Maerker et al., 2021; Minegishi et al., 2021) resulting in Nodal signaling on the left and subsequently pitx2c expression in the left lateral plate mesoderm (Schweickert et al., 2000, 2010; Vonica and Brivanlou, 2007). Lateralized expression of pitx2c directs the asymmetric development of organs such as the heart (Bowers et al., 1996). Therefore, cilia are critical for breaking bilateral symmetry to establish proper LR patterning. Defects in this process can lead to situs inversus, heterotaxy, and congenital heart disease (Babu and Roy, 2013; Wallmeier et al., 2020).
Cilia are primarily composed of microtubules, ciliary associated proteins, and the ciliary membrane (Wallmeier et al., 2020; Hilgendorf et al., 2024). The basal body, located at the base of cilia, is responsible for cilia assembly and orientation. There are a variety of types of ciliated cells. For example, motile and immotile monociliated cells are located in the LRO, central nervous system, and many other organs. Multiciliated cells, which have an array of cilia on the cell surface, are present in the respiratory tract, brain ventricles, and female reproductive tract. In these contexts, multiciliated cells generate brisk extracellular fluid flow to clear debris or pathogens, circulate fluids and nutrients, or transport ova. Defects in cilia formation or movement are known to result in a spectrum of human diseases including recurrent infections, bronchiectasis, hydrocephalus, and infertility (Mitchison and Valente, 2017).
Recently, multiple studies have demonstrated that CCDC57 (coiled-coil domain containing 57) is essential for motile cilia function by regulating the association between basal body polarity and axonemal orientation to control directional ciliary beating (Pan et al., 2024). CCDC57 also regulates centriole duplication and mitosis by interacting with microtubules and the microcephaly protein CEP63, thereby influencing cell cycle progression and chromosome stability (Gurkaslar et al., 2020). ccdc57-knockout in zebrafish and mouse lead to hydrocephalus and spinal curvature, consistent with a ciliopathy, but a role for ccdc57 has not been described in LR patterning (Pan et al., 2024; Xie et al., 2023; Li et al., 2023).
In this study, we used Xenopus tropicalis as a model to investigate the relationship between ccdc57, cilia function and LR patterning. We found that when ccdc57 was depleted, the cilia in the multiciliated cells of the embryonic epidermis became abnormal, and cilia driven flow over the embryonic epidermis was significantly decreased. Depleted embryos exhibited various cilia-related phenotypes, including LR asymmetry defects and edema. Finally, we also identified an individual with situs inversus who carries compound heterozygous variants in CCDC57. Based on rescue experiments in Xenopus, we demonstrate that these variants cause a reduction in protein function.
RESULTS
Ccdc57 depletion causes edema and alters situs in Xenopus tropicalis
To test the impact of ccdc57 in early embryonic development, we depleted ccdc57 in X. tropicalis using two different methods: morpholino oligo (MO) and F0 CRISPR (Fig. 1A). We designed a MO targeting the start site of the ccdc57 mRNA transcript and two non-overlapping sgRNAs targeting different exons of ccdc57 (Crispr1 and Crispr2).
*Depletion of ccdc57 leads to edema and cardiac looping defects. (A) Illustration of the ccdc57 targeting strategy using both MO and CRISPR-Cas9. The MO blocks the start codon in the coding region, while two different CRISPRs target exons 11 and 16. (B) Normal tadpole heart looping (D-loop) and the abnormal heart loop (L-loop and A-loop) at stage 42 Xenopus. These are ventral views with anterior to the top and the red outline highlights the heart and outflow tract. For embryos with edema, the internal organs exhibit abnormal morphology, and heart looping cannot be scored. Scale bar: 500 μm. (C) The percentage of edema in Crispr1, Crispr2 and MO injected groups. ‘n’, represents the number of embryos. ****P<0.0001 according to Chi-Square test. (D) After selecting out the edema embryos, the percentage of abnormal heart looping in Crispr1, Crispr2 and MO injected groups. ‘n’, represents the number of embryos without edema. ***P<0.0001 according to Chi-Square test.
When we depleted ccdc57 using the MO (5 or 10 ng per embryo), the predominant phenotype was edema detectable after stage 38, which appeared similar to that seen with other examples of loss of cilia function (del Viso et al., 2012). 49.8% and 90.8% of embryos developed edema when injected with 5 or 10 ng of MO, respectively, compared to just 3.4% in uninjected controls (UIC) and 2.6% using the standard MO Control (Fig. 1B,C). Similarly, embryos injected with Crispr1 and Crispr2 developed edema in 27.7% and 14.6% of embryos, respectively, compared to control groups (UIC 3.4%, Crispr Control 3.4%) (Fig. 1B). Because of the edema, cardiac looping is difficult to score as the outflow tract is stretched and obscured. However, in the tadpoles without edema, we observed cardiac looping defects (7.4% Crispr1, 7.0% Crispr2, 21.1% 10 ng MO, 8.0% 5 ng MO compared to 1.0% UIC, 0.7% Crispr Control and 0.9% MO Control) (Fig. 1C).
To address specificity and efficacy of our ccdc57 depletion, we used multiple tests. First, we used Inference of CRISPR Edits (ICE) analysis to confirm CRISPR targeting (Conant et al., 2022). We amplified the cut site by PCR in either individual embryos or pools of embryos and deconvoluted Sanger traces to identify the size and percentage of indels. The percentage of INDELs is 80% and 89% for Crispr1 and Crispr2, respectively, while the out of frame INDEL percentage is 69% and 61% (Fig. S1). Additionally, CRISPR based depletion phenotypes are similar to those seen by a different depletion strategy, MOs. We also compared results to the standard control MO and to a Crispr Control (that targets tyrosinase which affects pigmentation). Finally, we demonstrate rescue of the MO using the human CCDC57 mRNA (see below).
Ccdc57 knockdown disrupts LR patterning
Due to the edema in ccdc57 depleted embryos, scoring cardiac looping was challenging. Therefore, to further determine if LR patterning was affected, we examined LR molecular markers. The LR signaling cascade begins at the LRO, which in Xenopus forms at stage 16 (Fig. 2A). At this point, cilia driven extracellular fluid flow is just initiating and dand5 is symmetrically expressed flanking the LRO (which we term ‘early dand5’). By stage 19, cilia driven flow has completed its signaling and dand5 is suppressed on the left (which we term ‘late dand5’). This asymmetry leads to pitx2c in the left lateral plate mesoderm that we can assay at stage 28 in Xenopus, which precedes any edema. Therefore, we began our analysis of global LR patterning with pitx2c.
*Depletion of ccdc57 alters global LR patterning. (A) Illustration of relevant stages for this experiment. Embryos were injected at 1-cell stage and then collected at stage 16 or 19 for dand5 and stage 28 for pitx2c. (B) The depletion of ccdc57 leads to pitx2c abnormalities. The leftmost panel (blue outline) shows the normal pitx2c expression in the left mesoderm. The other panels show examples of abnormal pitx2c expression as right sided (red), bilateral (green), or absent (purple). All views are ventral with anterior to the top. The arrow indicates pitx2c expression in lateral mesoderm, scale bar: 200 μm. (C) The graph depicts the percentage of embryos with abnormal pitx2c expression. ‘n’, represents the number of the embryos cumulatively in injection groups ****P<0.0001 (Chi-square test or Fisher's exact test). (D) At stage 16 (early), dand5 is expressed bilaterally in the LRO (leftmost box outlined in blue). Examples of abnormal dand5 expression are shown including R<L (green) and R>L (red). These are ventral views of dissected LROs with anterior to the top, scale bar: 100 μm. (E) The percentage of embryos with dand5 expression types at stage 16. ‘n’, represents the number of the embryos. ‘ns’ indicates no significant difference between the MO and UIC group. Fisher's exact test. (F) At stage 19 (post flow), the cilia-driven flow suppresses dand5 signal on the left side (leftmost box with blue outline). Embryos with abnormal expression included, R<L (green) and R=L (red). Scale bar: 100 μm. (G) The percentage of embryos with dand5 expression types at stage 19. ‘n’, represents the number of the embryos. Fisher's exact test. ***P<0.0001.
We collected embryos depleted of ccdc57 using either MO or CRISPR at stage 28 and performed in situ hybridization to detect pitx2c transcripts. We observed abnormal pitx2c expression in both groups, indicating disruptions in LR patterning. The percentage of abnormal pitx2c is 0.7% and 0% in UIC for CRISPR and MO, respectively, compared to 28.8% in Crispr1, 19.0% in Crispr2, 11.0% in 5 ng of MO and 18.9% in 10 ng of MO (Fig. 2B,C). These results suggested that global LR patterning is altered in ccdc57 depleted embryos.
Looking earlier in the LR cascade, at stage 16, we found that expression of dand5 in MO-injected embryos was similar to the UIC group (Fig. 2D,E). However, by stage 19, we observed significant expression abnormalities in MO-injected embryos. In 98.4% of stage 19 UIC embryos, dand5 signal was suppressed on the left (R>L) and only 1.6% of UIC embryos had abnormal R<L signal. In contrast, 38.2% of ccdc57 depleted embryos (10 ng of MO) had R>L expression of dand5, 35.3% showed bilateral R=L signal, and 26.5% displayed R<L expression (Fig. 2F,G). From these data, we conclude that global LR patterning is altered in ccdc57 depleted embryos and that normal asymmetry of dand5 is not established at stage 19 consistent with a defect in cilia function at the LRO.
Ccdc57 is required for cilia-driven fluid flow and ciliary structure
Given that ccdc57 depletion leads to global LR patterning defects and alters late dand5 (but not early dand5), we hypothesized that Ccdc57 affects ciliary function in embryos. To explore this further, we examined two cellular contexts in which cilia drive extracellular fluid flow – the Xenopus embryonic epidermis and the LRO. The Xenopus embryonic epidermis has multiciliated cells that drive extracellular fluid flow much like the human respiratory epithelium (Walentek, 2021; Collins et al., 2021).
First, we analyzed the extracellular fluid flow generated by the epidermal cilia at stages 23-24 with OCT, optical coherence tomography, imaging as we described previously (Kim et al., 2024). Using OCT imaging, we can readily detect the extracellular fluid movement across the embryonic epidermis driven by multiciliated cells (MCCs) by tracking extracellular particles (cellular debris) (Tang et al., 2019). We categorized the cilia-driven fluid flow into three groups: normal fluid flow, slow fluid flow (barely detectable fluid flow) and no fluid flow. (Fig. 3A-C, for examples of flow classes see Movies 1-3). In embryos depleted of ccdc57 by MO, the distribution of embryos in these categories shifted from normal flow towards slow flow, and as the MO concentration increased, the proportion of embryos with no fluid flow increased (Fig. 3D). In the UIC, 100% of the embryos exhibited normal fluid flow. In the 5 ng MO injection group, only 16.2% of embryos maintained normal fluid flow, while 70.3% showed slow fluid flow and 13.5% had no fluid flow. This trend became more pronounced in the 10 ng MO group, where normal fluid flow dropped to 5.4%, slow fluid flow was observed in 54.0% of embryos, and 40.5% showed no fluid flow.
*Depletion of ccdc57 alters epidermal cilia and LRO area. (A-C) The extracellular fluid flow driven by epidermal cilia is visualized using OCT imaging. Endogenous particles (cellular debris) are present within the intravitelline space, and their movement can be tracked by OCT in vivo. Temporal color coding depicts particle trajectory over time. The color bar represents color versus the corresponding frame number in the color-coded image. Based on the trajectory map over 300 frames, we classified the flow as normal flow (A), slow flow (B), or no flow (C), scale bar: 100 μm. (D) Based on OCT imaging, percentages of embryos with normal, slow, and no-flow in uninjected controls and ccdc57 depleted tadpoles. ‘n’, represents the number of the embryos, Chi-square test, ****P<0.0001. (E,F) Cilia are reduced in MCCs when ccdc57 is depleted with MO. Cilia were marked with anti-acetylated-tubulin (green) and actin marked with phalloidin (red). Zoomed-in cilia image is shown in the blue boxed area. Scale bars: 50 μm. (G,H) Cilia in the LRO stained by anti-arl13b (red) and actin via phalloidin (green) to mark the cell borders. Scale bars: 50 μm. (I) Total area of the LRO. t-test. ***P<0.0001. (J) Cell number in LRO. t-test. ‘ns’ indicates no significant difference. (K) Cilia number in LRO. t-test. ‘ns’ indicates no significant difference. (L) Number of cilia per cell. t-test. ‘ns’ indicates no significant difference.
Given the loss of fluid flow, we considered two possibilities: 1) either the cilia were largely abnormal structurally or 2) largely structurally normal but immotile. We collected embryos injected with MO and performed anti-acetylated tubulin immunofluorescence staining. We found that in the MO-injected groups, by gross inspection, the number of cilia per multiciliated cell appeared reduced and their length and morphology were abnormal (Fig. 3E,F). The number of MCCs appeared unchanged between the MO and the control groups, but the structure of the cilia appeared abnormal. We can see less cilia per multiciliated cell.
We co-injected the 10 ng MO with membrane RFP (which also labels the ciliary membrane) and examined ciliary structure and motility under confocal microscopy, comparing these embryos with control embryos injected with membrane RFP alone. Consistent with the OCT and immunofluorescence results described above, we observed cilia that were almost completely immotile. In addition, the number of cilia per cell was markedly reduced, and the remaining cilia appeared rigid rather than curved (Movies 4,5). Therefore, based on these results, we conclude that Ccdc57 is critical for cilia structure and motility.
Given the LR patterning defects in ccdc57 depleted embryos, we next wanted to determine if Ccdc57 was also responsible for the structure of monocilia within the LRO. Compared to the UIC group, MO-injected embryos exhibited an enlarged LRO area. However, there is no significant difference in total number of cilia and cells in the LRO (Fig. 3G-L).
Biallelic CCDC57 variants identified in a patient with situs abnormalities
In our database of patients suspected to have primary ciliary dyskinesia (PCD), we identified a 27-year-old East Asian male who was diagnosed with situs inversus at birth but did not exhibit neonatal respiratory distress or related phenotypes (Fig. 4A,B). In recent years, the patient has experienced recurrent sinusitis but does not have typical bronchiectasis or other lower respiratory tract phenotypes. The nasal nitric oxide was 430 ppb (in normal range). Whole-exome sequencing revealed that the patient carries compound heterozygous missense variants in CCDC57 [NM_198082:c.493G>A(p.G165S),NM_198082:c.1555A>G(p.S519G)], which was confirmed by Sanger sequencing (Fig. 4C). Variants are confirmed to be in trans through Sanger sequencing of the family members. We also excluded the possibility of pathogenic variants in other known PCD- or cilia-related genes.
*Dextrocardia in patient with CCDC57 variants. (A) The pedigree of the individual with CCDC57 variants. The arrow indicates the individual with situs inversus. (B) CT scan of the patient reveals dextrocardia. R, right. (C) Sanger sequencing also demonstrates the two variants of CCDC57 determined by exome sequencing. (D) The multiciliary beat frequency of the patient and healthy controls measured at 37°C. Each data point represents the average ciliary beat frequency within a random area. t-test. **P<0.001. (E) The domain prediction of the CCDC57 protein and the locations of the identified variants. The Alphafold3 prediction of the CCDC57 protein structure. The superposition of the PDB 3D Viewer reveals that the amino acid changes from the patient variants have an impact on the helical angles of the protein. Additionally, the position of the microtubule-binding domain also displays significant changes in the presence of the patient variants.
Next, we captured nasal ciliated cells, cultured them at 37°C, and observed ciliary movement under high-speed video microscopy (HSVM) also at 37°C. The videos showed no gross observable abnormalities in the cilia beating patterns. However, analysis using CiliarMove revealed that the patient's ciliary beat frequency was slightly lower compared to normal controls (Fig. 4D, Movies 6,7) (Sampaio et al., 2021).
We then performed cilia-related immunofluorescence staining (DNAH5, DNALI1, RSPH9, CEP164) on airway epithelial cells obtained from a healthy control and our situs inversus patient. The results showed no gross abnormalities in the outer dynein arms, inner dynein arms, radial spokes and basal body. (Figs S2,S3). According to the AlphaFold-3 structural prediction model, the identified variants are located within or at the boundaries of the coiled-coil domain. Both variants are predicted to alter the local structural stability and may affect the overall spatial organization of the protein (Fig. 4E).
CCDC57 variants are loss-of-function based on rescue
Our patient carries compound heterozygous variants, but the pathogenicity of these missense variants is unknown. To test whether these variants affected protein function, we injected 10 ng of ccdc57 MO into Xenopus embryos at the single-cell stage and assessed whether the introduction of human CCDC57 mRNA into one cell at the two-cell stage could rescue ciliary formation and function of the MCCs on the embryonic epidermis (Fig. 5A). In this scenario, ccdc57 is depleted throughout the embryos, while embryos are selected in which the human mRNA is expressed only on the right or left side. Therefore, by comparing the left and right side of the embryonic epidermis, we can test if the human CCDC57 mRNA rescues the MO depletion. Embryos were collected at stage 28, after which we observed the fluid flow on both sides of each embryo and examined epidermal cilia using anti-acetylated tubulin staining.
*Patient variants in CCDC57 affect protein function. (A) Illustration of injection scheme. The MO was injected at one cell stage depleting ccdc57 throughout the embryo, and at the 2-cell stage, the human mRNA (with or without patient variants) and GFP-tracer mRNA were injected in one cell and embryos selected with targeting to either the right or left side. (B) Cilia driven epidermal flow is visualized using red microspheres over the period of 20 s. (C,D) Cilia-driven epidermal flow visualized by OCT. Temporal color coding depicts particle trajectory over time. The color bar represents color versus the corresponding frame number in the color-coded image. Based on the trajectory map over 300 frames. (E,F) Anti-acetylated tubulin staining of the two sides of the same embryo, the cilia on the MO only side are reduced compared to the opposite side where the human mRNA is injected. (G) The immunofluorescence staining of anti-acetylated tubulin (green) and phalloidin (red) in both CCDC57-variant mRNA injected embryos. (H) Velocity of particles measured by OCT are calculated on both sides of the embryo and the ratio determined between the sides. Therefore, a ratio of 1 indicates that two sides have similar flow velocities (UIC and MO alone injected). UICs and MO both have a ratio of 1 as the velocity on both sides of the embryo are similar. In the case of the wild-type human mRNA, the mRNA injected side is rescued, and the ratio is greater than one. This is significantly different compared to the same experiment with patient variant mRNAs. t-test. *P<0.05. *P<0.01.
We found that, compared to the MO-only control side, the side co-injected with human CCDC57 mRNA exhibited significantly faster fluid flow (Fig. 5B-D, Movies 8,9). Acetylated tubulin staining revealed a rescue of cilia morphology on the side co-injected with human CCDC57 mRNA compared to the MO side (Fig. 5E,F).
Subsequently, we attempted to rescue the cilia phenotype by injecting CCDC57 mRNAs with patient variants (G165S-CCDC57-hmRNA and S519G-CCDC57-hmRNA). Under immunofluorescence microscopy, injection of the patient's variant RNA did not significantly improve the ciliary phenotype (Fig. 5G). We analyzed particle flow velocity on the rescued and non-rescued sides using OCT and calculated the velocity ratio between the two sides. The results indicated that, compared to human CCDC57 mRNA, injection of the patient's variant RNA failed to rescue particle flow velocity on the embryo surface (Fig. 5H).
DISCUSSION
CCDC57 belongs to the coiled-coil domain containing protein family. Proteins in this family are widely recognized to be involved in cilia structure (Becker-Heck et al., 2011; Hjeij et al., 2014). Pathogenic variants in multiple genes within this family are well-established causes for ciliopathy-related phenotype. Previous studies have shown that CCDC57 is primarily localized to the centrioles, where it plays a role in centriole replication and cell mitosis, and is associated with microtubule stability (Gurkaslar et al., 2020). In recent years, several studies using mouse and zebrafish models have suggested that Ccdc57 may be involved in the polarity of ependymal cilia, affecting cerebrospinal fluid flow and leading to phenotypes such as hydrocephalus and spinal curvature (Pan et al., 2024; Xie et al., 2023; Li et al., 2023). However, a role for ccdc57 in LR patterning was uncertain, and given that some proteins affect selected cilia, we sought to investigate whether ccdc57 also affects LR patterning.
In our ccdc57-depleted Xenopus model, we observed a range of cilia-related phenotypes, including defects in cardiac looping and global markers of LR patterning. Cilia labeling of MCCs showed a disrupted cilia structure (reduced number of cilia, abnormal morphology, and loss of motility) and OCT identified a significant reduction in fluid flow over the embryonic epidermis. These findings suggest that CCDC57 is an essential component of cilia that drives LR patterning and extracellular fluid flow as seen in the respiratory tract. While we detected clear abnormalities in the cilia of the epidermal MCCs, the cilia of the LRO looked largely normal, and therefore, we could not identify a likely reason for why late dand5 is abnormal in ccdc57 depleted embryos. Live imaging of the LRO may reveal abnormal cilia motility or orientation that alters LRO flow, a future research avenue we are actively pursuing. We also note that the LRO morphology was abnormal with a larger area, whether this plays a role in regulating dand5 at the LRO margin remains to be seen.
We also identified an individual with LR patterning defects who carried compound heterozygous variants in CCDC57 gene. Human RNA rescue experiments supported the hypothesis that these CCDC57 variants result in at least a partial impairment of ciliary function. Although he did not present with typical bronchiectasis, he had a long history of chronic sinusitis and the cilia beating frequency was reduced. Based on the comprehensive results of multiple assessments, although we confirmed that CCDC57 deficiency impairs ciliary function, a definitive diagnosis of PCD in the patient remains inconclusive. We have sought to identify additional cases of ciliopathy associated with CCDC57; however, to date, this remains the only case found across all accessible databases. We anticipate that further cases will be identified in the future. With the advancement of research in recent years, an increasing number of cilia-related genes have been discovered. Interestingly, for many of these patients, HSVM may show normal beating frequency, and TEM may not reveal significant structural abnormalities, yet these genes can still cause disease (Dougherty et al., 2020). In such cases, mucociliary transport (MCT) analysis is often required to assess ciliary beating patterns or polarity, providing insights into ciliary dysfunction (Wallmeier et al., 2021; Howes et al., 2024). Previous studies on Ccdc57 knockout mice showed impaired mucociliary transport function in airways, which is consistent with the reduced particle flow velocity observed via OCT in X. tropicalis in our study (Pan et al., 2024). This represents a new trend in the analysis and understanding of ciliary motility disorders, and OCT is an important tool for conducting MCT analysis (Tang et al., 2019).
Our work used the Xenopus model to demonstrate that depletion of ccdc57 through CRISPR-Cas9 targeting and MOs led to disrupted left-right patterning and abnormal cilia function. Additionally, through rescue experiments, we confirmed that the CCDC57 variants identified in our suspected individual are dysfunctional suggesting that CCDC57 may be a cause of the patient's PCD. Together, our work not only elucidates the essential role of CCDC57 in motile cilia function and LR axis formation but also underscore its relevance as a novel genetic contributor to human laterality disorders, paving the way for more precise molecular diagnostics of laterality defects and ciliopathies.
MATERIALS AND METHODS
Ethics and informed consent
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 the included individual and healthy controls. Healthy controls were recruited as healthy volunteers from the hospital, and all were screened to exclude the possibility of PCD or other cilia-related disorders.
Animal husbandry
X. tropicalis were housed and cared for in our aquatics facility according to established protocols that were approved by the Yale Institutional Animal Care and Use Committee (IACUC). Ovulations and IVFs were performed as previously described (Lane et al., 2022b; Lane and Khokha, 2022).
Microinjections of MO and mRNA in Xenopus
Microinjections of Xenopus embryos were performed per standard protocols (Lane et al., 2022a). 1.0 mm borosilicate glass needles (World Precision Instruments) were pulled to a fine taper and used to inject embryos at 1-cell stage with either 5-10 ng of translation blocking MO 5′-ATTCCTCCTCTTTTGGCAACATTTT-3′ (GeneTools) or 2 nl of a solution containing 400 pg of sgRNA plus 1.5 ng of Cas9 protein. For F0 CRISPRs, we incubated the sgRNA and Cas9 protein together briefly before injecting into the 1-cell Xenopus embryo. We used the EnGen sgRNA synthesis kit (NEB) to generate sgRNAs using the following oligos:
Crispr 1-taatacgactcactataGGTTATTTGTGAGTCTCACCgttttagagctagaa Crispr 2-taatacgactcactataGGCTCGAGGTTTTCTGGTACgttttagagctagaa.
We also included an MO-control group and a CRISPR control group to exclude potential off-target effects as well as the possible toxic effects of the MO and Cas9. The control MO is a standard control MO 5′-CCTCTTACCTCAGTTACAATTTAA 3′ from GeneTools. The control Crispr oligo was target to tyrosinase.
Crispr-Control – taatacgactcactataGGCTGTTGTAGGCAATCGGGgttttagagctagaa.
Injections were traced with either 90 pg of Dextran RUBY or Dextran Alexa Fluor 488 (Invitrogen). For live cilia imaging, 90 pg of membrane RFP was injected to label the cilia.
Post-injections, embryos were flooded with 3% Ficoll in 1/9× Marc's Ringer (MR) for 1 h for recovery. For CRISPR/Cas9 injection, embryos (including controls) were put into a 28°C incubator for 1 h. Then embryos were transferred into a dish containing 1/9× MR with 50 μg/ml of gentamicin and incubated at 22-26°C along with sibling control embryos.
Injections were confirmed by fluorescent lineage tracing with a Zeiss Lumar fluorescence stereomicroscope the day after injection, and tadpoles were incubated until the required stage.
Human reference CCDC57 (HsCD00296923, DNASU) was sub cloned into pCSDest2 vector using Gateway recombination techniques (LR clonase II, Thermo Fisher Scientific). To create variants, site-directed mutagenesis was conducted on wild-type human CCDC57 according to standard methods using the Q5 Site-Directed Mutagenesis Kit (New England BioLabs). All clones were confirmed by Sanger sequencing.
Capped mRNAs were generated in vitro by linearizing plasmids with an appropriate restriction enzyme and transcribing with the mMessage machine kit (mMESSAGE mMACHINE™ SP6 Transcription Kit Ambion), following the manufacturer's instructions.
ICE verification
CRISPR efficiency was determined through Sanger sequencing. CRISPR injected embryos were grown to approximately stage 43. Embryos were anesthetized by brief incubation in 2% tricaine, then transferred to individual tubes. Genomic DNA was extracted by immersing embryos in 100 μl of 50 mM sodium hydroxide per embryo, incubating at 95°C for 10 min, vortexing, then neutralizing by adding 20 μl of 1 M Tris pH 7.4. The region around the cut site was amplified using the following primers:
Crispr1-F: TAAACCACAAACCAGCGAAAC
Crispr1-R: AAACTTACATCTCCCACAAGA
Crispr2-F: TGTATGCTCACCTGCTCTGT
Crispr2-R: AAAGGTGCCAAGGGTTTCAA.
PCR products were purified using the Monarch PCR & DNA clean up kit, then sent for Sanger sequencing using the forward primer for each. Sequences were analyzed for insertion/deletion efficiency using the online ICE tool (Synthego, https://ice.editco.bio/#/).
Whole-mount in situ hybridization
In situ hybridization was performed according to standard protocols (Khokha et al., 2002). The details of the probes employed have been previously described (Deniz et al., 2023).
For dand5, embryos were raised to stage 16 (early dand5) and stage 19 (late dand5), then fixed in MEMFA overnight at 4°C. The embryos were washed 3×15 min with PBS. LROs were dissected and dehydrated with several washes from 25% to 100% ethanol, and then the in situ hybridization procedure was performed as previously described (Khokha et al., 2002).
For pitx2c, embryos were raised until stage 28, fixed with MEMFA over night at 4°C, dehydrated with several washes from 25% to 100% ethanol, then processed through the in situ hybridization procedure.
Immunofluorescence and microscopy
Human respiratory epithelial cilia
Nasal ciliated cells from the proband and a healthy control were obtained by nasal brush and fixed in 4% paraformaldehyde overnight at 4°C. Briefly, the slides were incubated overnight at 4°C with primary antibodies: DNAH5 (HPA037470, Sigma-Aldrich, 1:100), DNALI1 (ab155490, Abcam, UK, 1:100), RSPH9 (23253-1-AP, Proteintech 1:100), CEP164 (22227-1-AP, Proteintech 1:100), 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 DAPI (Vector Laboratories, #H-1200-10) 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).
LRO
Stage 16 embryos were fixed with 4% PFA in PBS for 2 h at room temperature, then washed three times with PBS. The LROs were dissected and incubated in the blocking buffer [3% bovine serum albumin (BSA) in PBS+0.1% Triton X-100] for 1 h at room temperature. LROs were incubated overnight with anti-ADP-ribosylation factor-like protein 13B (Arl13b clone N295B/66, NeuroMab) diluted 1:200 in blocking buffer at 4°C overnight. The LROs were washed 3×10 min with PBS, followed by incubation in anti-mouse Alexa Fluor 647 (Thermo Fisher Scientific) diluted 1:500 in blocking buffer for 2 h at room temperature. Then, specimens were washed 3×10 min with PBS and incubated in Alexa Fluor 488 Phalloidin (Thermo Fisher Scientific) diluted 1:100 in blocking buffer for 1 h at RT. The embryos were washed 2×10 min with PBS before mounting between two coverslips with ProLong Gold Antifade (Thermo Fisher Scientific) and imaged with a Zeiss 880 airyscan confocal microscope.
Epidermal cilia
Stage 26-28 embryos were collected for epidermal cilia staining. Embryos were fixed 4%PFA in PBS for 2 h in room temperature, then washed three times with PBS. The embryos were incubated in the blocking buffer for 1 h in room temperature and stained with mouse anti-acetylated tubulin (Sigma, cat. no. #T6793) diluted 1:1000 as the primary antibody overnight. The embryos were washed 3×10 min with PBS, followed by incubation in anti-mouse IgG Alexa Fluor 488 (Thermo Fisher Scientific) diluted 1:500 in blocking buffer for 2 h at room temperature. Then, embryos were washed 3×10 min with PBS and incubated in Alexa Fluor 647 Phalloidin (Thermo Fisher Scientific) diluted 1:100 in blocking buffer for 30 min at RT. The embryos were washed 2×10 min with PBS before mounting between two coverslips with ProLong Gold Antifade (Thermo Fisher Scientific) and imaged with a Zeiss 880 airyscan confocal microscope.
Whole-exome sequencing and Sanger validation
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) per the manufacturer's instructions and used for subsequent exon sequencing. Library capture, sequencing, and data analysis were performed by Novogene Bioinformatics Institute, Beijing, China as described previously (Guo et al., 2021). 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.
Sanger sequencing was performed in the patient to validate the variants. Primer sequences were designed using an online primer design tool.
High speed video microscopy analysis
Nasal ciliated epithelia from the patient and healthy control were obtained by nasal brush and ALI cultured at 37°C for 3 weeks. HSVM were photographed using an upright Olympus BX53 microscope (BX53, Olympus) with a 40× objective lens. Videos were recorded using a sCMOS camera (Prime BSI™, Photometrics) at a rate of 500 fps at 37°C. Only intact ciliated edges (>50 µm) were used for functional analysis. Seven separate ciliated epithelial strips from the mucus-free regions were measured. Ciliary beat frequency (CBF) was calculated using the validated automated open-source software (CiliarMove) (Sampaio et al., 2021).
Extracellular fluid flow imaging
To visualize cilia driven extracellular fluid flow, a Thorlabs Ganymede 900 nm spectral domain-OCT Imaging System was used as described previously (Tang et al., 2019). Stage 22-23 embryos (still within the vitelline membrane and without any evidence of edema) were collected to analyze under OCT and classified as ‘normal flow’, ‘slow flow’ and ‘no flow’ as described. To visualize extracellular fluid flow using beads, stage 28 embryos were collected and cultured in 1× PBS with 1.00 µm dyed carboxyl polystyrene microspheres (DCCR004, Bangs Lab), and subjected to OCT.
Quantification and statistical analysis
All Xenopus experiments were performed a minimum of three times, with each replicate a different clutch of embryos (biological replicates). To assess whether there were significant differences between groups, t-tests, Fisher's exact tests and chi-square tests were performed, as detailed in the respective figure legends. A P-value less than 0.05 was considered statistically significant.
Supplementary Material
10.1242/biolopen.062495_sup1Supplementary information
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