Mutation of ccdc57 results in scoliosis in zebrafish

In an ENU-based screen for zebrafish mutants with mineralized craniofacial and skeletal tissue defects, we identified tft^168N mutants that displayed severe spinal curvatures in adults (Fig 1A). Both Micro-CT and Alizarin red staining showed that tft^168N mutants displayed abnormal three-dimensional curvatures and deformities of the spinal vertebrae when viewed from both dorsal and lateral positions (Figs 1B and S1A). Interestingly, the severity of the spinal curvatures was comparable in male and female zebrafish mutants, in that the cobb angles measured from dorsal-ventral and medio-lateral curvature showed similar distribution patterns (S1B–S1E Fig). To identify the gene responsible for this phenotype, we performed Next Generation sequencing and identified a mutant locus at the genomic region encoding the ccdc57 gene. The ccdc57 gene contains 19 exons encoding 979 amino acids, and the tft^168N allele introduced a stop codon in the 11th exon of ccdc57 resulting in the truncation of the final 407 C-terminal amino acids (Fig 1C). To further validate this gene mutation, we generated ccdc57 mutants via CRISPR/Cas9 methods and recovered two mutant alleles, one with a 2-bp insertion (+2) and another with a 7-bp deletion (Δ7) in the first exon (Fig 1C). Both of these two ccdc57 mutant alleles displayed scoliosis (Fig 1D). Moreover, complementation testing between the Δ7 and tft^168N alleles showed that these two alleles failed to complement each other, confirming that mutations in ccdc57 were causative for scoliosis in these mutants (Fig 1D). Of note, when dissected from fixed vertebrae, ccdc57 mutant spinal cords also displayed curvature defects that closely mimicked those of the spinal curvatures (Fig 1E).

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Fig 1. Mutation of ccdc57 leads to scoliosis.

(A) Representative images of male and female wild type and tft^168N mutants. (B) Micro-CT images showing lateral and dorsal views of 3-months-old wild type and tft^168N mutants. (C) Genomic structure and sequences of wild type, tft^168N mutant and two ccdc57 mutant alleles generated with CRISPR/Cas9. The underlined sequence in wt indicates PAM sequence of sgRNA target. (D) Representative images of male and female wild type and ccdc57 mutants with different genetic background as indicated. (E) The dissected brain and spinal cord in wild type and ccdc57 mutant. Scale bars: 1 cm in panels A, B, D, E.

https://doi.org/10.1371/journal.pbio.3002008.g001

To define the progression of scoliosis in ccdc57 zebrafish mutants, we conducted developmental analyses of the mutant phenotype. Zygotic ccdc57^Δ7 mutants displayed body curvature at 3 days postfertilization (dpf) (S2A Fig). Interestingly, body curvature was not apparent in 3 dpf ccdc57^tft168N mutants (S2A Fig). Such disparity may be due to the presence of truncated forms of Ccdc57 proteins in the ccdc57^tft168N mutants, as the tft^168Nallele encodes the N-terminal 572 amino acids of Ccdc57 (Fig 1C). Moreover, maternal Ccdc57 protein may contribute to early embryonic development. Indeed, maternal zygotic (MZ) ccdc57^tft168N mutants displayed body curvature defects at both 3 and 5 dpf, similar to those of MZ ccdc57^Δ7 mutants (S2A Fig). In the following studies, we focused on the ccdc57^Δ7 mutant allele to evaluate the function of Ccdc57 in spinal development and scoliosis.

In zebrafish, ciliogenesis defects are a major cause of body curvature development [7]. We therefore examined cilia development in ccdc57 mutants. Surprisingly, all of the cilia examined were grossly normal in the ccdc57 mutants as visualized using anti-glycylated tubulin antibody (S2B–S2F Fig). Although abnormal notochord differentiation can contribute to the development of congenital scoliosis, we did not detect any notochord defects in ccdc57 mutants at early stages, suggesting an independent mechanism of scoliosis progression in the absence of Ccdc57 (S2G Fig). In fact, scoliosis was first detected at approximately 17 dpf in both ccdc57^tft168N and ccdc57^Δ7 mutants, with initial bending of the mutant spine occurring in the middle portion of the trunk (S2H and S2I Fig).

Hydrocephalus in ccdc57 mutants

Recently, abnormal CSF flow has been linked to the development of scoliosis [6,8]. Therefore, to better assess the etiology of scoliosis progression in ccdc57 mutants, we dissected whole brains from ccdc57 mutants and wild type sibling controls. The ccdc57 mutant brain was larger and appeared transparent due to an expanded ependymal epithelium filled with CSF (Fig 2A and 2B). Histological analysis of cross-sectioned brains revealed the presence of dilated ventricles in the optic tectum and rhombencephalon of ccdc57 mutants, as well as in the central spinal canal (Fig 2C–2E). In addition, micro-CT analysis confirmed the presence of dilated ventricles in ccdc57 mutant brains (Fig 2F).

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Fig 2. Hydrocephalus in ccdc57 mutants.

(A, B) External phenotypes of telencephalons in wild type and ccdc57 mutant. Arrows point to the expanded ependymal epithelia due to excess CSF accumulation. (C) Diagram of the brain tissue in zebrafish showing the sites of crossing section in different ventricles. CCe, corpus cerebelli; TeO, tectum opticum; CC, central canal; TecV, tectal ventricle; RV, rhombencephalic ventricle. (D) Histological cross-sections showing ventricles (arrows) at different sites as indicated. (E) Bar graph showing relative size of brain ventricles at different sites as indicated. (F) Micro-CT images showing the ventricles (arrows) in TecV and CC of wild type and mutant. (G) Diagram showing the position of diencephalic choroid plexus (dChP) in adult zebrafish. OB, olfactory bulb; Tel, telencephalon. (H, I) Three-dimensional reconstruction of dChP visualized with DAPI staining in wild type and ccdc57 mutant. These still images are from S1 Movie and visualized from ventral side of the dChP. A, anterior; P, posterior. Scale bars: 100 μm in panels A and B, 1 mm in panel D, and 50 μm in panels H and I. The data underlying the graphs shown in the figure can be found in S1 Data.

https://doi.org/10.1371/journal.pbio.3002008.g002

As described, CSF is produced by the CP. Therefore, we further examined the CP in wild type and ccdc57 mutants focusing on the epithelial monolayer connecting the telencephalon and optic tectum known as the forebrain CP or diencephalic CP (dChP) [36–38] (Fig 2G). Three-dimensional reconstructions of the dChP in wild type zebrafish revealed a chapeau-like structure with multiple folds covering the brain tissues (Fig 2H and S1 Movie). In contrast, ccdc57 mutant dChP appeared stretched and lacked foldings, due to excess CSF accumulation (Fig 2I and S2 Movie). Next, we investigated whether ccdc57 mutant larvae showed signs of hydrocephalus. Injection of Rhodamine- or FITC-conjugated fluorescent beads into 2 or 3 dpf zebrafish larvae revealed no obvious differences in the brain ventricle size between mutants and siblings (S3 Fig). Together, these data suggested that, similar to the late appearance of scoliosis, ccdc57 mutants developed hydrocephalus at later stages of development.

Planar polarity defects of cilia and basal bodies in ccdc57 mutant ependymal cells

Excess accumulation of CSF in cerebral ventricles is one of the major causes of hydrocephalus. In zebrafish, directed CSF flow is facilitated by the coordinated beating of motile cilia in multiciliated ependymal cells lining the brain ventricles. Focusing on the telencephalon and dChP regions, we identified multiciliated cells largely restricted to the central portion of the ependymal layer (S4 Fig). Next, we monitored motile cilia beating in these ependymal cells. In adult wild type zebrafish, all motile cilia bent to the direction of the fluid flow, and multicilia bundles beat synchronously (Fig 3A and S3 Movie). In contrast, multicilia bent in a variety of directions in ccdc57 mutants (Fig 3A and S4 Movie). Even within the same ciliary bundle, individual cilia beating appeared disorganized (S4 Movie). We next performed scanning electron microscopy (SEM) analyses to visualize the ultrastructure of the ependymal cilia bundles. Compared to the clustered distribution in wild type fish, ciliary bundles appeared highly disorganized in ccdc57 mutants (Fig 3B). Aberrant distribution of these ependymal cell ciliary bundles was further confirmed via immunostaining with anti-glycylated tubulin antibody to visualize cilia (Fig 3C).

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Fig 3. Mutation of ccdc57 results in planar polarity defects in ependymal cells.

(A) Still images showing the beating direction of motile cilia to drive fluid flow (pink arrow) in adult wild type and mutants. The representative kymographs of cilia movement were shown at the bottom. (B) Scanning electron microscopy showing the multicilia of the ependymal cells in wild type and ccdc57 mutant. (C) Confocal images showing cilia of the ependymal cells in wild type and ccdc57 mutant as indicated. Cilia were labeled with anti-mono-glycylated tubulin in green and the tight junction were stained with Claudin-5 antibody in red. Nuclei were counterstained with DAPI. (D) Confocal images showing the basal body distribution in the ependymal cells of wild type and ccdc57 mutant as indicated. The basal bodies were labeled with anti-γ tubulin in green and cytoskeleton was stained with phalloidin in purple. Tight junctions were stained with Claudin-5 antibody in red. Nuclei were counterstained with DAPI. Arrows indicate the displacement of basal bodies from the center of the cells. The schematic diagram of the displacement of basal bodies were shown at the bottom. (E) Angular distribution of the basal bodies in ependymal cells from wild type and ccdc57 mutant. (F) Statistical analysis showing relative displacement distance of basal bodies from center of the cell. (G) Statistical analysis showing the number of basal bodies in each ependymal cell of wild type and ccdc57 mutant. BB, basal body. (H) Model illustrating the distribution of multicilia on ependymal cells of wild type and ccdc57 mutant. All the data analyzed in this figure are generated from 3- to 4-months-old adult zebrafish. Scale bars: 5 μm in panels B and D; 7.5 μm in panel C. The data underlying the graphs shown in the figure can be found in S1 Data.

https://doi.org/10.1371/journal.pbio.3002008.g003

Cilia are anchored to the surface of ependymal cells through basal bodies. We therefore investigated basal body localization in wild type and ccdc57 mutant ependymal cells. In wild type zebrafish, basal bodies appeared localized to the same side of each ependymal cell (Fig 3D and 3E). In contrast, basal bodies were randomly distributed in ccdc57 mutants, with many localized to the center region of ependymal cells, indicating polarity defects in the absence of Ccdc57 (Fig 3D–3F). Notably, the number of basal bodies was comparable between mutant and wild type ependymal cells (Fig 3G). Next, we characterized the basal bodies of ependymal cells inside the ChP. In wild type zebrafish, multiciliated ependymal cells mainly localized to the center of the folds of the ChP (S5A and S5B Fig and S5 and S6 Movies). Of note, wild type ChP ependymal cells also exhibited planar polarity in basal body distribution (S5C Fig), and this planar polarity was absent in ccdc57 mutant ChP ependymal cells (S5C–S5E Fig). Together, these data suggested that loss of Ccdc57 resulted in abnormal distribution of multicilia in the ependymal cells, together with basal body planar polarity defects (Fig 3H).

Cell polarity defects of ccdc57 mutant ependymal cells

We next asked whether ependymal cell polarity was affected in the absence of Ccdc57. Since basal bodies rely on the cytoskeleton microtubule network to localize cilia, the misplacement of basal bodies in ccdc57 mutants may be due to abnormal organization of cytoskeletal microtubules. We therefore examined the cytoskeleton using alpha tubulin antibody staining and found that the microtubule skeleton appeared disorganized in ccdc57 mutants as compared to the highly polarized microtubule network in wild type cells (S6A and S6B Fig). The position of mitochondria also depends on the microtubule network, we further analyzed the distribution of ependymal cell mitochondria using Tom20 staining. Our results showed that ccdc57 mutant ependymal cells displayed abnormal mitochondria distribution patterns as compared with those of wild type cells (S6C and S6D Fig).

We next examined the shape of ependymal cells via Claudin-5 antibody-labeled cellular tight junctions. At 3 months postfertilization (mpf), wild type zebrafish ependymal cells appeared symmetrically distributed throughout the ependymal layer, with narrow and elongated cells located in the center and larger round cells located laterally (Fig 4A and 4B). Moreover, the elongated cell axes were parallel to the anterior–posterior axis, indicating polarity of these cells (Fig 4C). Intriguingly, the symmetrical distribution pattern of ependymal cells was also found to be established in 17 dpf wild type zebrafish larvae (Fig 4D–4F). In contrast, polarized ependymal epithelial cell organization was severely disrupted in 3 mpf ccdc57 mutants (Fig 4B and 4C). At 17 dpf when spine curvatures are first apparent, ccdc57 mutants also exhibited abnormal-shaped ependymal cells in the central region, suggesting an earlier defect of ependymal cell polarity (Fig 4E and 4F). Notably, since multiple motile cilia formation does not occur until 1 mpf (Fig 4G) [38], these results suggested that Ccdc57 participates in the regulation of ependymal cell polarity in early zebrafish development, as well as in later developmental basal body positioning.

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Fig 4. Loss of Ccdc57 results in ependymal cell polarity defects.

(A, B) Distribution pattern of ependymal cells in 3-months-old wild type and ccdc57 mutant as indicated with tight junction marker Claudin-5 staining. Nuclei were counterstained with DAPI in blue. The displayed region of ependymal epithelia was indicated in panel A. Heatmap showing the relative size of each ependymal cells as displayed in the confocal images. (C) Statistical analysis showing orientation of the midline ependymal cells along the anterior–posterior (A–P) axis in wild type and ccdc57 mutant. The angles between the longer axis of each midline ependymal cells and the A–P axis were used to evaluate cell polarity with 0 degree indicating horizontal cell. (D, E) Distribution pattern of ependymal cells in the brains of 17 dpf wild type and ccdc57 mutant. (F) Statistical analysis showing orientation of the midline ependymal cells in wild type and mutant as indicated. (G) Confocal images showing the ependymal layer labeled with Claudin-5 (red) and glycylated tubulin (green) antibodies in 25 and 30 dpf wt and ccdc57 mutants. Enlarged views of the boxed regions are displayed in the bottom. Nuclei were counterstained with DAPI. Scale bars: 25 μm in panel B; 50 μm in panel E; 50 μm in panel G. The data underlying the graphs shown in the figure can be found in S1 Data.

https://doi.org/10.1371/journal.pbio.3002008.g004

Ccdc57 orchestrates the synchronized beating of motile cilia in the spinal canal

During larval zebrafish development, ependymal cells are derived from radial glial cells, which also are involved in establishing their translational cell polarity [31]. Therefore, we next asked whether radial glial cilia were defective in the absence of Ccdc57. At larvae stages, spinal canal ependymal cells are a specialized type of radial glia harboring primary motile cilia that drive CSF flow. In ccdc57 mutants, the number and length of spinal canal cilia were comparable to those of wild type larvae (S2E and S2F Fig). Further examination of cilia localization by immunostaining with anti γ-tubulin antibody showed that both wild type and ccdc57 mutant cilia were localized to the posterior apical surface of each radial glia cell in the spinal canal, with no apparent differences in basal body localization (Fig 5A and 5B).

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Fig 5. Ccdc57 orchestrates synchronized beating of motile cilia in the central canal.

(A) Confocal images showing the localization of basal bodies in the floor plate cells at 2 dpf and 5 dpf. Basal bodies were stained with anti-γ tubulin (gama-Tub, green) antibody, and nuclei were counterstained with DAPI in blue. The white dotted line connects the center of cells. (B) Statistical analysis showing the angles of basal bodies with the anterior–posterior (A–P) axis as illustrated in the diagram. (C) Still images showing kymographs of floor plate cilia movement in 5 dpf of wild type and ccdc57 mutant. (D) Beating frequency of floor plate cilia in wild type and ccdc57 mutants. (E) Statistical analysis showing the rotation angles of cilia in floor plate. The anterior and posterior angles were measured as illustrated in the diagram. (F, G) Relative tilting directions of floor plate cilia in wild type and ccdc57 mutants. The tilting direction was evaluated by the bisector of each angles. (H) Confocal images showing relative localization of Dvl protein (green) and basal body (gama-Tub, red) in floor plate cells. (I, J) Subcellular localization of Dvl protein (green) in the floor plate ependymal cells. The statistical results were shown in panel J. (K) Confocal images showing the localization of GFP-Prickle (green) in floor plate ependymal cells as indicated. (L) Confocal images showing the distribution of Dvl protein (green) on ependymal cells of adult zebrafish. The displacement distance of Dvl vesicles was also shown on the right. Scale bars: 5 μm in panel A; 5 μm in panels H and I; 10 μm in panel K; 7.5 μm in panel L. The data underlying the graphs shown in the figure can be found in S1 Data.

https://doi.org/10.1371/journal.pbio.3002008.g005

We next examined the beating pattern of these spinal canal cilia using high speed video microscopy. By imaging cilia motility in the caudal central canal of 5 dpf wild type larvae, we found that all of the cilia beat in a similar manner, and a synchronized beating wave was easily observed throughout the field of view (Fig 5C and S7 Movie). However, the orchestrated ciliary beating was disrupted in ccdc57 mutant spinal cords, with individual cilia beating independently of one another (Fig 5C and S8 Movie). Of note, the beating frequency of individual cilia was comparable between ccdc57 mutant and wild type larvae (Fig 5D). Ciliary beating angle measurements demonstrated that the beating angle increased significantly in ccdc57 mutants as compared to wild type siblings (Fig 5E). We further used the bisector of each angle to evaluate the tilting direction of motile cilia and found that the tilting angles were significantly increased in ccdc57 mutants (cat. 72.11° ± 14.16° in wild type versus 85.52° ± 5.06° in mutants) (Fig 5F and 5G).

To further explore the beating defects of spinal canal cilia, we performed fluorescent bead tracing experiments at different stages of zebrafish development. By monitoring the movement of fluorescent beads injected into the central canal, we observed bidirectional CSF flow, as shown previously, in the central canal of wild type larvae at both 30 hpf and 3 dpf [39,40] (S9 and S11 Movies). The bidirectional CSF flow was largely maintained in ccdc57 mutants, although residential circular particle movements were also observed (S10 and S12 Movies). Strikingly, we noticed that the injected fluorescent beads were able to be transported to the end of the central canal by 6 hours postinjection (hpi) in the majority of wild type larvae, while none of the ccdc57 mutants contained fluorescent beads in the caudal spinal canal, and the fluorescent bead transport distances were significantly reduced (S7 Fig). The bead transport defects were observed in ccdc57 mutants at all stages analyzed (3 dpf, 5 dpf, and 17 dpf) (S7 Fig). Together, these data suggest that Ccdc57 orchestrates the synchronized beating of spinal canal motile cilia, whose deficiency leads to abnormal CSF-flow.

Localization of core PCP components in ccdc57 mutants

The posterior tilting of spinal canal motile cilia is mainly regulated by the PCP pathway [41]. To further characterize this phenomenon, we examined the localization of two major PCP pathway components, Prickle and Dishevelled 1 (Dvl1), in wild type and ccdc57 mutant fish. Similar to their basal bodies, Dvl1 appeared localized to the apical posterior region of each radial glia cell (Fig 5H), with Dvl1-positive vesicles localized to the region surrounding the basal bodies (Fig 5H). Noticeably, the Dvl1 distribution angles were comparable between ccdc57 mutants and wild type control embryos (Fig 5I and 5J). The localization of Prickle was opposite to that of Dvl1 vesicles, in the anterior apical surface, as demonstrated by GFP-Prickle labeling. The Prickle localization was similar in wild type and ccdc57 mutants (Fig 5K). These data suggested that localization of PCP components in larval zebrafish was not affected by the absence of Ccdc57. Intriguingly, in adult zebrafish, Dvl1 expressing vesicles appeared localized to one side of mature ependymal cells, corresponding to the location where multicilia formed (Fig 5L). In contrast, in adult ccdc57 mutants, Dvl1 expressing vesicles appeared dispersed randomly throughout the entire cell, clearly indicating a late onset cell polarity defect in ccdc57 mutants (Fig 5L).

CCDC57 encodes a centrosomal satellite protein required for cell polarity

To reveal the mechanisms of Ccdc57 regulation of basal body positioning, we further examined the subcellular localization of CCDC57 in RPE-1 cells. Similar to previous reports, CCDC57 mainly localized to centriolar satellites in RPE-1 cells [42]. Flag-tagged CCDC57 protein colocalized with centriolar satellite proteins OFD1 and PCM1 and did not localize to cilia (ARL13B) or to the distal appendage of the basal body (CEP164) (Fig 6A–6C). Mass spectrometry analysis of pulled-down proteins using Anti-FLAG conjugated beads showed that CCDC57 interacted with multiple centrosomal proteins, including CEP170, OFD1, and CP110 (Fig 6D), and we further validated the CCDC57-OFD1 interactions via immunoprecipitation (Fig 6E). Moreover, siRNA knockdown of OFD1 eliminated localization of CCDC57 to the centriolar satellite. In contrast, siRNA knockdown of CCDC57 gene expression had no effect on centrosomal localization of OFD1(Fig 6F).

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Fig 6. CCDC57 is a centrosomal satellite protein required for cell polarity.

(A-C) Confocal images showing the relative localization of CCDC57 with OFD1, PCM1, CEP164, and ARL13B in RPE-1 cells. (D) Mass spectrometry analysis of the CCDC57 interacting proteins. (E) Immunoprecipitation results showing the interaction between CCDC57 and OFD1. (F) Confocal images showing the localization of CCDC57 and OFD1 in siRNA knockdown RPE-1 cells. (G) Scratch-wound assay showing the polarized localization of Golgi (GM130, green) during directional cell migration in control and CCDC57 siRNA-treated cells. (H) Model illustrating the statistical analysis of cell polarity by Golgi position. (I) Dot plots showing the angles of Golgi facing the migration edge in control and CCDC57 siRNA-treated cells. (J, K) Images showing the cell migration state and statistical analysis of the migration distance. In all panels, nuclei were counterstained with DAPI in blue. Scale bars: 2.5 μm in panel A; 5 μm in panel B; 3 μm in panel C; 5 μm in panel F; 50 μm in panel G. The data underlying the graphs shown in the figure can be found in S1 Data.

https://doi.org/10.1371/journal.pbio.3002008.g006

During migration, cells can establish a front–rear polarity characterized by the polarized distribution of Golgi and centrosomes in the leading edge [43]. We therefore further tested the role of CCDC57 in establishing cell polarity in RPE-1 cells by wound-scratch assay. After scratching, the microtubule cytoskeleton of leading edge RPE-1 cells appeared polarized, with the Golgi facing toward the scratched space to direct cell migration. In control siRNA-treated RPE-1 cells, the majority of leading edge cells contained Golgi apparatus located within 60 degrees of the direction of migration relative to the nucleus (Fig 6G–6I). In contrast, the polarized orientation of Golgi apparatus was significantly compromised in CCDC57 siRNA-treated RPE-1 cells, as demonstrated by significantly increased orientation angles (Fig 6G–6I). Furthermore, migration distance was also reduced in CCDC57 siRNA-treated cells as compared to control siRNA-treated cells (Fig 6J and 6K). Together, these in vitro studies further confirmed the role of the centrosomal protein, CCDC57, in establishing cell polarity.

Abnormal RF assembly and urotensin expression in ccdc57 mutant larvae

We next sought to identify the causes of the body curvature observed in ccdc57 mutants. Epinephrine signals are essential for urotensin expression and body straightening [8]. In line with this, epinephrine treatment was also able to rescue body curvature in ccdc57 mutant embryos (S8 Fig). The RF is essential for body axis straightening through transferring the epinephrine signals to the CSF-cNs [20,21]. We found that wheat germ agglutinin (WGA) can be used to label and image the RF (S9A–S9C Fig). In various ciliary mutants, WGA-labeled RF appeared either discontinuous or absent (S9D–S9G Fig). Similarly, WGA staining of the RF was also diminished or absent in ccdc57 mutant larvae (Fig 7A). Interestingly, body curvature severity closely correlated with the severity of RF assembly defects in ccdc57 or kif3a mutants (S9H–S9M Fig). Next, we examined the expression of urotensins in ccdc57 mutant larvae using whole-mount in situ hybridization (WISH) analysis. While urp1 expression appeared relatively normal in the anterior trunk of ccdc57 mutant larvae, the number of urp1-expressing cells was markedly decreased in the posterior trunk (Fig 7B and 7C). Together, these data suggested that Ccdc57 deficiency interrupts assembly of the RF and down-regulates the expression of urotensin genes. The fact that differences in urotensin gene expression mainly occurred in the posterior trunk may explain why ccdc57 mutant larvae exhibit only mild body curvature.

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Fig 7. Mutation of ccdc57 results in RF defects and abnormal expression of Urotensins in the spinal cord.

(A) Representative images of Reissner fiber (RF, white arrows) in 2 dpf wild type and ccdc57 mutant larvae. cc, central canal; nc, notochord; fp, floor plate. (B) Whole-mount in situ hybridization results showing the expression of urp1 in 24 hpf control and ccdc57 mutant larvae as indicated. The enlarged views of the staining in the posterior region are shown in the bottom. (C) Statistical analysis showing the number of urp1-expressing cells in the anterior and posterior part of the trunk. (D) Confocal images showing RF (white arrow) in wild type and ccdc57 adult mutants. RF was stained with wheat germ agglutinin (WGA, red), and nuclei were counterstained with DAPI. (E) qPCR analysis showing the expression of urotensin genes (urp1 and urp2) in different parts of the adult trunk as illustrated in the diagram. (F) In situ hybridization results showing the expression of urp2 in the brains of adult wild type and ccdc57 mutant as indicated. Arrows point to the expression of urp2 in the posterior part of the brain. (G) In situ hybridization results showing the expression of urp2 in the spinal cord of wild type and ccdc57 mutant as indicated. The purple arrowhead indicates the sites of enriched urp2 expression. (H) The line graphs showing the relative optical density of the expression of urp2 in wild type and ccdc57 mutants. (I) Model illustrating the distribution of urp2 in wild type and ccdc57 mutant. Scale bars: 7.5 μm in panel A; 10 μm in panel D. The data underlying the graphs shown in the figure can be found in S1 Data.

https://doi.org/10.1371/journal.pbio.3002008.g007

Ectopic accumulation of Urotensin neuropeptides in ccdc57 adult mutants

Next, we attempted to discover the relationship between the development of scoliosis and CSF flow defects. First, we investigated the assembly of the RF in wild type and ccdc57 mutants, based on the previously characterized roles for the RF in regulating body axis development [10,19,22]. In wild type adults, the RF appeared thick and straight (Fig 7D). In contrast, the RF appeared much thinner, discontinuous, and/or absent in adult ccdc57 mutants (Fig 7D). Of note, the severity of RF assembly defects correlated with the level of scoliosis in the mutants (S10A and S10B Fig). Next, we focused on the expression of urotensin neuropeptides. Intriguingly, we found that from a lateral view, adult ccdc57 mutant spines always contained a strong dorsal bending in the anterior trunk and a second dorsal bending in the tail region (Fig 1B and 1E). To better characterize this feature, we dissected adult wild type and ccdc57 mutant trunks into three segments (Head, Middle, and Tail) and isolated total RNA from each segment (Fig 7E). Unexpectedly, qPCR results showed that the expression of urp1 and urp2, the major urotensins regulated by CSF signaling, was increased over approximately 10 times in the Head segments of ccdc57 mutants as compared with those of wild type control siblings (Fig 7E). In contrast, Middle trunk segments displayed lower expression levels of these genes in ccdc57 mutants, while urotensin expression again appeared up-regulated in Tail segments of ccdc57 mutant adult spines (Fig 7E). To further validate these results, we performed ISH analysis on dissected spinal cords for the expression of urp2, one of the major urotensin genes expressed at later stages. Our results showed that the expression of urp2 was dramatically increased in the brainstem region of ccdc57 mutants (Fig 7F). In contrast, urp2 expression was virtually absent in the middle part of the spine, with some urp2 expression observed in the tail region (Fig 7G and 7H). To further examine whether urp2 expression in the tail region correlated with the second dorsal bending of the spine, we dissected the spinal cord at the second bending site and further cut it at the apex into anterior and posterior fragments (S10C Fig). Interestingly, compared with wild type control siblings, urp2 gene expression was up-regulated in the posterior fragment of the dissected spinal cord, while urp2 was not detected in the anterior fragment (S10C Fig). Together, these results suggest an interesting relationship between urp2 expression and spinal curvature (Fig 7I).

Correlation between urotensin expression and spine curvature in ciliary mutants

To further gauge the relationship between urotensin expression and spinal curvature, we evaluated spine curvature phenotypes in several zebrafish scoliosis mutants. Both tmem67 and ofd1 mutants displayed scoliosis. Micro-CT results showed that these mutants also displayed initial dorsal bending in the anterior spine (Fig 8A). The observed scoliosis in ofd1 and ccdc57 mutants further demonstrated the interactions between Ofd1 and Ccdc57 that were revealed by our IP pulldown experiments (Fig 6E). Of note, qPCR results confirmed the enhanced expression of urp1 and urp2 in the anterior spines of ccdc57, tmem67, and ofd1 mutants (Fig 8B).

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Fig 8. Ectopic accumulation of urotensin neuropeptides is associated with spine curvature in zebrafish ciliary mutants.

(A) Representative images of wild type and scoliosis zebrafish mutants. Micro-CT images are shown on the bottom. (B) qPCR analysis showing the expression of urotensin genes in the heads of wild type and scoliosis mutants. (C) Enlarged views of the head regions of wild type, ccdc57, and uts2r3 mutants. (D, E) Statistical analysis of the dorsal curvature angles in different mutants as indicated. The angles were measured between the direction of the parasphenoid bone and the Weberian vertebrae as illustrated in the diagram (D). (F) In situ hybridization results showing the expression of urp2 in the brains of wild type and uts2r3 mutant as indicated. (G) Expression of urp2 in the spinal cords of wild type and uts2r3 mutant. The strongly increased expression of the urp2 in the tail region was shown on the enlarged views. (H) Dissected spinal cords from wild type and uts2r3 mutants as indicated. The two boxed regions correspond to those in panel G. Scale bars: 1 cm in panels A and H. The data underlying the graphs shown in the figure can be found in S1 Data.

https://doi.org/10.1371/journal.pbio.3002008.g008

We have previously shown that mutation of uts2r3, the major Urotensin receptor, leads to scoliosis. Noticeably, the spinal curvature of uts2r3 mutants was clearly different from that of ciliary mutants. The anterior dorsal bending phenotype appeared relatively minor in uts2r3 mutants, while all uts2r3 mutants bent to the ventral side first, and displayed strong dorsal bending in the posterior portion of the trunk (Fig 8A). To better characterize spinal bending, we measured the angle between the parasphenoid bone and the Weberian vertebrae orientation in wild type, ccdc57, and uts2r3 mutants (Fig 8C). The measured angles of ccdc57 mutants were significantly larger than those in control and uts2r3 mutants (Fig 8C–8E). Consistent with these results, urp2 expression levels were dramatically increased in the anterior spines of ccdc57 mutants, while only a slight increase was observed in uts2r3 mutants (Fig 8B), as validated via WISH assay (Fig 8F). Noticeably, the location of increased urp2 expression in uts2r3 mutant spines also correlated with the observed second bending in uts2r3 mutant spines (Fig 8G and 8H). Together, these data provide strong evidence that spinal bending closely correlates with the activation of Urotensin signaling.

Abnormal urotensin signals in idiopathic scoliosis patients

Finally, we sought to investigate whether urotensin signaling is also involved in the regulation of spinal curvature in human scoliosis patients. Although it is difficult to obtain scoliosis patient spinal cord tissue for analysis, we were able to collect bilateral paravertebral muscle tissue from scoliosis patients during surgery. We then compared Urotensin signaling pathway gene expression in paravertebral muscle tissue harvested from the convex and concave sides of spinal curvature sites (Fig 9A). Strikingly, although UTS2 expression appeared similar between convex and concave muscle tissue locations, we observed a remarkably asymmetric expression of UTS2R in bilateral paravertebral muscles of AIS patients (n = 46) (Fig 9B and 9C). The expression of UTS2R was significantly higher in convex side as compared to concave side muscle tissue (Fig 9B). According to the ratio of UTS2R expression in the convex versus the concave (convex/concave), we classified these AIS patients into two groups with the ratio of 2 as cutoff point. Strikingly, patients with >2-fold difference in expression (n = 26) had remarkably severe curvature magnitude as compared to patients with <2-fold difference (n = 20) (58.62 ± 9.48 degrees versus 51.35 ± 6.58 degrees) (Fig 9D and 9E). Thus, these data strongly support the hypothesis that abnormal Urotensin signaling may make a significant contribution to the severity of spinal curvature observed in AIS patients.

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Fig 9. Abnormal urotensin signals in adolescent idiopathic scoliosis (AIS) patients.

(A) Diagram indicating the position of bilateral paravertebral muscles harvested from AIS patients. (B, C) Relative expression of UTS2R and UTS2 in bilateral paravertebral muscles of AIS patients (n = 46). (D) Distribution of Cobb angles within the two groups of AIS patients exhibiting a ratio of UTS2R expression on the convex versus concave sides greater (>) or less (<) than 2. (E) Representative X-ray images showing two AIS patients with convex/concave ratios greater (5.21) or less than 2 (1.24). Measurements of Cobb angles are also indicated. The data underlying the graphs shown in the figure can be found in S1 Data.

https://doi.org/10.1371/journal.pbio.3002008.g009

In summary, here we showed that Ccdc57 is a centrosomal protein required for the proper establishment of basal body and cilia polarity. Loss of Ccdc57 disrupted the planar polarity of ependymal cells, affected the polarity and beating pattern of cilia present in both radial glial and ependymal cells, and led to CSF flow defects. The CSF flow defects resulted in the up-regulation of Urotensin signals in the anterior portion of the spine, eventually leading to spinal dorsal bending and scoliosis.

Ethics statement

All zebrafish studies were conducted according to standard animal guidelines and approved by the Animal Care Committee of Tufts University and Ocean University of China. Human tissue collection was approved by the Ethics Committee of the Nanjing University Medical School Affiliated Nanjing Drum Tower Hospital, China (No. 2019-066-01).

Zebrafish strains and mutants

All zebrafish strains were maintained at 14-hour light/10-hour dark cycle at 28.5°C. The tft^168N mutant line was identified from an ENU-based mutagenesis screen conducted in the Yelick Laboratory, Tufts University. The chromosomal location of the tft^168N allele was mapped by the Goessling Laboratory, Harvard-MIT Division of Health Sciences and Technology to an interval on Chromosome 12 containing 3 premature stop codon gene mutations, including a tgt/tga nucleotide substitution resulting in a predicted C/* mutation at amino acid 573 of ccdc57. Further analysis via quantitative real-time PCR (qRT-PCR) validated this homozygous mutation in phenotypic tft^168N mutants. Further validation of the ccdc57 mutation and scoliosis phenotype was performed via generation of CRISPR/Cas9 ccdc57 mutants. Zebrafish mutants were generated using CRISPR/Cas9 method with the following target sequences: ccdc57: 5′-GGGAAGAGGTCAGTGAGCTT -3′, ofd1: 5′-TATCAGACCTTCAAGAGCCG-3′, tmem67:5′-GGCAAGTGTCAGTGTCCTGA-3′. Cilia mutants (MZkif3a, ift74, ift88, and uts2r3) and Tg (urp1: GAL4; UAS: Kaede) transgenic lines were the same as previously reported [8,9,55–57].

Spinal cord and brain dissection

To dissect zebrafish spinal cords, humanely killed adult zebrafish were fixed in 4% of paraformaldehyde (PFA) overnight at 4°C. After washing 3 times with 1X PBS (phosphate buffered saline (PBS)) for 20 minutes each, the skin, muscle, and neural arches were carefully removed with tweezers, and the spinal cord was separated from the vertebral bodies. For brain dissection, zebrafish were first humanely killed using Tricaine in ice-cold water, then the skulls were removed with tweezers in 1X PBS to expose the brain. The dissected brain and spinal cord were further processed for WISH according to standard protocols.

Alizarin red and Calcein staining

Adult zebrafish were fixed in 4% PFA at 4°C for 1 week. After a 1-hour wash with PBST (0.1% Tween20), fixed zebrafish were stained with 0.01% Alizarin red (Sigma) in 1% KOH for another week. Next, the samples were washed in PBST for 3 days (1 wash/day) at room temperature and further cleared with 0.5% trypsin digestion for 24 hours at room temperature. After multiple washes with PBST, the skin was manually removed and the skeleton was imaged on ZEISS stemi 508 microscope. For Calcein staining, live zebrafish were incubated in 0.2% Calcein (Sigma) (pH 7.5) for 15 minutes and then washed twice with system water. In vivo stained zebrafish were anesthetized with 0.01% tricaine methanesulfonate (MS222), mounted in 3% methyl cellulose, and imaged using a fluorescent Leica M165FC microscope.

Micro-CT and vibratome sectioning of brain tissue

To visualize mineralized skeletal structures via Micro-CT, zebrafish were first anesthetized with 0.01% tricaine methanesulfonate (MS222). Micro-CT images were captured using a PerkinElmer Quantum GX2 microCT scanner. For histological analysis, the dissected brain was fixed in 4% PFA overnight at 4°C and then embedded in 3% low melting agarose. Transverse serial sections through the brain were collected using Leica vibratome vt1000s at a thickness of 90 μm each and imaged using Leica M165FC microscope.

High-speed video microscopy

To record the motility of ependymal cell cilia, the brain was dissected from the adult zebrafish, and the ependymal cell layer was gently peeled away from the top of the telencephalon with tweezers. The ependymal tissues were positioned on a glass cover slip containing 1X PBS and placed upside down in the center of a depression slide. Cilia movement was recorded using a 100X oil objective on a Leica Sp8 confocal microscope equipped with a high-speed camera (Motion-BLITZ EoSens mini1; Mikro-tron, Germany). Recordings of in situ cilia movement in the posterior portion of the spinal canal of 5 dpf larvae were conducted as previously described [58]. In all experiments, cilia movement was captured at a rate of 500 frames per second, and playback was set at 25 frames per second. Image processing was performed using ImageJ software (National Institutes of Health, Bethesda, MD, USA).

Scanning electron microscopy

Ependymal epithelia were dissected from the brain, fixed in 2.5% glutaraldehyde overnight at 4°C, then washed 3 times with 1X PBS. Next, samples were serially dehydrated to 100% ethanol and transferred into isoamyl acetate. After critical point drying, samples were sputter coated with gold–palladium alloy before imaging. Images were acquired using Hitachi-3400N scanning electron microscope (Hitachi, Tokyo, Japan).

Immunofluorescence assay

Immunostaining of whole-mount larvae was performed using standard protocols [59]. For immunofluorescence of ependymal cells, the brain was first dissected from adult zebrafish and then fixed in Dent’s fixative (80% methanol and 20% dimethylsulfoxide). Fixed brains were incubated with primary antibodies overnight at 4°C, followed by washing 3 times with PBST at 4°C, then incubated with secondary antibody overnight at 4°C. Immunostained adult zebrafish ependymal cell layer tissues were carefully collected with tweezers for imaging. The ependymal cell tissues of zebrafish less than 1 month old were imaged directly and did not require dissection from the brain.

For RF staining, 48 hpf zebrafish larvae were fixed in Dent’s fixative overnight at 4°C, washed 3 times with 1 X PBS, then incubated with WGA dissolved in PBS (1:200) overnight at 4°C. After 4 times wash with 1X PBS, the samples were imaged with Leica Sp8 confocal microscope. For staining in adult zebrafish, the spinal cord was first dissected, fixed in 4% PFA overnight at 4°C, and further processed for WGA staining. After staining, the tissues surrounding the spinal cord were removed to expose the RF for imaging.

The following antibodies were used: anti-glycylated tubulin (1:500, EMD); anti-γ-tubulin (1:500, Sigma); anti-cldn5 (1:250, Thermo Fisher Scientific); anti-DVL1 (1:200, BiCell Scientific); anti-TOM20 (1:500, Abcam); anti-alpha-tubulin (1:500, Sigma); anti-polyglutamylated tubulin (1:500, Adipogen); and WGA (1:200, Invitrogen).

Cell culture

HEK293T cells were cultured in Dulbecco’s Modified Eagle Medium-high glucose (D6429, Sigma-Aldrich), and RPE-1 cell lines were cultured in Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12 (D8437, Sigma-Aldrich). Both media were supplemented with 10% fetal bovine serum (FSP500, ExCell Bio) and 100 IU/mL penicillin/streptomycin (15140122, Gibco).

Plasmid and siRNA transfection

Transfections of HEK293T cells were performed using polyethyleneimine (PEI) (24765–2, Polysciences). Transfections of siRNA were performed using Lipofectamine RNAiMAX (Thermo Fisher, 13778150). All transfections were performed according to the manufacturer’s instructions. The final concentrations of control siRNA (D-001810-10, Dharmacon) and CCDC57 siRNA (L-021433-02, Dharmacon) were 30 nM.

Lentiviruse package and infections

The pLV-Flag-CCDC57 plasmids were cotransfected with package plasmids psPAX2 (Addgene #12260) and pMD2.g (Addgene #12259) into HEK293 cells. Supernatant media containing lentivirus was filtered through a 0.45-μm membrane filter and added to plated RPE-1 cells in the presence of 5 μg/ml polybrene (sc-134220, Santa Cruze). After 48 hours, the infected cells were selected with 10 μg/mL puromycin for 14 days in vitro culture.

Immunofluorescence microscopy of cultured cells

The methods used were as previously described [60]. Briefly, cells grown on coverslips were fixed with 4% PFA for 10 minutes at room temperature and then permeated by cold methanol for 10 minutes at −20°C. After washing with PBS, the cells were incubated with the primary antibodies at room temperature for 1 hour. After washing twice with PBS, cells were incubated with secondary antibodies and 4′, 6-diamidino-2-phenylindole (DAPI) for 1 hour in dark. After washing 3 times with PBS, the coverslips were mounted with mounting buffer (S36963, Invitrogen). All antibodies were diluted in blocking buffer (1% BSA in PBS). Images were acquired using laser scanning confocal microscope (FV3000, Olympus) with a 40× oil-immersion objective. ImageJ software was used for quantification. For IF, the following antibodies were used: mouse anti-flag (1:1,000, Sigma-Aldrich); rabbit anti-PCM1(1:1,000, Cell Signaling Technology); rabbit anti-CEP164 (1:4,000, Proteintech); and OFD1 (1:5,000, homemade).

Coimmunoprecipitation and mass spectrometry analysis

HEK293T cells transfected with pLenti-Flag-CCDC57 were lysed in lysis buffer (20 mM Tris–HCl (pH = 7.5), 150 mM NaCl, 1 mM EDTA, 1 mM NaF, 0.3% Triton X-100) with 1% protease inhibitor cocktail (K1007, APExBIO). After centrifugation at 15,000g for 10 minutes at 4°C, the supernatant was incubated with FLAG M2 beads (A2220, Sigma-Aldrich) for 2 hours at 4°C, followed by 3 washes with lysis buffer.

Proteins were eluted from FLAG M2 beads using 0.2 mg/mL FLAG peptides (F3290, Sigma-Aldrich), mixed with SDS sample buffer and resolved using SDS-PAGE. Finally, protein bands were excised from the SDS-PAGE gels, digested with trypsin, and the extracted peptides were analyzed using mass spectrometry.

Western blot analysis

Proteins were separated by 10% SDS-PAGE and transferred to NT nitrocellulose membranes (66485, BIOTRACE). After blocking with 5% milk, membranes were incubated with anti-FLAG (1:5,000; F1804; Sigma-Aldrich) and anti-OFD1 (1:30,000) for 1 hour at room temperature. After washing with PBS 3 times, the membranes were incubated with secondary antibodies for 1 hour at room temperature. The ChemiDoc Touch Imaging System (Bio-RAD) and Odyssey Laser Imaging System (LI-COR) were used for image acquisition.

Wound healing assays

RPE1 cells transfected with negative control or CCDC57 siRNAs were seeded on glass coverslips in 12-well plates. When reaching 100% confluence, the culture medium was replaced with serum-free media. After 24-hour starvation, 1 mL pipette tips were used to generate scratches on the glass coverslips, and PBS was used to wash away the detached cells. Subsequently, cells were cultured in serum-free medium for an additional 6 hours and then fixed with 4% PFA. Golgi apparatuses were stained with GM130 (1:500; M179-3, MBL). Cells with Golgi apparatuses located in the sector facing the wound were considered positive for migration. Three groups (>100 cells per group) were used for quantification. GraphPad Prism software was used for statistical analysis. Quantitative data were presented as mean ± standard deviation (SD), and Student t tests were performed to determine statistical significance.

Pharmaceutical treatments and whole-mount in situ hybridization

Epinephrine (Sigma) treatment was performed as previously described [8]. Briefly, mutant and control zebrafish embryos were incubated with epinephrine (10 mg/ml) from bud stage until analysis. Images were captured using a Leica M165FC microscope. WISH was performed according to standard protocols. The following primers were used for in situ hybridization: urp1 forward: 5′-ACATTCTGGCTGTGGTTTG-3′, reverse: 5′- TGTATGGGGAAAACAAAGG-3′; urp2 forward: 5′-CAGCCCAAATAACAGAGACAAGAG-3′, reverse: 5′- AGAGGGTCAGTCGTGTTATTGAGG -3′.

Quantitative PCR

The expression levels of urotensin genes were evaluated using qRT-PCR analysis. Guided by the curvature of the spines, scoliosis mutant trunks were cut into three segments to include the highest or lowest portions of each spinal curve. Total RNA was isolated from dissected larvae or adult tissues using Trizol (Takara). cDNA was synthesized using PrimeScript 1st strand cDNA Synthesis Kit (Takara). qPCR was performed on the Step One real-time PCR system (Thermo Scientific) using the Eva-Green Master Mix (ABM). The following primers were used for PCR analysis: urp1 forward: 5′-TCTGGCGGTGCTCTACATTC-3′, reverse: 5′- AGCAGGACAGGAAGCACAGT-3′; urp2 forward: 5′- CCGGAGAACCAGATGCCTTT −3′, reverse: 5′-ATTTGGGCTGCTTGTTGCTG-3′. Relative gene expression levels were quantified using the comparative Ct method (2^−ΔΔCt method) based on Ct values for target genes and zebrafish β-actin.

Fluorescent beads tracing experiments

To evaluate hydrocephalus in zebrafish larvae, Rhodamine- or FITC-conjugated fluorescent dye (70 kDa) were injected directly into the brain ventricles at 2 or 3 dpf. Briefly, ccdc57 mutants and control siblings were first anesthetized using 0.01% tricaine and then incubated with 20 mM 2,3-butanedione monoxime (BDM, Sigma) for 6 minutes to stop the heart beating. The larvae were then placed on the surface of a 1% agarose plate and further injected with fluorescent dye. Fluorescence images were collected using Leica M165FC fluorescent microscope. The size of each brain ventricle was measured and quantified using ImageJ software.

For fine particle movement analysis, we injected 20 nm or 100 nm fluorescent beads (F8888, Thermo Fisher Scientific) into the central canal of 30 hpf (20 nm) or 3 dpf (100 nm) zebrafish larvae. The injection methods were similar to previously reported [39,40]. Briefly, 30 hpf or 3 dpf larvae were anesthetized using 0.4 mg/ml tricaine and then mounted in 1.5% low melting point agarose in the lateral position. The 20-nm or 100-nm fluorescent beads were injected into the center of the diencephalic ventricle. At 1 hour after injection, time-lapse images were acquired at room temperature on an inverted Leica DMI8 spinning disk confocal microscope equipped with an Andor iXon Life 888 EMCCD using a 40X water immersion objective (N.A. = 1.1). These images of the fluorescent beads in the rostral part of central canal were acquired at a frame rate of 10 Hz using Fusion software. The data were further analyzed with ImageJ software.

To examine the migration of fluorescent beads in central canal, we microinjected 100 nm fluorescent beads into the central canal at the position above the end of the yolk extension. The migrated distance of fluorescent beads was captured by THUNDER Imager Model Organism and quantified using ImageJ software.

3D reconstruction of the choroid plexus

The adult brain was dissected and fixed in 4% PFA overnight at 4°C. After immunofluorescent staining described above, the dChP was imaged using a 40X water objective on a Leica Sp8 confocal microscope. For 3D reconstruction of the CP, we first performed image binarization for acquired image stacks using ImageJ software (version 1.52p) and then performed surface rendering of the binary stack using Imaris software (version 9.6.0) to obtain 3D model.

Cell polarity analysis

ImageJ software was used to quantify cell polarity. For basal body angle analysis, angles were measured between a line made between the centers of the cell and basal bodies and the horizontal axis, as illustrated in Fig 3E. The displacement distance ratio was calculated by dividing the distance between the center of the cell and the center of the basal body by the distance from the cell center to the cell membrane. Cell orientation and centers were calculated by MorphoLibJ plugins, and the raw data were analyzed by Microsoft Excel software to generate graphs.

Human tissue collection

Paraspinal muscles were collected from 46 female AIS patients with main thoracic curve during corrective surgery. Bilateral deep paraspinal muscle biopsies of 1.5 × 1.5 × 1.5 cm³ were collected at the apical vertebral of the main curve for all the subjects. All patients or their guardians provided informed consent for the tissue collection.

Statistical analysis

Statistical analysis was performed using ImageJ, Microsoft Excel, or GraphPad Prism 6 software. All data were presented as mean ± SD as indicated in the figure legends. A value of p < 0.05 was considered statistically significant. The numerical data used in all figures are included in S1 Data.