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IK is essentially involved in ciliogenesis as an upstream regulator of oral-facial-digital syndrome ciliopathy gene, ofd1

Cell & Bioscience volume 13, Article number: 195 (2023) Cite this article

Abstract

Background

The cilia are microtubule-based organelles that protrude from the cell surface. Abnormalities in cilia result in various ciliopathies, including polycystic kidney disease (PKD), Bardet-Biedl syndrome (BBS), and oral-facial-digital syndrome type I (OFD1), which show genetic defects associated with cilia formation. Although an increasing number of human diseases is attributed to ciliary defects, the functions or regulatory mechanisms of several ciliopathy genes remain unclear. Because multi ciliated cells (MCCs) are especially deep in vivo, studying ciliogenesis is challenging. Here, we demonstrate that ik is essential for ciliogenesis in vivo.

Results

In the absence of ik, zebrafish embryos showed various ciliopathy phenotypes, such as body curvature, abnormal otoliths, and cyst formation in the kidney. RNA sequencing analysis revealed that ik positively regulated ofd1 expression required for cilium assembly. In fact, depletion of ik resulted in the downregulation of ofd1 expression with ciliary defects, and these ciliary defects in ik mutants were rescued by restoring ofd1 expression. Interestingly, ik affected ciliogenesis particularly in the proximal tubule but not in the distal tubule in the kidney.

Conclusions

This study demonstrates the role of ik in ciliogenesis in vivo for the first time. Loss of ik in zebrafish embryos displays various ciliopathy phenotypes with abnormal ciliary morphology in ciliary tissues. Our findings on the ik–ofd1 axis provide new insights into the biological function of ik in clinical ciliopathy studies in humans.

Background

Primary cilia are microtubule-based organelles that protrude from eukaryotic cell surfaces. In most cell types, primary cilia are highly dependent on the phase of cell cycle because they are dynamically regulated during cell cycle progression. They begin to form in the G0 and G1 phases, disassemble typically from the S/G2 phase, and finally undergo rapid resorption before entry into mitosis [1]. Primary cilia function as cellular antennae and intermediate external stimuli to spatiotemporally determine cell fate in the homeostasis of tissue development [2, 3]. Thus, dysfunction of primary cilia in tissues or organs leads to a variety of genetic pleiotropy and syndromic disorders, commonly termed ciliopathies, such as polycystic kidney disease, Bardet–Biedl syndrome, nephronophthisis, and retinitis pigmentosa [4,5,6,7,8]. Ciliopathies are induced by various factors, including genetic mutations, loss of intraflagellar transport proteins, improper docking of basal body, and abnormal pre-mRNA splicing of ciliary component genes [9,10,11,12,13,14]. Although an increasing number of human diseases are attributed to ciliopathies, causative factors and their mechanistical and molecular functions in ciliary defects are largely unexplored in vivo. Therefore, elucidating the molecular correlations and functions of various genes associated with ciliopathy are challenging.

The nuclear protein IK, also called as RED because of existence of sequence repeated arginine(R), glutamic acid(E) and aspartic acid(D) residues, is necessary for cell progression in mitosis and attributes in genomic stability by contributing to pre-mRNA splicing. Structurally, IK binds with SMU1 and the interacted two proteins perform the coordinated functions as a dimer complex with stabilizing reciprocally. Thus, lack of one protein makes to degrade reciprocal protein and leads to perturbed cellular function [15].In mitosis, IK recruits a core mitotic checkpoint protein mitotic arrest deficient 1 (MAD1) at the spindle poles and functions in mitotic progression of the spindle assembly checkpoint [16, 17]. Furthermore, IK-depleted cells show suppressed activity of PP1/PP2A which subsequently contributes to the sustained activity of Aurora kinase A, a member of mitotic serine/threonine kinase [18]. Meanwhile, IK also known to be required for activation of spliceosomal B complexes as it functions in the pre-mRNA splicing of short introns [19]. Although the expression level of ik is different in various tissues, the precise role of ik in specific tissues remains still unclear. Homozygous knockout of ik in mice results in embryonic lethality and we previously demonstrated that depletion of ik impairs the development of skeletal muscle in vivo using zebrafish model [20]. However, further studies on the physiological role of IK in vivo need be more required.

Oral-facial-digital 1 (ofd1) is the first gene reported in oral-facial-digital syndrome (OMIM 311200) [21,22,23,24] and encodes OFD1 protein located in the centrosome and basal body of primary cilia [21]. During ciliogenesis, it modulates the length of centriole and formation of distal appendages in the mother centriole [25,26,27]. Thus, the absence of ofd1 induces the abnormal elongation of distal regions of centrioles, and the long centrioles form structurally destabilized microtubules with abnormal post-translational modifications [27]. In human, mutations in ofd1 result in abnormal ciliary morphology and occurs OFD syndrome characterized by malformations of the face, oral cavity, and digits [28,29,30]. The patients with OFD syndrome present with several features common to ciliopathies, such as cystic disease and patterning abnormalities [28]. To date, the studies of ciliary dysfunction in the absence of OFD1 has been described using diverse in vitro and in vivo models [25, 27, 31,32,33,34]. In zebrafish, loss of ofd1 results in bent body axes, edema, and perturbed intravascular fluid flow in Kupffer’s vesicles with shorted and disrupted axonemes in cilia [31]. However, the molecular mechanisms controlling the expression of ofd1 beyond cilium assembly are poorly understood.

Here, we found that IK is positively associated with ofd1 expression, which is required for cilium assembly. Depletion of IK downregulates ofd1 expression and restoration of ofd1 expression overcomes IK-depletion-induced ciliary defects. Our findings show that IK–OFD1 axis is critical for normal ciliary development in vivo and provide new biological insights into clinical human ciliopathy.

Results

Zebrafish ik mutants display ciliopathy-like phenotypes

Zebrafish is considered to be the most suitable in vivo model to investigate the function of ik because homozygous knockout of ik is embryonic lethal in mice but not in zebrafish embryos. To explore the function of IK in vivo, we examined the spatiotemporal expression of ik during embryogenesis in zebrafish using whole-mount in situ hybridization (WISH) and quantitative RT-PCR (Fig. 1A, Additional file 1: Fig. S1). ik was expressed from 1-cell close to the blastoderm margin, but the overall maternally expressed level was significantly low (approximately 14 somites). However, after the period of maternal to zygotic transition (MZT) (approximately 21 h after fertilization in zebrafish), ik expression became intense (after 1 day post fertilization), specifically in organs from the mesodermal lineage, including the head, pronephros, and muscle. During the zygotic stage, ik was ubiquitously distributed in the tailbud and became progressively enriched, particularly in ciliated tissues such as the head and pronephric duct (Additional file 2: Fig. S2). These spatiotemporal expression analyses show that ik expression is much higher at the zygotic stage than at the early stage and imply that IK may have a functional role in ciliated tissues of zebrafish. Next, to characterize the role of IK in vivo, we analyzed the phenotypes of previously generated homozygous ik knockout zebrafish mutants using CRISPR/Cas9 system [20] and observed that ik mutants exhibited ciliopathy phenotypes. At 2 days post fertilization (dpf), ik mutants showed a distinct body curvature of 100% (n = 112) and abnormal otolith phenotypes such as fused or multiple form (83%; n = 93 out of 112) (Fig. 1B). Also, cyst formation in pronephros was observed in 80% (n = 90 out of 112). In addition to these phenotypes, the randomized left–right (LR) axis of the heart looping is known as a major ciliopathy phenotype caused by defects in motile cilia in Kupffer’s vesicles during early development of zebrafish [35] because ciliated organ Kupffer’s vesicles determine the LR symmetry of the brain, gut, and heart at the early stage. To determine the alteration in the LR asymmetry patterning in ik mutants, we performed WISH analysis to detect LR asymmetry in the heart using cardiac myosin light chain 2 (cmlc2) as a marker for the whole heart of zebrafish embryos. Wild-type (WT) embryos and ik mutants showed no differences in LR organization, with a correct S-shaped loop (D-loop) (Fig. 1C). These results indicate that loss of ik causes major ciliopathy phenotypes but is not involved in determining LR asymmetry. Next, we depleted ik expression in zebrafish embryos using morpholino antisense oligonucleotides (MOs) targeting exon 2 at the one-cell stage to perform the rescue experiment (Fig. 1D, E) [20, 36]. ik morphants revealed various pleiotropic ciliopathy phenotypes, including body axis curvature (74.6%; n = 50 out of 67), otolith defects (50%; n = 11 out of 22), and pronephric cysts (85.71%; n = 30 out of 35) (Fig. 1F). To verify the specificity of ik MO knockdown, we additionally co-injected synthetic mRNA encoding full-length wild-type ik with MO. The co-injected ik mRNA rescued all defective ciliary phenotypes of ik morphants revealing the possibility of off-target effects was excluded.

Fig. 1
figure 1

zebrafish ik mutants display ciliopathy-like phenotypes (A) Whole-mount in situ hybridization (WISH) analysis of zebrafish ik mRNA at different developmental stages (1-cell, 4-cell, 256-cell, Germ-ring, Bud, 14 somites, 1 dpf, 2 dpf, 3 dpf, and 4 dpf). Scale bar, 250 μm (B) Brightfield microscopic images of the body curvature, otolith, and pronephric cysts pronephros in WT embryos and ik mutants (ik −/−) at 2 dpf. Pronephric cyst of ik mutants is marked with red arrowhead. Stacked bar graph displays the percentage of embryos with a ventrally curved body, abnormal otolith phenotype, and cyst formation (C) WISH analysis of cmlc2 for the location of whole heart in WT embryos and ik mutants at 2 dpf (D) Schematic representation of the coding region of zebrafish ik. The target region (exon 2) of the morpholino employed in this study was represented using a blue line (E) RT-PCR analysis of ik mRNA expression in control morphants (MO), ik MO, and ik MO/ik mRNA co-injected embryos at 2 dpf. gapdh served as a normalization control and band intensity ratio of ik/gapdh mRNA expression was marked (F) Brightfield microscopic images of control MO, ik MO, and ik MO/ik mRNA co-injected embryos at 2 dpf. Whole body, otolith, and kidney cyst were captured. Pronephric cysts of ik MO are marked with red arrowhead

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Loss of ik results in abnormal ciliary morphology

To determine the mechanism underlying ciliopathy phenotypes in ik-depleted embryos, we next focused on studying the impact of ik on ciliogenesis and abnormal ciliogenesis in hair cells was confirmed in ik mutants by immunostaining for ac-α-tubulin, a specific marker of cilia. In hair cells of ik mutants, the number of cilia was reduced by approximately 50% of WT embryos, and their length was shortened by approximately 30% (Fig. 2A). Furthermore, we used Tg(brn3c:GFP) zebrafish line with ciliary bundles on hair cells labeled with green fluorescent protein (GFP). Because the brn3c gene, a member of the POU (Pit-Oct-Unc) domain transcription factor brn3 subfamilies (brn3a, brn3b, and brn3c), is selectively expressed in auditory and vestibular hair cells, the Tg(brn3c:GFP) line serves as an appropriate model for studying cilia in hair cells. We observed ik morphant Tg(brn3c:GFP) embryos showed a decrease in ciliary length and bundle number in hair cells (Fig. 2B). However, when full-mRNA of ik was co-injected into ik morphant Tg(brn3c:GFP) embryos, the abnormal ciliary phenotypes of ik morphant Tg(brn3c:GFP) embryos were almost completely rescued compared to those of the control group. Next, the effects of ik depletion on ciliogenesis were examined in the pronephros. ik mutants showed randomly misdirected cilia in pronephros, and we interestingly observed that disturbed cilia of the pronephros in ik mutants were specifically in the anterior but not in the posterior region (Fig. 2C). To determine whether aberrant ciliogenesis is owing to the alteration of the ultrastructure of motile cilia, we analyzed the organization or composition of axonemes of motile cilia using a transmission electron microscope (TEM). Similar to that of the WT embryos, ik mutants also had 9 + 2 arranged cilia in the pronephros (Fig. 2D, Additional file 3: Fig. S3). The result indicates that the abnormal ciliary morphology owing to the depletion of ik is not because of the damage to axoneme arrangement. Similar to that in ik mutants, 58% ik morphants showed randomly misdirected cilia in pronephros, and this abnormality was rescued by co-injection of ik mRNA (Fig. 2E). ik morphants also exhibited aberrant cilia only in the anterior region and not in the posterior region similar to ik mutants. These results show that IK functions specifically in the anterior region and not in the posterior region in the pronephros.

Fig. 2
figure 2

Loss of ik results in abnormal ciliary morphology (A) Whole-mount immunostaining of cilia in the inner ear (white boxes) using anti-acetylated-α-tubulin (green) of WT embryos (n = 3) and ik mutants (n = 5) at 2 dpf. Scale bar, 20 μm. Quantified graphs of cilia lengths and numbers in the inner ear between WT embryos and ik mutants at 2 dpf. *p < 0.05, ***p < 0.001 (B) Confocal microscopic images of control MO, ik MO, and ik MO/ik mRNA co-injected Tg(brn3c:GFP) embryos at 3 dpf. Scale bar, 20 µm. Quantification of ciliary length and number in the inner ear of control MO (n = 12), ik MO (n = 17), and ik MO/ik mRNA co-injected embryos (n = 10). **p < 0.01, ***p < 0.001 (C) Whole-mount immunofluorescence (left) of overall cilia in the pronephros in WT and in ik mutants at 2 dpf using anti-acetylated-α-tubulin (green). Scale bar, 100 µm. Enlarged view (right) of pronephric cilia in proximal and distal tubules stained with anti-acetylated-α-tubulin (green) and nuclei stained with DAPI (blue) at 2 dpf WT embryos and ik mutants. Scale bar, 20 µm. Stacked bar graph displays the percentage of embryos of cilia phenotype in the anterior pronephric duct of WT embryos and ik mutants at 2 dpf. (D) TEM results showing the ultrastructure of cilia in the pronephric duct of ik mutants. Cross-section showing the “9 + 2” configuration (black circles). Scale bar, 100 nm (E) Whole-mount immunofluorescence (left) of overall cilia in the pronephros of control MO, ik MO, and ik MO/ik mRNA co-injected embryos at 3 dpf with anti-acetylated-α-tubulin (green). Scale bar, 100 µm. Enlarged view (right) of pronephric cilia in the anterior and posterior pronephric ducts stained using anti-acetylated-α-tubulin (green) and nuclei stained with DAPI (blue) at 2 dpf control MO, ik MO, and ik MO/ ik mRNA co-injected embryos. Scale bar, 20 µm. Stacked bar graph displays the percentage of embryos based on ciliary morphology in the anterior pronephric duct (percentage of abnormal ciliary morphology of embryos: control MO, 0%; ik MO, 58%; ik MO + ik mRNA, 0%)

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ik mutants show impaired kidney development with abnormal ciliary morphology

Tissues with ciliary defects generally show impaired development [37, 38] and abnormal primary cilia cause impaired kidney development, such as autosomal recessive polycystic kidney disease (ARPKD) [39]. Because ik mutants also revealed abnormal ciliogenesis in the pronephros, they may be defective in kidney development. Thus, we measured the spatial expression of the segment marker genes of the proximal tubule to examine whether the development of proximal tubule is abnormal. The expression of slc4a4 in the proximal convoluted tubule (PCT) and slc13a1 in the proximal straight tubule (PST) was reduced in ik mutants (Fig. 3A). Interestingly, the spatial mRNA expression of slc4a4 in ik mutants showed shorter Y-shaped PCT structure than that of the WT. And spatial mRNA expression of the PST marker slc13a1 also showed a shorter PST structure with weaker expression than that of the WT. However, the expression of distal late segment marker slc12a3 was slightly increased in ik mutants, and the spatial expression of slc12a3 showed an unaltered structure compared with that of the WT. The levels of segmental marker genes were confirmed by reverse transcription polymerase chain reaction (RT-PCR) (Fig. 3B). This result implies that ik mutants have abnormal kidney development, showing impaired ciliary morphology specific to the anterior but not the posterior region. foxj1a, essential gene for motile cilia formation in the kidney, is rapidly induced in response to impaired kidney development or cyst formation [40]. Therefore, we examined whether foxj1a expression was increased in ik mutants with abnormal kidney development and cyst formation. RNA sequencing (RNA-seq) analysis revealed that foxj1a expression was increased in ik mutants by 1.8 at log2-fold changes relative to that of the WT embryos (Fig. 3C), and this enhanced expression was also confirmed by RT–PCR (Fig. 3D). Additionally, the spatiotemporal expression of foxj1a was analyzed by WISH at 1.5–2.5 dpf (Fig. 3E). At 1.5 dpf, foxj1a expression was upregulated in ciliary tissues, including the pronephros, in ik mutants compared to that in WT embryos, and it was continuously maintained at 2.5 dpf. Next, we performed rescue experiments using co-injection of ik mRNA into ik morphants to examine whether overexpression of ik directly affected foxj1 expression. Unlike ik morphants showing significantly enhanced foxj1a expression, ik morphants co-injected with ik mRNA showed foxj1a expression similar to that of the WT embryos (Fig. 3F).

Fig. 3
figure 3

ik mutants show impaired kidney development with abnormal cilia morphology (A) WISH analysis of pronephric duct markers slc4a4, slc13a1, and slc12a3 in WT embryos and ik mutants at 2 dpf. Scale bar, 250 μm (B) RT-PCR analysis of slc4a4, slc13a1, and slc12a3 expression in WT embryos and ik mutants at 2 dpf. gapdh served as a normalization control. Relative bands intensities of slc4a4, slc13a1, and slc12a3 normalized to gapdh intensity was graphically represented (in triplicates) using Image J software. **p < 0.01, ***p < 0.001 (C) Linear filter model showing correlation of overall gene expression based on RNA-seq data between WT and ik mutants at 3 dpf. The point of foxj1a plot is indicated. (D) RT-PCR analysis of foxj1a expression in WT embryos and ik mutants at 1.5 and 2.5 dpf. gapdh served as a normalization control. Relative band intensity of foxj1a normalized with respect to gapdh intensity is graphically represented (in triplicates) *p < 0.05, **p < 0.01 (E) WISH analysis of foxj1a in WT and ik mutants at 1.5, 2, and 2.5 dpf. Scale bar, 250 μm (F) WISH analysis of foxj1a in control MO, ik MO, and ik MO/ik mRNA co-injected embryos at 2.5 dpf. Scale bar, 250 μm

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ofd1, a necessary gene for cilia assembly, is downregulated by ik depletion

As ik was found to be essential for proper ciliogenesis with the development of ciliary tissues, RNA-seq data were analyzed to identify cilia assembly-related genes that were differentially expressed between WT and ik mutants. The log2-transformed transcriptome expression in RNA-seq of ik mutants was analyzed according to the strand-specific index with statistical significance set at p < 0.05. Differentially expressed genes involved in cilium assembly (Gene Ontology(GO):0060271) were also identified (Fig. 4A). Among the filtered cilium assembly genes, including ift57, ift172, dzip1l and ofd1, we focused on ofd1 gene with 0.59 at log2-fold changes as a target of ik, because the previously known physiological phenotypes of ofd1 mutants were considerably similar to ik mutants showing ciliopathy such as body curve, otolith abnormalities and a wide spectrum of malformations [31]. Indeed, ofd1 expression was downregulated in ik mutants (Fig. 4B) and was spatially reduced in the head and pronephros (Fig. 4C). To exclude the possibility of off-target effects, rescue experiments were additionally performed. The downregulated ofd1 expression in ik morphants was enforced by co-injection of ik mRNA as indicated by RT-PCR (Fig. 4D) and WISH experiments (Fig. 4E). This means that ofd1 is a downstream target of ik in vivo. Next, we performed the rescue experiment in vitro to confirm that IK functions as a transcriptional upstream regulator of OFD1. IK was knockdowned in human RPE cells, and wildtype-IK or mutant-IK (T485A; mutation of threonine to alanine cannot be phosphorylated) was re-transfected (Additional file 4: Fig. S4). Similar to the in vivo results, the OFD1 mRNA level was downregulated in IK-knockdowned (KD) cells and rescued by wildtype-IK. In contrast, when the mutant-IK (T485A) was transfected into IK-KD cells, the OFD1 mRNA expression still remained at low level without rescue. These results provide compelling evidence that IK is a transcriptional upstream regulator of OFD1. Previously, as IK was also known to play a role as a pre-mRNA splicing factor [19], we analyzed whether the downregulated ofd1 expression in zebrafish ik mutants was induced by pre-mRNA splicing error. As a result, we observed that the ofd1 pre-mRNA was normally spliced in ik mutants showing identical sequence with WT (Additional file 5). Furthermore, IK negatively regulates Aurora kinases, which are not only involved in the cell cycle but also play a role in cilia-associated signaling [16,17,18]. Thus, we investigated the expression of aurora kinases to examine the possibility that the downregulation of ofd1 in ik mutants might be due to aurora kinases. In RT-PCR analysis, the mRNA level of aurora kinase a and b showed no difference between WT and ik mutants (Additional file 6: Fig. S5A). Moreover, knockdown of Aurora kinase A or B in RPE cells also did not affect to expression of OFD1 at protein level (Additional file 6: Fig. S5B). This implies that the downregulation of odf1 in ik mutants occurs as a direct effect of ik, independent of aurora kinases.

Fig. 4
figure 4

ofd1, a necessary gene for cilia assembly, is downregulated by the loss of ik (A) Heat map of seven differentially expressed cilium assembly genes at least 1.5 > log2 fold changes in ik mutants relative to those of WT embryos. Upregulated DEGs relative to the mean are indicated by red color. Downregulated DEGs are shown by blue color (scale bar, log 2 of mRNA ratio) (B) RT-PCR analysis of ofd1 mRNA expression in WT embryos and ik mutants. gapdh served as a normalization control. Relative band intensity of ofd1 normalized with respect to gapdh intensity is graphically represented (in triplicate) using Image J software. **p < 0.01 (C) WISH for ofd1 in WT embryos and ik mutants at 1 dpf. Scale bar, 100 μm. Enlarged view of the pronephros in WT and ik mutants at 1 dpf. Scale bar, 150 μm (D) RT-PCR analysis of ofd1 mRNA expression in control MO, ik MO, and ik MO/ik mRNA co-injected embryos at 1.5 dpf. gapdh served as a normalization control. Relative band intensity of ofd1 normalized with respect to gapdh intensity is graphically represented (in three individual experiments) using Image J software. **p < 0.01, ***p < 0.001 (E) WISH for ofd1 in control MO, ik MO, and ik MO/ik mRNA co-injected embryos at 1 dpf. Enlarged view of the pronephros in control MO, ik MO, and ik MO/ik mRNA co-injected embryos at 1 dpf. Scale bar, 100 μm

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Ciliopathy phenotypes of ik mutants are rescued by co-injection of ofd1

Next, to determine whether ofd1 could rescue the ciliopathy phenotypes in ik mutants, we analyzed the phenotypes of ik morphants co-injected with ofd1 mRNA (Fig. 5A). As expected, ik morphants co-injected with ofd1 mRNA showed 73.7% rescued WT body curvature compared to that of ik morphants (4%) and normal otolith phenotype (82.6%) in contrast with that of ik morphants (54.8%). In addition, cyst formation observed in ik morphants by approximately 53% was reduced to 21.4% in ik morphants co-injected with ofd1 mRNA. These results indicate that ofd1 is a downstream target of ik. To exclude the off-target effect of co-injection of ofd1 mRNA, we examined spatial ofd1 expression in ik morphants and ofd1 mRNA co-injected ik morphants using WISH experiments (Fig. 5B). The downregulated ofd1 expression in ik morphants was enhanced by overexpressed ofd1 mRNA, indicating no off-target effect in rescue experiments. Based on the rescued phenotype of kidney cyst formation in ik morphants co-injected with ofd1 mRNA, we inferred that co-injection of ofd1 mRNA into ik morphants would also moderate the impairment of kidney development or cyst formation and affect foxj1a expression. Thus, foxj1a expression was assessed by WISH experiment and RT-PCR (Fig. 5C, D). Consequently, the significantly upregulated foxj1a expression in the pronephros of ik morphants was alleviated by co-injection of ofd1 mRNA in WISH (Fig. 5C). The mRNA level analysis of foxj1a in RT-PCR also demonstrated similar results (Fig. 5D). These results indicate that the ciliopathy phenotypes in the loss of ik result from the loss of ofd1.

Fig. 5
figure 5

The ciliopathy phenotypes of ik mutants are rescued by ofd1 co-injection (A) The lateral morphological view of control MO, ik MO, and ik MO/ofd1 mRNA co-injected embryos (body curvature and otolith imaged at 2 dpf and pronephric cyst imaged at 3 dpf). Pronephric cyst in ik MO is indicated by red arrowhead. Stacked bar graph displays the phenotypic percentage of embryos. B WISH for ofd1 in control MO, ik MO, and ik MO/ofd1 mRNA co-injected embryos at 1 dpf. Scale bar, 100 μm. Enlarged view of the pronephros in control MO, ik MO, and ik MO/ik mRNA co-injected embryos at 1 dpf. Scale bar, 150 μm. C WISH for foxj1a in control MO, ik MO, and ik MO/ofd1 mRNA co-injected embryos at 2.5 dpf. Scale bar, 100 μm (D) RT-PCR analysis of foxj1a mRNA expression in control MO, ik MO, and ik MO/ofd1 mRNA co-injected embryos at 1.5 dpf. gapdh served as a normalization control. Relative band intensity of foxj1a normalized with respect to gapdh intensity is graphically represented using Image J software (in three individual experiments). *p < 0.05, ***p < 0.001

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ik acts as the key regulator of ofd1 and regulates proper ciliogenesis in vivo

Finally, to elucidate the correlation between ik and ofd1 in ciliogenesis, we examined ciliary morphology of ik morphants co-injected with ofd1 mRNA using the Tg(brn3c:GFP) zebrafish line. As expected, ik morphants co-injected with ofd1 mRNA showed rescued cilia length and bundle numbers in hair cells compared to those of ik morphants. It also revealed comparable number of hair cells in the inner ear to that of the control morphant group (Fig. 6A). Similarly, in immunostaining for cilia in the pronephros, the randomly disorganized cilia morphology of ik morphants was predominantly restored by co-injection of ofd1 mRNA (Fig. 6B). Taken together, ik contributes to proper ciliogenesis in vivo by regulating ofd1.

Fig. 6
figure 6

ofd1 co-injection can rescue ciliary dysmorphology of ik morphants (A) Confocal microscopic images of control MO, ik MO, and ik MO/ofd1 mRNA co-injected Tg(brn3c:GFP) embryos at 2 dpf. Scale bar, 20 µm. Bar graphs show quantification of ciliary length and number in the inner ears of control MO, ik MO, and ik MO/ofd1 mRNA co-injected embryos at 2 dpf. **p < 0.01, ***p < 0.001 (B) Whole-mount immunostaining of pronephric cilia using anti-acetylated-α-tubulin (red) at 2 dpf control MO, ik MO, and ik MO/ ofd1 mRNA co-injected embryos. Scale bar, 20 µm. Stacked bar graph displays the percentage of ciliary morphology of embryos in the anterior pronephric duct (percentage of abnormal ciliary morphology of embryos: control MO, 5.12% (37/39); ik MO, 86.6% (4/30); ik MO + ofd1 mRNA, 16.6% (15/18)

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Discussion

Ciliopathies emerge from functional and structural abnormalities of cilia [41]. Over the last two decades, the functions of genes related to ciliopathies have been revealed through genetic analysis of animal models and humans [42, 43]. Although 187 ciliopathy-associated genes have been characterized until recently, the roles of 241 candidate genes for ciliary function remain unclear [11]. The function of these candidate genes remains to be elucidated to understand the pathophysiology of ciliopathies. In this study, we showed that knockdown of ik induced ciliopathy phenotypes with disrupted ciliogenesis, which was accompanied by downregulation of ciliopathy-associated gene ofd1.

In general, failure to align the LR asymmetry sides of the cardiovascular system is often proposed as the main feature of ciliopathy [44]. However, ik mutants showed normally positioned LR loops in the cardiovascular system (Fig. 1C). We suggest two possible explanations for this phenomenon. First, this can be explained by the phenotypic expression of ik during embryogenesis. As embryonic transcripts are quiescent during early embryogenesis, maternally provided proteins and RNA directly contribute to early embryonic development [45,46,47,48]. And then, when the embryo enters the MZT period, transcriptional factors produced from the activated zygotic genome are governed by gene expression programs. We observed that ik transcript was detected at the early stage, but the expression level was significantly weaker than that at the zygotic period (Fig. 1A). Therefore, considering that LR asymmetry patterning arises in early somite stages [49], ik knockdown at an early stage would not affect phenotypes characterized during early embryogenesis. Second, foxj1 is known to be required for LR body asymmetry during early embryogenesis [50]. Given the expression of foxj1a is significantly higher in ik mutants than in the WT embryos, foxj1a might perform complementary functions in the absence of ik. To understand the precise role of ik in ciliogenesis in vivo, further studies are needed to focus on the spatiotemporal effect of ik during embryogenesis.

Understanding ciliated cell types in specific tubular segments provides insights into the relationship between cilia and kidney development [37, 51]. Our study also demonstrates that ik mutants resulted in abnormal ciliary morphology in the pronephros, but interestingly, it was restricted only to proximal renal tubules and not distal tubules. Furthermore, we observed structurally abnormal development of PCT and PST segments in ik mutants (Fig. 3A). In nearly all vertebrates, two main ciliated cells exist: a single non-motile primary ciliated cell and MCCs [52]. Although ciliopathy syndromes linked to primary cilium defects have been extensively studied, studies on MCCs have been experimentally challenged because MCCs are located deep inside organs, and monitoring their behavior is difficult. However, zebrafish embryos have optical transparency; therefore, they have been used as a powerful model to study MCCs. In zebrafish, MCCs are present in several tissues, including the pronephros [53]. In the pronephros, MCCs are dispersed mainly in the PST region but are also adjacent within the PCT and DE segments. Given that disorganized ciliated cells were only observed in the PCT and PST in ik mutants, depletion of ik may specifically impair the proliferation or differentiation of MCCs in the proximal tubule. Interestingly, human clinical studies also showed similar results that patients with kidney diseases have abnormal MCCs with central microtubular pair (9 + 2 structure), whereas healthy people only contain non-motile primary cilia with 9 + 0 structure [54, 55], specifically in the proximal tubules not in the distal tubules [56]. These results suggest the possibility that abnormal differentiation of MCCs into proximal tubular cells would specifically affect kidney-related ciliopathy in vivo. Regarding the fact that ik mutants have phenotypes similar to those of human kidney patients, investigation of the role of IK using ik mutants will elucidate the molecular details of kidney ciliopathy in humans.

OFD type I syndrome is a rare ciliopathy disorder; however, more than seven million global patients have symptoms of renal cystic disease and other complicated disorders [57]. Because no effective pharmacological therapies for OFD syndrome is currently available, more biological and therapeutic approaches have been required. Here, we found that ik mutants had reduced ofd1 mRNA expression. Because IK was also known as a pre-mRNA splicing factor in vitro, we additionally demonstrated whether there exists damage in pre-mRNA splicing of ofd1 but downregulated ofd1 in ik mutants was not from the pre-mRNA splicing error (Additional file 5). Based on these results, there are two possibilities to explain the cause of downregulated ofd1 expression. First, it might be the result of a transcription factor defects of ofd1 in ik mutants. Although we have clearly demonstrated in this study that IK directly contributes to expression of OFD1 at transcription level through rescue experiments using IK mutant construct in vitro (Additional file 4: Fig. S4), the specific transcription factor of OFD1 has not yet been identified. Therefore, it is necessary to investigate the specific transcription mechanism for ofd1 gene. Second, we previously reported that knockdown of ik led genomic instability showing abnormal chromosome features in vitro [58]. Thus, loss of ik is also likely to induce genomic instability even in vivo leading to reduction of the ofd1 mRNA expression. This possibility is also further supported by recent report showing mutation of SMU1, which is a reciprocal interactor of IK and functions as a unit by stabilizing each other, results in chromosomal abnormality with aberrant sister chromatid exchanges [59]. They referred that defects of these splicing factors could disrupt links between RNA metabolism and genomic recombination, thereby leading to abnormal gene expression. Therefore, it is worthwhile to explore the association between loss of ik and chromosomal instability to elucidate the regulatory mechanism of ofd1. In the underlying molecular mechanism mediated by the ofd1, ofd1 inhibits autophagy by promoting the degradation of the unc-51-like kinase ulk1 complex component atg13 [60, 61]. Thus, the stabilized ulk1 complex by the loss of ofd1 enhances autophagy and causes OFD type I syndrome. As the ofd1 expression was downregulated in loss of ik leading to ciliopathy phenotypes, further experiments are required to address whether loss of ofd1 enhances autophagy in ik mutants. Considering that IK is well known to play a role in regulating mitotic cell cycle progression with mitotic kinase [16,17,18] and ciliogenesis is dependent on cell cycle progression under the control of autophagy, understanding the ciliogenesis–cell cycle–autophagy crosstalk between IK and OFD1 will provide another molecular aspect for understanding human ciliopathy.

Conclusions

In conclusion, this study is the first to demonstrate the role of ik in ciliogenesis in vivo. In the absence of ik, zebrafish embryos display various ciliopathy phenotypes with abnormal ciliary morphology in ciliary tissues. We also address that ofd1, an essential gene for ciliogenesis, is a downstream target of ik. Elucidating the exact role of ik provides valuable insights into the understanding of ciliogenesis and ciliopathy in vivo.

Materials and methods

Zebrafish maintenance and husbandry

WT AB*, Tg(brn3c:GFP), and Tg(bactin2:Arl13b-GFP) zebrafish (Danio rerio) were used in this study. ik mutant zebrafish embryos have been previously described [20]. Embryos/larvae were raised at 28.5 °C and were natural spawning of adult zebrafish. Zebrafish were maintained in a tank rack system with automatic water recycling, water changes, and monitoring and adjustment of water parameters. Fish were kept under a 14 h light and 10 h dark cycles and were fed artemia (INVE aquaculture, Belgium) and pelleted dry food (Tetra, Blacksburg, VA, USA) thrice a day. Embryos were anesthetized with 0.2 mg/ml tricaine (3-aminobenzoic acid ethylester; Sigma-Aldrich, St. Louis, Missouri, USA) and embedded in methylcellulose. All zebrafish husbandry and experimental protocols complied with institutional guidelines and were approved by the local ethics board (Sookmyung Women’s University Animal Care and Use Committee, SMWU-IACUC-1712-036).

Cell culture and transfection

The human retinal pigment epithelial cell line, hTERT-RPE1 were maintained in DMEM/F12 media supplemented with 10% FBS (Gibco, USA) at a humidified 37 °C, 5% CO2. IK siRNA transfection into RPE1 cells was performed using Lipofectamine RNAiMax Transfection Reagent (Invitrogen, USA) for 48 h according to the manufacturer's protocol with the siRNA oligonucleotides (5′- CUACCAAGGAGUUGAUCAA-3′) [58]. To knockdown of AURKA and AURKB, the siRNA of ON-TARGET plus SMART pool (Dharmacon) were used and transfection was performed according to the manufacturer's protocol. For rescue experiment, wildtype-full length human IK or mutant-IK T485A cloned into pcDNA 3.1 vector (Invitrogen, USA) was re-transfected using X-tremeGENE HP DNA Transfection Reagent (Roche, Germany) into siRNA transfected cells for 24 h according to the manufacturer's protocol.

RT-PCR

Total RNA was extracted using TRIzol (Ambion, Foster City, California, USA), according to the manufacturer’s instructions. RNA concentration was measured using a NanoDrop (Thermo Fisher Scientific, Waltham, Massachusetts, USA), and 2 μg of RNA sample was reverse-transcribed into cDNA using a GoScript reverse transcription system kit (Promega, Madison, Wisconsin, USA).

Primer sequences for RT-PCR were designed using the National Center for Biotechnology Information (NCBI) primer-blast program (https://www.ncbi.nlm.nih.gov/tools/primer-blast/) (Additional file 7: Table S1).

WISH

WISH was performed with standard protocols as previously described [62, 63]. Zebrafish embryos were fixed with 4% paraformaldehyde in phosphate-buffered saline (PBS) at 4 °C overnight and then rinsed with PBST (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 2 mM KH2PO4, 0.1% Tween 20) for rehydration. To increase the permeability of RNA probes, embryos were incubated with proteinase K (10 μg/ml), depending on their developmental stages. Embryos were refixed in 4% paraformaldehyde for 20 min at 25 °C and prehybridized in hybridization solution (HYB solution) containing 50% formamide, 5 × SSC (SSC: 3 M NaCl and 0.3 M sodium citrate, pH 7), 50 μg/ml heparin, 500 μg/ml RNA, 46 mM citric acid, pH 6, and 0.1% Tween 20 for at least 1 h at 65 °C. The HYB solution was removed, and the RNA probe was added to embryos at 65 °C and incubated overnight. The embryos were washed at 65 °C for 30 min once with 50% formamide in 2 × , 1 × , and 0.2 × SSC. For staining, embryos were incubated with blocking solution [2% sheep serum, 2 mg/ml bovine serum albumin (BSA) in PBST] for 2 h and then with anti-digoxigenin Fab fragment conjugated with alkaline phosphatase (Roche, Penzberg, Germany) at 1:5000 dilution at 4 °C overnight. Embryos were washed with PBST and visualized using BM Purple (Roche) as a substrate for alkaline phosphatase. After sufficient staining intensity was achieved, PBST was added to stop the reaction. Images were captured using a Leica DVM6 digital microscope (Leica, Germany) at Korea Basic Science Institute. Primers used for the RNA probe are provided in the Additional file 8: Table. S2, and foxj1a primers were designed as previously described [64].

Immunofluorescence

For immunostaining of cilia in zebrafish embryos, embryos were fixed in 4% paraformaldehyde at 4 °C overnight and blocked with blocking solution (1 × PBS, 3% BSA, and 1% Triton X-100) at RT for 2 h. Zebrafish embryos were incubated with anti-acetylated-α-tubulin (Proteintech, CL594-66200, 1:500 dilution) at 4 °C overnight. Next, embryos were washed PBST (with 1% Triton X-100) for 30 min and incubated with Alexa Fluor 488-conjugated secondary antibodies (Sigma, 1:500 dilution) at 4 °C overnight. The nuclei of the fixed embryos were stained with 4', 6'-diamidino-2-phenylidole (DAPI). Images were acquired using a confocal microscope (LSM700, Zeiss, Oberkochen, Germany) at the Chronic and Metabolic Diseases Research Center of Sookmyung Women’s University.

Immunoblotting

The Immunoblotting was performed as described previously [20]. The primary antibodies used for immunoblotting were as follows: rabbit polyclonal anti-OFD1 (Proteintech, 22851–1-AP), mouse monoclonal anti-β-actin (Santa Cruz, sc-47778), rabbit monoclonal p-AURKA Thr288 (Cell Signaling, #3079) and rabbit monoclonal p-AURKB Thr232 (Cell Signaling, #2914).

TEM

Embryos were whole-mount fixed with Karnovsky’s fixative solution containing 2% glutaraldehyde and 2% paraformaldehyde overnight at 4 °C. After washing thrice with 0.05 M sodium cacodylate, they were postfixed with 1% aqueous osmium tetroxide. Then, embryos were stained en bloc using 0.5% aqueous uranyl acetate and were dehydrated through a graded series (30–100%) of ethanol, 20 min for each step. Embryos were incubated with progressively concentrated propylene oxide dissolved in ethanol and infiltrated with increasing concentrations of Spurr’s resin. The samples were embedded in Spurr’s resin, baked in an oven at 65 °C overnight, and sectioned using an ultramicrotome (EM UC7, Leica, Wetzlar, Germany). The sections were observed using a TEM (Talos L120C, FEI, Czech Republic) at the NICEM transmission electron microscope laboratory (Seoul National University, Seoul, Republic of Korea).

Microinjection of morpholino oligonucleotide (MO) and mRNA

The MO sequence of ik was 5′-GGAGCCAGAGGATTAGAGTACACAT-3′, as previously described [20] and was obtained from GeneTools (Corvallis, OR, USA). For knockdown of ik, embryos at one-cell stage were injected with 4 ng of MO using a PV820 Pneumatic PicoPump (World Precision Instruments, Sarasota, Florida, USA) and reared in E3 medium (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4, and 0.1% methylene blue) at 28.5 °C. Images were captured using a microscope (Olympus, Biotek Cytation 5) and a confocal microscope (LSM700, Zeiss, Oberkochen, Germany). For rescue experiments, full-length ik or ofd1 mRNA was transcribed in vitro using a T7 mMESSAGE mMACHINE kit (Ambion, Foster City, California, USA) according to the manufacturer’s instructions. Embryos were then co-injected with 250 pg ik or ofd1 mRNA with 4 ng of ik MO per egg.

RNA-seq analysis

We used a previously analyzed RNA-seq dataset of ik mutants to evaluate differentially expressed genes (DEGs) [20]. In brief, genome-wide RNA-seq data of zebrafish embryos at 3 dpf were analyzed by eBiogen (Seoul, Korea). RNA with an absorbance ratio > 1.8 and integrity > 7.0 was confirmed. Fragments per kilobase of transcript per million mapped reads (FPKM) were used to determine gene expression. Upregulated or downregulated genes were identified using the ExDEGA program (EBIOGEN Inc., Seoul, Republic of Korea). Gene Ontology (GO) by the DAVID analysis was performed through Quick GO (https://www.ebi.ac.uk/QuickGO). Heatmap profiles of DEGs were prepared according to GO analysis using the Multiple Experiment Viewer software program v.4.9. A gene set representing > 1.5-fold change in ik KO embryos is presented by red color, and blue color represents < 1.5-fold change.

Statistical analysis

The values in the graphs are presented as mean ± standard deviation (SD). Multiple comparisons within groups were performed using one-way analysis of variance (ANOVA), and differences between the means of individual groups were evaluated using Student’s t-tests. A p-value < 0.05 was considered as the threshold for significant differences (*p < 0.05, **p < 0.01, ***p < 0.001) and is provided in the figure legends.

Availability of data and materials

All datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request.

Abbreviations

MOs:

Morpholino antisense oligonucleotides

MZT:

Maternal to zygotic transition

OFD1:

Oral-facial-digital 1

PCR:

Proximal convoluted tubule

TEM:

Transmission electron microscope

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Acknowledgements

Zebrafish in Figs 2A, C, 3A were illustrated by Biorender.

Funding

This study was supported by the Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Education: NRF-2021R1I1A1A01060442 (to H.I.K), NRF-2022R1F1A1074668 (to M.J.K), NRF-2021R1A2C3003414 (to Y.Y), NRF-2021R1A6A1A03038890 (to Y.Y), and the Korea Basic Science Institute (National Research Facilities and Equipment Center); No. 2021R1A6C101A564 (to Y.Y).

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Authors and Affiliations

  1. Research Institute of Women’s Health, Sookmyung Women’s University, Seoul, 04312, South Korea

    Hye In Ka, Mina Cho, Se Hwan Mun, Sora Han, Min Jung Kim & Young Yang

  2. Chronic and Metabolic Diseases Research Center, Sookmyung Women’s University, Seoul, 04312, South Korea

    Hye In Ka, Se Hwan Mun & Young Yang

  3. Seoul Center, Korea Basic Science Institute, Seoul, 02841, South Korea

    Seung-Hae Kwon

  4. Department of Biological Sciences, Sookmyung Women’s University, Seoul, 04312, South Korea

    Min Jung Kim & Young Yang

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  2. Mina Cho
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  3. Seung-Hae Kwon
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  4. Se Hwan Mun
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Contributions

HIK planned the research, performed the experiments and wrote the manuscript. MC performed the experiments and wrote the manuscript. SHK, SHM and SH prepared key materials and discussed the results. MJK and YY supervised the entire project, co‐wrote, and proofread the manuscript. All authors read and approved the final manuscript.

Corresponding authors

Correspondence to Min Jung Kim or Young Yang.

Ethics declarations

Ethics approval and consent to participate

All zebrafish husbandry and experimental protocols complied with institutional guidelines and were approved by the local ethics board (Sookmyung Women’s University Animal Care and Use Committee, SMWU-IACUC-1712-036).

Consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

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Supplementary Information

Additional file 1:

Figure S1. The relative ik mRNA expression at different developmental stages. The ik mRNA expression at different developmental stages was quantified uisng qRT-PCR and normalized to β-actin. *p < 0.05, **p < 0.01, ***p < 0.001.

Additional file 2:

Figure S2. Sections of whole-mount ISH probed with ik. (A) Dorsal (left) and lateral (right) view of 2-dpf embryo stained with the ik probe with planes of sections in follow panels as indicated (B) Section image of part of head with otic vesicle (arrow) of zebrafish embryo (C) Transverse cross section of the pronephric tubules (arrow).

Additional file 3:

Figure S3. Large field-of-view images of cilia in the pronephros of ik mutants imaged by TEM. (A) Cross-sectional image of zebrafish pronephros. Motile cilia were observed in the sample (arrow). Scale bar, 200 nm (B) High magnification image of (A). Scale bar, 100 nm.

Additional file 4:

Figure S4. The OFD1 mRNA expression in re-transfection of wildtype-IK or mutant-IK in RPE cells. The OFD1 mRNA expression in IK knock-downed human RPE cells which were re-transfected with cDNA encoding IK wild-type or T485A mutant.

Additional file 5:

Sequencing results of ofd1 in ik KO embryos at 2.5 dpf.

Additional file 6:

Figure S5. The expression analysis of aurora kinases. (A) The aurora kinase A (aurka) and B (aurkb) mRNA expression in WT embryos and ik mutants at 2 dpf (B) The OFD1 expression at the protein level in human RPE cells transfected with siAURKA or siAURKB for 48 h.

Additional file 7:

Table S1. List of primer sequences used for RT-PCR analysis.

Additional file 8:

Table S2. List of primer sequences used for WISH analysis.

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Ka, H.I., Cho, M., Kwon, SH. et al. IK is essentially involved in ciliogenesis as an upstream regulator of oral-facial-digital syndrome ciliopathy gene, ofd1. Cell Biosci 13, 195 (2023). https://doi.org/10.1186/s13578-023-01146-9

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Keywords

Cell & Bioscience

ISSN: 2045-3701

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