September 2018
Volume 59, Issue 11
Open Access
Genetics  |   September 2018
IFT52 as a Novel Candidate for Ciliopathies Involving Retinal Degeneration
Author Affiliations & Notes
  • Xue Chen
    Department of Ophthalmology, The First Affiliated Hospital of Nanjing Medical University, State Key Laboratory of Reproductive Medicine, Nanjing, China
    Department of Ophthalmology and Vision Science, Eye and ENT Hospital, Shanghai Medical College, Fudan University, Shanghai, China
    Key Laboratory of Myopia of State Health Ministry (Fudan University) and Shanghai Key Laboratory of Visual Impairment and Restoration, Shanghai, China
  • Xiaoguang Wang
    Department of Ophthalmology, Ningxia Eye Hospital, People's Hospital of Ningxia Hui Autonomous Region (First Affiliated Hospital of Northwest University for Nationalities), Yinchuan, China
  • Chao Jiang
    Department of Ophthalmology, The First Affiliated Hospital of Nanjing Medical University, State Key Laboratory of Reproductive Medicine, Nanjing, China
  • Min Xu
    Department of Ophthalmology, Northern Jiangsu People's Hospital, Yangzhou, China
  • Yang Liu
    Department of Ophthalmology, Ningxia Eye Hospital, People's Hospital of Ningxia Hui Autonomous Region (First Affiliated Hospital of Northwest University for Nationalities), Yinchuan, China
  • Rui Qi
    Medical College of Northwest Minzu University, Lanzhou, China
  • Xiaolong Qi
    Department of Ophthalmology, Ningxia Eye Hospital, People's Hospital of Ningxia Hui Autonomous Region (First Affiliated Hospital of Northwest University for Nationalities), Yinchuan, China
  • Xiantao Sun
    Department of Ophthalmology, Children's Hospital of Zhengzhou, Zhengzhou, China
  • Ping Xie
    Department of Ophthalmology, The First Affiliated Hospital of Nanjing Medical University, State Key Laboratory of Reproductive Medicine, Nanjing, China
  • Qinghuai Liu
    Department of Ophthalmology, The First Affiliated Hospital of Nanjing Medical University, State Key Laboratory of Reproductive Medicine, Nanjing, China
  • Biao Yan
    Department of Ophthalmology and Vision Science, Eye and ENT Hospital, Shanghai Medical College, Fudan University, Shanghai, China
    Key Laboratory of Myopia of State Health Ministry (Fudan University) and Shanghai Key Laboratory of Visual Impairment and Restoration, Shanghai, China
  • Xunlun Sheng
    Department of Ophthalmology, Ningxia Eye Hospital, People's Hospital of Ningxia Hui Autonomous Region (First Affiliated Hospital of Northwest University for Nationalities), Yinchuan, China
  • Chen Zhao
    Department of Ophthalmology, The First Affiliated Hospital of Nanjing Medical University, State Key Laboratory of Reproductive Medicine, Nanjing, China
    Department of Ophthalmology and Vision Science, Eye and ENT Hospital, Shanghai Medical College, Fudan University, Shanghai, China
    Key Laboratory of Myopia of State Health Ministry (Fudan University) and Shanghai Key Laboratory of Visual Impairment and Restoration, Shanghai, China
    Department of Ophthalmology, Children's Hospital of Zhengzhou, Zhengzhou, China
  • Correspondence: Chen Zhao, Department of Ophthalmology and Vision Science, Eye and ENT Hospital, Shanghai Medical College, Fudan University, 138 Yi xue yuan Road, Xuhui Qu, Shanghai 200032, China; dr_zhaochen@163.com
  • Xunlun Sheng, Department of Ophthalmology, Ningxia Eye Hospital, People's Hospital of Ningxia Hui Autonomous Region, West Huaiyuan Road, Yinchuan, China; shengxunlun@163.com
  • Biao Yan, Department of Ophthalmology and Vision Science, Eye & ENT Hospital, Shanghai Medical College, Fudan University, 138 Yi xue yuan Road, Xuhui Qu, Shanghai 200032, China; yanbiao1982@hotmail.com
  • Footnotes
     XC, XW, and CJ contributed equally to the work presented here and should therefore be regarded as equivalent authors.
Investigative Ophthalmology & Visual Science September 2018, Vol.59, 4581-4589. doi:10.1167/iovs.17-23351
  • Views
  • PDF
  • Share
  • Tools
    • Alerts
      ×
      This feature is available to authenticated users only.
      Sign In or Create an Account ×
    • Get Citation

      Xue Chen, Xiaoguang Wang, Chao Jiang, Min Xu, Yang Liu, Rui Qi, Xiaolong Qi, Xiantao Sun, Ping Xie, Qinghuai Liu, Biao Yan, Xunlun Sheng, Chen Zhao; IFT52 as a Novel Candidate for Ciliopathies Involving Retinal Degeneration. Invest. Ophthalmol. Vis. Sci. 2018;59(11):4581-4589. doi: 10.1167/iovs.17-23351.

      Download citation file:


      © ARVO (1962-2015); The Authors (2016-present)

      ×
  • Supplements
Abstract

Purpose: Mutations in the intraflagellar transport protein 52 homolog (IFT52) gene are reported to interrupt ciliary function and cause short-rib thoracic dysplasia (SRTD), a specific form of skeletal ciliopathy. However, the roles of these mutations in retinal ciliopathy are inexplicit. We herein aim to study the impact of IFT52 mutations in retinopathies.

Methods: A patient with syndromic ciliopathy, presenting mild SRTD (skeletal ciliopathy) and Liber congenital amaurosis (LCA; retinal ciliopathy), and nine unaffected family members were recruited. Comprehensive systemic evaluations, including ophthalmic tests, were received by the patient. Whole genome sequencing (WGS) was applied for genetic annotation. An in vitro cell system was employed to study the pathogenicity of the variant.

Results: WGS identified a homozygous missense variation in IFT52, c.556A>G (p.T186A), carried by the patient but absent in both unaffected siblings. In silico analysis supported the pathogenic nature of this highly conserved variant. Structural analysis suggested that this substitution could generate a novel hydrogen bond between the mutated residue 186 and proline at residue 192, thus potentially interrupting the tertiary structure and the stability of the IFT52 protein. In vitro cellular study indicated that this mutation might disturb the stability of encoded IFT52 protein and dramatically disrupt cilia elongation in hTERT-RPE1 cells in a loss-of-function manner.

Conclusions: This report expands ocular phenotypes of IFT52 mutation-caused ciliopathy to include retinal ciliopathy and demonstrates its deleterious nature in interrupting primary ciliary function. Our study hence highlights the need for screening for IFT52 mutations in LCA patients and ophthalmic reviews of patients carrying IFT52 mutations.

Ciliopathies refer to a group of multisystemic genetic disorders triggered by mutations in genes required for ciliary assembly and maintenance.1 Intraflagellar transport (IFT) is a highly conserved biological process by which various proteins are transported along the ciliary axoneme from cytoplasm to ciliary tip.2 Specifically, mutations in genes encoding IFT-B complex components could disrupt ciliary assembly and generate a wide spectrum of ciliopathies.35 The IFT protein 52 homolog (IFT52; MIM: 617094) gene, located on 20q13.12, encodes the proline-rich Ift52 protein, a particle of the IFT-B core complex. IFT52 mutations have been previously recognized to cause autosomal recessive short-rib thoracic dysplasia (SRTD) 16 with or without polydactyly (SRTD16, MIM: 617102), a distinct type of skeletal ciliopathies.3,4 SRTD16 presents various forms and is characterized by a constricted thoracic cage, short ribs, shortened tubular bones, and a trident appearance of the acetabular roof.6,7 Polydactyly is variably present. The present report of the patient further expands the IFT52 mutation relevant phenotypes to include Liber congenital amaurosis (LCA; MIM: 204000) with a milder systemic defect than the two previously reported cases. 
Materials and Methods
Patients
Our study, which conformed to the tenets of the Declaration of Helsinki, was approved and prospectively reviewed by ethics committee of the People's Hospital of Ningxia Hui Autonomous Region. Written informed consent was obtained from all participants or their legal guardians before enrollment. One patient and eight unaffected family members were recruited from family YWH of Hui ethnicity (see Fig. 2A). First cousin consanguineous marriage was reported. Medical history was obtained from each participant. The proband received systemic clinical evaluations, including comprehensive ophthalmic examinations. Peripheral blood samples were collected from all participants using 5-mL tubes with EDTA. Genomic DNA was extracted using a DNA blood kit (QIAmp; Qiagen, Valencia, CA, USA). Another 100 healthy controls from the same ethnic group free of ocular problems and other major systemic diseases were also included for DNA extraction. 
Figure 1
 
Systemic evaluations of patient YWH-IV:1. (A, B) Fundus photos of patient YWH-IV:1 suggest waxy optic disk and attenuated retinal vessels. Sporadic pigment deposits are also noticed in the midperipheral retina (indicated by arrow). (C, D) OCT examinations of patient YWH-IV:1 reveal loss of ellipsoid in bilateral macular region. (E) Both scotopic and photopic ERG responses are diminished. (F) Patient YWH-IV:1 presents small chest with short ribs and micromelic limbs. (G) Sandal gap is found in her right foot, between her fourth and fifth phalanges. (H, I) Dental dysplasia is identified. Panoramic dental X-ray suggests that one left mandibular premolar, one right mandibular premolar, and one right mandibular incisor are missing (I).
Figure 1
 
Systemic evaluations of patient YWH-IV:1. (A, B) Fundus photos of patient YWH-IV:1 suggest waxy optic disk and attenuated retinal vessels. Sporadic pigment deposits are also noticed in the midperipheral retina (indicated by arrow). (C, D) OCT examinations of patient YWH-IV:1 reveal loss of ellipsoid in bilateral macular region. (E) Both scotopic and photopic ERG responses are diminished. (F) Patient YWH-IV:1 presents small chest with short ribs and micromelic limbs. (G) Sandal gap is found in her right foot, between her fourth and fifth phalanges. (H, I) Dental dysplasia is identified. Panoramic dental X-ray suggests that one left mandibular premolar, one right mandibular premolar, and one right mandibular incisor are missing (I).
Figure 2
 
Family pedigree and genetic annotation of the identified mutation. (A) Pedigree of family YWH. +: wild-type allele; MU: IFT52 c.556A>G (p.T186A). (B) Sequence chromatograms of the identified mutation. (C) Orthologous protein sequence alignment of IFT52 from humans (H. sapiens), chimpanzees (P. troglodytes), dogs (C. lupus), cows (B. taurus), pigs (S. scrofa), mice (M. musculus), chickens (G. gallus), zebrafish (D. rerio), and fruit flies (D. melanogaster). Conserved residues are shaded. The mutated residue 186 is boxed and indicated. (D) Schematic representation of the relative linear location of the identified IFT52 mutation in context of genome structure (upper) and protein structure (below). Regions of IFT52 protein that interact directly with another three IFT-B components (IFT88, IFT70, and IFT46) are indicated. (E) Crystal structural analysis of the wild-type and mutant IFT52 protein. A novel hydrogen bond between residues 186 and 192 is generated by the mutation. (F) Expression of Ift52 in multiple murine tissues including heart, liver, spleen, lung, kidney, brain, muscle, stomach, intestines, colorectum, neural retina, RPE, and optic nerve is demonstrated. A 243-bp PCR product of the murine Ift52 (top panel) is detected in all tested tissues. PCR product of the murine Rplp0 (109 bp) is analyzed in parallel as a loading control.
Figure 2
 
Family pedigree and genetic annotation of the identified mutation. (A) Pedigree of family YWH. +: wild-type allele; MU: IFT52 c.556A>G (p.T186A). (B) Sequence chromatograms of the identified mutation. (C) Orthologous protein sequence alignment of IFT52 from humans (H. sapiens), chimpanzees (P. troglodytes), dogs (C. lupus), cows (B. taurus), pigs (S. scrofa), mice (M. musculus), chickens (G. gallus), zebrafish (D. rerio), and fruit flies (D. melanogaster). Conserved residues are shaded. The mutated residue 186 is boxed and indicated. (D) Schematic representation of the relative linear location of the identified IFT52 mutation in context of genome structure (upper) and protein structure (below). Regions of IFT52 protein that interact directly with another three IFT-B components (IFT88, IFT70, and IFT46) are indicated. (E) Crystal structural analysis of the wild-type and mutant IFT52 protein. A novel hydrogen bond between residues 186 and 192 is generated by the mutation. (F) Expression of Ift52 in multiple murine tissues including heart, liver, spleen, lung, kidney, brain, muscle, stomach, intestines, colorectum, neural retina, RPE, and optic nerve is demonstrated. A 243-bp PCR product of the murine Ift52 (top panel) is detected in all tested tissues. PCR product of the murine Rplp0 (109 bp) is analyzed in parallel as a loading control.
Animals
C57BL/6 mice housed in the Model Animal Research Center, Nanjing University, conformed to Institutional Animal Care and Use Committee–approved protocol. All animal experiments conformed to the ARVO Animal Statement for the Use of Animal in Ophthalmic and Vision Research and were approved by the local ethics committee. 
Whole Genome Sequencing (WGS) and Bioinformatics Analysis
WGS was performed on patient YWH-IV:1 to reveal her disease-causative mutations per a previously described protocol.8 Briefly, DNA was first prepared with a sequencing kit (TruSeq Nano DNA HT Library Prep Kit; Illumina, San Diego, CA, USA). Sequencing was conducted on the HiSeq X10 platform (Illumina). Obtained reads were then aligned to the human hg19 genome using Burrows-Wheeler Aligner v.0.6.1.9 Genome Analysis Tool Kit v.3.6 (Intel Corp, Santa Clara, CA, USA) was next applied for base quality recalibration and local realignment. Atlas-SNP2 and Atlas-Indel2 were further used for variant calling.10 All revealed variants were filtered against seven single-nucleotide polymorphism (SNP) databases, including dbSNP144 (http://hgdownload.cse.ucsc.edu/goldenPath/hg19/database/snp144.txt.gz; available in the public domain from University of California Santa Cruz, Santa Cruz, CA, USA), the HapMap project (ftp://ftp.ncbi.nlm.nih.gov/hapmap; available in the public domain from the National Center for Biotechnology Information, Bethesda, MD, USA), the 1000 Genomes Project (ftp://ftp.1000genomes.ebi.ac.uk/vol1/ftp; available in the public domain from The European Bioinformatics Institute, Cambridgeshire, UK), the YanHuang database (http://yh.genomics.org.cn/; available in the public domain from Bejing Genomics Institute at Shenzhen, Shenzhen, China), the Exome Variant Server (http://evs.gs.washington.edu/EVS/; available in the public domain from NHLBI Exome Sequencing Project), the Exome Aggregation Consortium database (http://exac.broadinstitute.org/; available in the public domain from the Exome Aggregation Consortium), and the Genome Aggregation database (http://gnomad.broadinstitute.org/; available in the public domain from the Genome Aggregation database. A full list of contributing groups can be found at http://gnomad.broadinstitute.org/about.). Variants found homozygous in the seven SNP databases or with a minor allele frequency of over 0.01 were discarded. Noncoding variants were discarded. Only variants located within an annotated exon or within 10 bp (base pairs) on either side were included for further analysis. 
Sanger Sequencing, RT-PCR, and Real-Time PCR
Sanger sequencing was performed for intrafamilial cosegregation analysis and prevalence test in 100 unrelated controls using a previously defined protocol.11 Primer information was provided upon request. RNA isolation, cDNA synthesis, PCR and real-time PCR were conducted using a previously defined protocol.11,12 Extracted RNA (1 μg) was used for cDNA synthesis using a kit (PrimeScript RT; Takara, Otsu, Shiga, Japan). RT-PCR was used to evaluate the expression pattern of Ift52 in different tissues of the C57BL/6 mice, including heart, liver, spleen, lung, kidney, brain, muscle, stomach, intestines, colorectum, neural retina, retinal pigment epithelium (RPE), and optic nerve.13 A 243-bp product of murine Ift52 (NM_172150.4) and a 109-bp product of murine housekeeping gene Rplp0 (NM_007475.5) were generated with 100 ng synthetic cDNA. For cellular study, cells were harvested at 48 hours posttransfection for RNA extraction. Real-time PCR was conducted using a dye (FastStart Universal SYBR Green Master; Roche, Basel, Switzerland) with a real-time PCR system (StepOne Plus Real-time PCR System; Applied Biosystems, Darmstadt, Germany) to detect the intracellular expression of the IFT52 gene. 
In Silico Analyses and Structural Modeling
To test the evolutionary conservation of the mutated spot, software (Vector NTI AdvanceTM 2011; Thermo Fisher Scientific, Waltham, MA, USA) was applied to align the protein sequence of IFT52 in the following orthologues: Homo sapiens (ENSP00000362121.3), Pan troglodytes (ENSPTRP00000023200.3), Canus lupus (ENSCAFP00000013725.3), Bos taurus (ENSBTAP00000026122.5), Sus scrofa (ENSSSCP00000007845.2), Mus musculus (ENSMUSP00000018002.6), Gallus (ENSGALP00000005566.3), Danio rerio (ENSDARP00000066190.4), Drosophila melanogaster (FBpp0078962.5), and Caenorhabditis elegans (R31.3.1). Crystal structures of the wild-type and mutant IFT52 protein were obtained with SWISS-MODEL online server (http://www.swissmodel.expasy.org/; provided in the public domain by the University of Basel, Basel, Switzerland). Predicted structures were displayed using open source software (PyMol v. 1.5; Schrödinger, New York, NY, USA). Sorting Intolerant From Tolerant Human Protein DB (http://sift.bii.a-star.edu.sg/; available in the public domain from The European Bioinformatics Institute) and Polymorphism Phenotyping v. 2 (http://genetics.bwh.harvard.edu/pph2/index.shtml; available in the public domain) online software was used to determine the pathogenic impacts of the identified mutation. 
Reagents and Plasmids Construction
Scramble and IFT52 small interfering RNAs (siRNAs) were purchased from RiboBio Co. Ltd (Guangzhou, China). Open reading frame sequence of human IFT52 (NM_016004) was synthesized, amplified, and inserted into the expression vector pCMV-C-Flag (Beyotime, Shanghai, China) with a FLAG tag, using EcoRI and XbaI restriction sites to generate the Ac-IFT52WT plasmids for cell transfection.13,14 A lightning site-directed mutagenesis kit (QuikChange; Agilent Technologies, Santa Clara, CA, USA) was applied to introduce the missense mutation IFT52 p.T186A into the generated Ac-IFT52WT plasmid to get the recombinant plasmid Ac-IFT52T186A. We used Sanger sequencing to validate sequences of all produced plasmids in both directions. 
Cell Transfection
Human retinal pigment epithelial (hTERT-RPE1) cells were cultured in Dulbecco's modified Eagle's medium/Ham's F12 nutrient medium (1:1, DME/F12 medium; Invitrogen, Carlsbad, CA, USA) supplemented with 10% fetal bovine serum (Invitrogen), penicillin (100 U/mL), and streptomycin (100 g/mL) at 37°C, 5% CO2. For transfection assay, cells were seeded into six-well plates and transfected with 4 μg distinct plasmid and/or 100 pmol distinct siRNA using a transfection reagent (Lipofectamine 2000; Invitrogen) per the manufacturer's protocol. 
Immunoblotting
Immunoblotting was performed using a previously defined protocol.15 For protein extraction, cells were collected at 72 hours posttransfection in ice-cold RIPA buffer (Beyotime) with protease inhibitor cocktail (Roche, Basel, Switzerland). Proteins were then separated with 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis and transferred to a polyvinylidene fluoride membrane (Millipore, Billerica, MA, USA). The membrane was blocked, incubated with primary antibodies (Supplementary Table S1) at 4°C overnight, washed, and probed with horseradish peroxidase–conjugated secondary antibodies (Supplementary Table S1) at 37°C for 1 hour. Blots were then developed using the autoradiography with the enhanced chemiluminescence-Western blotting system (BioRad, Hercules, CA, USA) per manufacturers' protocols. Proteins were quantified with ImageJ software (http://rsb.info.nih.gov/ij/index.html; provided in the public domain by the National Institutes of Health, Bethesda, MD, USA). 
Immunofluorescent Staining
For immunofluorescent staining, hTERT-RPE1 cells were collected 48 hours posttransfection. Harvested hTERT-RPE1 cells were fixed with 4% paraformaldehyde, permeabilized with 0.5% Triton X-100, and blocked with milk. For coimmunostaining, cells were then incubated with a mixture of two primary antibodies (Supplementary Table S1) with different hosts (rabbit and mouse) at 4°C overnight. Cells were then washed, probed with fluorescence-conjugated secondary antibody (Alexa Fluor 594 Donkey Anti-Rabbit/Mouse IgG [H+L]; Invitrogen) (Supplementary Table S1) at 37°C for 1 hour, washed, and then probed (Alexa Fluor 488 Goat anti-Mouse/Rabbit IgG [H+L]; Invitrogen) (Supplementary Table S1) at 37°C for 1 hour. Cell nuclei were counterstained with 4′,6-diamidino-2-phenylindole (Sigma-Aldrich Corp., St. Louis, MO, USA). Images were collected with a confocal microscope (LSM 510; Carl Zeiss, Jena, Germany). 
Cilia Abundance and Ciliary Length Measurements
Cilia abundance and ciliary length were measured according to a previously described protocol.3 hTERT-RPE1 cells were serum starved and collected 48 hours posttransfection. Ciliary axonemes and basal bodies were labeled with antibodies to acetylated-α-tubulin (Ac-α-tubulin) and ADP-ribosylation factor-like protein 13B (ARL13B), respectively. Percentages of ciliated cells were calculated from 226 cells transfected with scramble siRNA and 166 cells transfected with IFT52 siRNA in five randomly selected regions from biological triplicates. Cilia lengths were measured for 48 untransfected cells, 28 cells transfected with scramble siRNA, 54 cells transfected with IFT52 siRNA, 51 cells transfected with IFT52 siRNA and Ac-IFT52WT, and 78 cells transfected with IFT52 siRNA and Ac-IFT52T186A. All cells were randomly selected from biological triplicates. ImageJ software was used to determine ciliary length. 
Statistics
We used software (GraphPad Prism v. 4.0; GraphPad Software, San Diego, CA, USA) for statistical analysis. Student's t-test was applied for comparisons between two groups, and 1-way ANOVA was used for comparing means of three or more samples. Data were presented as mean ± SEM, and P < 0.05 was taken as statistically significant. Experiments were conducted in both biological and technical triplicates with data averaged. 
Results
Clinical Presentations
The proband was referred to our ophthalmic clinic at age 5 to evaluate her nystagmus and severe visual impairment since infancy. The parents also noticed that she preferred to press and rub her eyes. No photophobia or night blindness was reported. No family history of similar vision loss was recorded. Her best corrected vision at that time was 0.1 logMAR (20/200 Snellen equivalent) OU. Funduscopy only identified attenuated retinal vessels. ERG was not attainable due to her noncooperation. At her most recent visit to our hospital at age 11, her best corrected vision was by hand motion for both eyes. Pigment deposits, waxy optic disk, and attenuated retinal vasculature were detected in her bilateral fundus (Figs. 1A, 1B). Optical coherence tomography (OCT) demonstrated loss of ellipsoid in bilateral macula (Figs. 1C, 1D), and both scotopic and photopic ERG responses were diminished (Fig. 1E). Based on her ophthalmic presentations, including severe visual impairments since infancy, oculodigital sign, and diminished ERG responses, we concluded her ophthalmic diagnosis as LCA. 
Other than ocular phenotypes, this patient presented severe growth retardation and mild mental retardation. She had narrow chest with short ribs and micromelic limbs (Fig. 1F). At age 11, her height was 116 cm (<5th percentile) and her weight was 20 kg (<5th percentile). Her body mass index was 14.86 (between 5th and 10th percentile).16 In addition, this patient had sandal gap in her right foot (Fig. 1G). Dental dysplasia was also noticed. One of her left mandibular premolars, one right mandibular premolar, and one right mandibular incisor were missing (Figs. 1H, 1I). Other results of systemic examinations, including audiometry, magnetic resonance imaging of the brain, and laboratory tests for hepatic and renal functions, were unremarkable. No visceral malformation was revealed. 
Genetic Evaluations and in Silico Analyses
Since this patient presented LCA phenotypes, we first focused on variants in known disease-causing genes for LCA and other types of inherited retinal degeneration (IRD; RetNet; https://sph.uth.edu/RetNet/home.htm; available in the public domain from The University of Texas Health Science Center, Houston, TX, USA). The WGS strategy reached a mean depth of 29.81-fold, and its detailed capture statistics are listed in Supplementary Table S2. By means of WGS, no potential mutations, including larger deletions or structural aberrations, were identified in any of the reported IRD relevant genes (Supplementary Table S3). WGS revealed that a homozygous variation, IFT52 c.556A>G (p.T186A), was carried by the proband but was absent in the two unaffected siblings (Fig. 2B). IFT52 is a syndromic ciliopathy-causing gene, which is similar to IFT140, CEP290, and IQCB1, another three LCA-causing genes also involved in syndromic ciliopathy.17 
Ift52 was found ubiquitously expressed in multiple murine tissues, including neural retina and RPE (Fig. 2F). The identified variant, located in the seventh exon of IFT52 and the GIFT domain of the IFT52 protein (Fig. 2D), was not revealed in all seven SNP databases and was absent in 100 unrelated controls. This variant, highly conserved among multiple species (Fig. 2C), was predicted to be damaged by both Sorting Intolerant From Tolerant Human Protein DB (score 0.01) and Polymorphism Phenotyping v. 2 (score 0.965) online software. Structural modeling of the wild-type and mutant proteins were further obtained based on the structure of murine IFT52 protein (Protein Data Bank ID: 5fms.1.A) with the sequence identity of 93.63 and similarity of 0.59.18 This substitution was found to generate a novel hydrogen bond between residue 186 and proline at residue 192, which could potentially interrupt the tertiary structure and the stability of the IFT52 protein (Fig. 2E). 
IFT52 p.T186A Might Disturb IFT52 Protein Stability
To determine whether the identified mutation will change the stability and intracellular localization of the IFT52 protein, we transfected IFT52 siRNA together with the FLAG-tagged Ac-IFT52WT/Ac-IFT52T186A plasmids into hTERT-RPE1 cells. We used IFT52 siRNA to knock down the endogenous IFT52 expression. Three pairs of IFT52 siRNAs were initially designed and tested. IFT52 siRNA-2 with best efficiency and stability was selected for further analyses (Fig. 3A). Decreased IFT52 intensity was found in cells transfected with IFT52 siRNA when compared to cells transfected with scramble siRNA, further implying that IFT52 expression was efficiently knocked down by IFT52 siRNA-2 (Fig. 3B). As revealed by immunofluorescence and immunoblotting, both IFT52 and FLAG expressions were decreased in hTERT-RPE1 cells expressing mutant IFT52 protein compared to cells expressing wild-type IFT52 protein (Figs. 3B–3E), implying that IFT52 p.T186A might partly interrupt the stability of encoded IFT52 protein. No obvious change was detected between the intracellular localization of wild-type and mutant IFT52 protein (Fig. 3B). 
Figure 3
 
IFT52 p.T186A mutation interrupts IFT52 protein stability. (A) IFT52 mRNA expressions are remarkably decreased in hTERT-RPE1 cells transfected with IFT52 siRNA-1/IFT52 siRNA-2/IFT52 siRNA-3 compared to cells transfected with scramble siRNA. (B) Immunofluorescence of FLAG (green, primary antibody: anti-mouse FLAG; secondary antibody: Alexa Fluor 488 Goat Anti-Mouse IgG H+L) and IFT52 (red, primary antibody: anti-rabbit IFT52; secondary antibody: Alexa Fluor 594 Donkey Anti-Rabbit IgG H+L) suggests declined IFT52 protein expression in hTERT-RPE1 cells transfected with IFT52 siRNA compared to cells transfected with scramble siRNA and decreased IFT52 and FLAG intensity in cells transfected with IFT52 siRNA plus Ac-IFT52T186A compared to cells transfected with IFT52 siRNA plus Ac-IFT52WT. (CE) Immunoblotting also implies reduced IFT52 protein expression in hTERT-RPE1 cells transfected with IFT52 siRNA plus Ac-IFT52T186A compared to cells transfected with empty vector or with IFT52 siRNA plus Ac-IFT52WT (C, D). FLAG expression is also declined in the IFT52 siRNA plus Ac-IFT52T186A transfected group compared to the IFT52 siRNA plus Ac-IFT52WT transfected group. *P < 0.05, **P < 0.01, ***P < 0.001.
Figure 3
 
IFT52 p.T186A mutation interrupts IFT52 protein stability. (A) IFT52 mRNA expressions are remarkably decreased in hTERT-RPE1 cells transfected with IFT52 siRNA-1/IFT52 siRNA-2/IFT52 siRNA-3 compared to cells transfected with scramble siRNA. (B) Immunofluorescence of FLAG (green, primary antibody: anti-mouse FLAG; secondary antibody: Alexa Fluor 488 Goat Anti-Mouse IgG H+L) and IFT52 (red, primary antibody: anti-rabbit IFT52; secondary antibody: Alexa Fluor 594 Donkey Anti-Rabbit IgG H+L) suggests declined IFT52 protein expression in hTERT-RPE1 cells transfected with IFT52 siRNA compared to cells transfected with scramble siRNA and decreased IFT52 and FLAG intensity in cells transfected with IFT52 siRNA plus Ac-IFT52T186A compared to cells transfected with IFT52 siRNA plus Ac-IFT52WT. (CE) Immunoblotting also implies reduced IFT52 protein expression in hTERT-RPE1 cells transfected with IFT52 siRNA plus Ac-IFT52T186A compared to cells transfected with empty vector or with IFT52 siRNA plus Ac-IFT52WT (C, D). FLAG expression is also declined in the IFT52 siRNA plus Ac-IFT52T186A transfected group compared to the IFT52 siRNA plus Ac-IFT52WT transfected group. *P < 0.05, **P < 0.01, ***P < 0.001.
IFT52 p.T186A Disrupts Cilia Elongation in hTERT-RPE1 Cells
Given the role of IFT52 protein in primary cilium formation and that IFT52 mutation has been previously found to disrupt ciliogenesis, we thus aimed to determine whether the mutation identified in this study would interrupt ciliary function in hTERT-RPE1 cells. Ciliary axonemes and basal bodies were labeled with antibody to Ac-α-tubulin and ARL13B. As we found that IFT52 p.T186A might interrupt the stability of encoded IFT52 protein, we first detected whether IFT52 insufficiency will disturb regular ciliary function. Our findings indicated that ciliary length was reduced in hTERT-RPE1 cells transfected with IFT52 siRNA compared to untreated cells and cells transfected with scramble siRNA (Figs. 4B–4D). No difference in percentages of ciliated cells was detected between cells transfected with scramble siRNA and IFT52 siRNA (Fig. 4A). Thus, our data suggested that IFT52 insufficiency interrupts cilia elongation but not cilia abundance. 
Figure 4
 
IFT52 p.T186A mutation impairs cilia elongation in hTERT-RPE1 cells. (A) Analysis of percentage of ciliated cells in hTERT-RPE1 cells transfected with scramble siRNA (n = 226) and IFT52 siRNA (n = 166). (B) Analysis of primary cilia length in untransfected hTERT-RPE1 cells (Ctrl; n = 48), cells transfected with scramble siRNA (n = 28), cells transfected with IFT52 siRNA (n = 54), cells transfected with IFT52 siRNA and Ac-IFT52WT (n = 51), and cells transfected with IFT52 siRNA and Ac-IFT52T186A (n = 78). (C, D) Immunofluorescence of IFT52 ([C] green, primary antibody: anti-rabbit IFT52; secondary antibody: Alexa Fluor 488 Goat anti-Rabbit IgG H+L), ARL13B ([D] green, primary antibody: anti-rabbit ARL13B; secondary antibody: Alexa Fluor 488 Goat anti-Rabbit IgG H+L), and Ac-α-tubulin (C, D) red, primary antibody: anti-mouse Ac-α-tubulin; secondary antibody: Alexa Fluor 594 Donkey Anti-Mouse IgG H+L shows reduced cilia length in hTERT-RPE1 cells transfected with IFT52 siRNA compared to untreated cells (Ctrl) and cells transfected with scramble siRNA. Scale bar: 10 μm. (E, F) Immunofluorescence of FLAG ([E], green, primary antibody: anti-rabbit FLAG; secondary antibody: Alexa Fluor 488 Goat Anti-Rabbit IgG H+L; [F] green, primary antibody: anti-mouse FLAG; secondary antibody: Alexa Fluor 488 Goat Anti-Mouse IgG H+L), Ac-α-tubulin ([E] red, primary antibody: anti-mouse Ac-α-tubulin; secondary antibody: Alexa Fluor 594 Donkey Anti-Mouse IgG H+L), and ARL13B ([F] red, primary antibody: anti-rabbit ARL13B; secondary antibody: Alexa Fluor 594 Donkey Anti-Rabbit IgG H+L) indicates reduced cilia length in hTERT-RPE1 cells transfected with IFT52 siRNA plus Ac-IFT52T186A compared to cells transfected with IFT52 siRNA plus Ac-IFT52WT. Scale bar: 10 μm. Error bars: SEM. ***P < 0.001.
Figure 4
 
IFT52 p.T186A mutation impairs cilia elongation in hTERT-RPE1 cells. (A) Analysis of percentage of ciliated cells in hTERT-RPE1 cells transfected with scramble siRNA (n = 226) and IFT52 siRNA (n = 166). (B) Analysis of primary cilia length in untransfected hTERT-RPE1 cells (Ctrl; n = 48), cells transfected with scramble siRNA (n = 28), cells transfected with IFT52 siRNA (n = 54), cells transfected with IFT52 siRNA and Ac-IFT52WT (n = 51), and cells transfected with IFT52 siRNA and Ac-IFT52T186A (n = 78). (C, D) Immunofluorescence of IFT52 ([C] green, primary antibody: anti-rabbit IFT52; secondary antibody: Alexa Fluor 488 Goat anti-Rabbit IgG H+L), ARL13B ([D] green, primary antibody: anti-rabbit ARL13B; secondary antibody: Alexa Fluor 488 Goat anti-Rabbit IgG H+L), and Ac-α-tubulin (C, D) red, primary antibody: anti-mouse Ac-α-tubulin; secondary antibody: Alexa Fluor 594 Donkey Anti-Mouse IgG H+L shows reduced cilia length in hTERT-RPE1 cells transfected with IFT52 siRNA compared to untreated cells (Ctrl) and cells transfected with scramble siRNA. Scale bar: 10 μm. (E, F) Immunofluorescence of FLAG ([E], green, primary antibody: anti-rabbit FLAG; secondary antibody: Alexa Fluor 488 Goat Anti-Rabbit IgG H+L; [F] green, primary antibody: anti-mouse FLAG; secondary antibody: Alexa Fluor 488 Goat Anti-Mouse IgG H+L), Ac-α-tubulin ([E] red, primary antibody: anti-mouse Ac-α-tubulin; secondary antibody: Alexa Fluor 594 Donkey Anti-Mouse IgG H+L), and ARL13B ([F] red, primary antibody: anti-rabbit ARL13B; secondary antibody: Alexa Fluor 594 Donkey Anti-Rabbit IgG H+L) indicates reduced cilia length in hTERT-RPE1 cells transfected with IFT52 siRNA plus Ac-IFT52T186A compared to cells transfected with IFT52 siRNA plus Ac-IFT52WT. Scale bar: 10 μm. Error bars: SEM. ***P < 0.001.
To better illustrate the property of the identified mutation, we further tested whether overexpressing of the wild-type and mutant IFT52 protein could rescue the phenotypes caused by IFT52 insufficiency. According to our results, reduction of ciliary length caused by IFT52 insufficiency could almost be completely rescued by coexpressing of wild-type IFT52 in hTERT-RPE1 cells (Figs. 4B, 4E, 4F). However, such interrupted cilia elongation could not be rescued by cotransfection of Ac-IFT52T186A plasmid. Average ciliary length was remarkably reduced in cells overexpressing mutant IFT52 protein compared to cells overexpressing wild-type IFT52 protein and the control group (Figs. 4B, 4E, 4F). To tell whether IFT52 p.T186A functions in a loss-of-function or dominant negative way, we further compared cilia length in hTERT-RPE1 cells transfected with IFT52 siRNA with cells cotransfected with IFT52 siRNA and Ac-IFT52T186A. No difference in cilia length was detected between the two groups (Fig. 4B). We therefore think that IFT52 p.T186A mutation functions in a loss-of-function manner rather than dominant negative. Taken together, the above observations suggested that IFT52 p.T186A mutation interrupts cilia elongation in a loss-of-function way. With these efforts, our data further supported that the detected variant is disease-causing mutation rather than rare polymorphism. 
Discussion
By means of WGS, this report identifies a novel IFT52 mutation, p.T186A, as a candidate for a recessive form of syndromic ciliopathy, presenting mild SRTD and LCA, hence highlighting the importance of IFT-B complex protein in retinal ciliopathy and the promising role of WGS in revealing novel disease-causing genes associate with monogenic disorders. The deleterious nature of the identified IFT52 mutation is supported by an in vivo functional study. Based on our results, this mutation could disturb the stability of encoded IFT52 protein and dramatically disrupt cilia elongation in hTERT-RPE1 cells. WGS has predominant roles in identifying larger deletions and structural aberrations. Herein, by means of WGS, no potential mutations, including larger deletions or structural aberrations, were identified in any of the reported IRD relevant genes. However, it is still possible that some deep intronic variants in known IRD-causing genes may be missed. With exception of this caveat, given the functional evidence taken together with a lack of other explanatory mutations, the IFT52 p.T186A mutation is most likely the candidate for the syndromic ciliopathy involving retinal degeneration and skeletal anomalies for this proband. 
Our study expands the ocular phenotypes caused by IFT52 mutations. The patient shares common but milder features with the two previous patients (Table), including narrow chest with short ribs, micromelic limbs, and sandal gap, but without respiratory distress, hypoplastic corpus callosum, or midface hypoplasia. In initial reports, ophthalmic involvement was found in a 3-year-old patient carrying IFT52 p.R142* with an uncharacterized retinopathy, presenting nystagmus, hypermetropia, and peripheral vision loss.4 However, her visual evoked potential was normal, suggesting that her retinal function was relatively preserved. No further details were revealed. In this study, our case presented typical LCA symptoms, including infantile-onset severe vision loss, nystagmus, oculodigital sign, and completely diminished ERG responses, which, to our knowledge, has not been previously linked to IFT52 mutations. 
Table
 
Summary of Reported Identified IFT52 Mutations
Table
 
Summary of Reported Identified IFT52 Mutations
The IFT52 protein, located near the basal bodies of the cilia, belongs to the IFT complex. IFT complex mutations, which interrupt cilia function, have been reported to be associated with a wide spectrum of retinopathies. Similar to our finding in this study, mutations in the IFT140 and IFT81 genes have been reported to cause syndromic diseases with retinopathy.1922 Patients carrying IFT140 mutations showed Mainzer-Saldino syndrome presenting early onset, severe retinal dystrophy, phalangeal cone-shaped epiphyses, and chronic renal failure.21 A patient carrying mutations in IFT81 presented retinal dystrophy with systemic disorders, including intellectual disability, cerebellar atrophy, and renal problems.19 In addition, IFT81 and IFT140 mutations contribute to nonsyndromic retinopathy,23 further supporting that interrupted IFT function correlates with retinal dysfunction. Meanwhile, IFT81 and IFT140 mutations also correlate with systemic ciliopathy without ocular presentations.19,20 Taken together, the variable ciliopathy phenotypes caused by IFT140 and IFT81 mutations resemble IFT52 mutation–correlated disease spectrum (Table) and support the involvement of IFT52 mutations in retinopathy. 
IFT52 interacts directly with another three IFT-B components (IFT88, IFT70, and IFT46) to form a tetrameric subcomplex of IFT-B core and is essential for the IFT-B complex maintenance and ciliary/flagellar assembly.24 IFT52 mutations could cause reduction and mislocalization of multiple IFT-B proteins, thus destabilizing the IFT-B core and further interrupting the assembly and stability of cilia.3,25 The above-mentioned IFT81 protein is also a core component of IFT-B complex, thus further highlights the crucial role of IFT-B in maintaining regular retinal function. Cilia are responsible for multiple biological processes in various tissues. Since cilia are devoid of protein synthesis, the IFT-mediated translocation of proteins is required to maintain regular ciliary functions, including in photoreceptor outer segments.26 Photoreceptors are found to degenerate following loss of cilia.27 Previous study has indicated that knocking down of ift52 in zebrafish could cause ectopic accumulation of opsins and subsequent photoreceptor degeneration.27 To date, 25 genes have been reported to be involved in LCA etiology (RetNet), among which three (IFT140, CEP290, and IQCB1) are syndromic ciliopathy genes like IFT52.17 A potential explanation for the diverse genotype-phenotype correlations of ciliary genes is a combination of their multifunctional natures and differential damaging effect of their mutant alleles.17 Further confirmative in vivo studies and screening for IFT52 mutations in more LCA patients are still needed. 
Taken together, our study reveals a novel IFT52 mutation, p.T186A, and expands phenotypes of IFT52 mutation-caused ciliopathy to include LCA. We also demonstrate the pathogenic nature of the identified mutation, which might disturb the stability of the encoded IFT52 protein and interrupt cilia elongation in hTERT-RPE1 cells. Our findings suggest that IFT52 mutation should be screened in LCA patients, and all patients with IFT52 mutations should receive comprehensive ophthalmic evaluations. 
Acknowledgments
Supported by the National Natural Science Foundation of China (81525006, 81670864, and 81730025 [CZ]; 81760180 [XS]; and 81700877 [XC]); the Jiangsu Province's Innovation Team (CZ); Natural Science Foundation of Jiangsu Province (BK20171087 [XC]); Shanghai Outstanding Academic Leaders (2017BR013 [CZ]); the Key Technology R&D Program of Ningxia Province (2014ZYH65 [XS]); Open Foundation of State Key Laboratory of Reproductive Medicine (Nanjing Medical University) (SKLRM-KA201607 [XC]); and a project funded by the Priority Academic Program Development of Jiangsu Higher Education Institutions. 
Disclosure: X. Chen, None; X. Wang, None; C. Jiang, None; M. Xu, None; Y. Liu, None; R. Qi, None; X. Qi, None; X. Sun, None; P. Xie, None; Q. Liu, None; B. Yan, None; X. Sheng, None; C. Zhao, None 
References
Hildebrandt F, Benzing T, Katsanis N. Ciliopathies. N Engl J Med. 2011; 364: 1533–1543.
Rosenbaum JL, Witman GB. Intraflagellar transport. Nat Rev Mol Cell Biol. 2002; 3: 813–825.
Zhang W, Taylor SP, Nevarez L, et al. IFT52 mutations destabilize anterograde complex assembly, disrupt ciliogenesis and result in short rib polydactyly syndrome. Hum Mol Genet. 2016; 25: 4012–4020.
Girisha KM, Shukla A, Trujillano D, et al. A homozygous nonsense variant in IFT52 is associated with a human skeletal ciliopathy. Clin Genet. 2016; 90: 536–539.
Bujakowska KM, Zhang Q, Siemiatkowska AM, et al. Mutations in IFT172 cause isolated retinal degeneration and Bardet-Biedl syndrome. Hum Mol Genet. 2015; 24: 230–242.
Huber C, Cormier-Daire V. Ciliary disorder of the skeleton. Am J Med Genet C Semin Med Genet. 2012; 160C: 165–174.
Schmidts M, Vodopiutz J, Christou-Savina S, et al. Mutations in the gene encoding IFT dynein complex component WDR34 cause Jeune asphyxiating thoracic dystrophy. Am J Hum Genet. 2013; 93: 932–944.
Fu Q, Xu M, Chen X, et al. CEP78 is mutated in a distinct type of Usher syndrome [published online ahead of print September 14, 2016]. J Med Genet. http://dx.doi.org/10.1136/jmedgenet-2016-104166.
Li H, Durbin R. Fast and accurate short read alignment with Burrows-Wheeler transform. Bioinformatics. 2009; 25: 1754–1760.
Challis D, Yu J, Evani US, et al. An integrative variant analysis suite for whole exome next-generation sequencing data. BMC Bioinformatics. 2012; 13: 8.
Zhao C, Lu S, Zhou X, Zhang X, Zhao K, Larsson C. A novel locus (RP33) for autosomal dominant retinitis pigmentosa mapping to chromosomal region 2cen-q12.1. Hum Genet. 2006; 119: 617–623.
Zhao C, Bellur DL, Lu S, et al. Autosomal-dominant retinitis pigmentosa caused by a mutation in SNRNP200, a gene required for unwinding of U4/U6 snRNAs. Am J Hum Genet. 2009; 85: 617–627.
Chen X, Liu Y, Sheng X, et al. PRPF4 mutations cause autosomal dominant retinitis pigmentosa. Hum Mol Genet. 2014; 23: 2926–2939.
Liu Y, Chen X, Xu Q, et al. SPP2 mutations cause autosomal dominant retinitis pigmentosa. Sci Rep. 2015; 5: 14867.
Chen X, Jiang C, Qin B, et al. LncRNA ZNF503-AS1 promotes RPE differentiation by downregulating ZNF503 expression. Cell Death Dis. 2017; 8: e3046.
Ziegler EE. 4.2 The CDC and Euro Growth Charts. World Rev Nutr Diet. 2015; 113: 295–307.
Xu M, Yang L, Wang F, et al. Mutations in human IFT140 cause non-syndromic retinal degeneration. Hum Genet. 2015; 134: 1069–1078.
Taschner M, Weber K, Mourao A, et al. Intraflagellar transport proteins 172, 80, 57, 54, 38, and 20 form a stable tubulin-binding IFT-B2 complex. EMBO J. 2016; 35: 773–790.
Perrault I, Halbritter J, Porath JD, et al. IFT81, encoding an IFT-B core protein, as a very rare cause of a ciliopathy phenotype. J Med Genet. 2015; 52: 657–665.
Duran I, Taylor SP, Zhang W, et al. Destabilization of the IFT-B cilia core complex due to mutations in IFT81 causes a spectrum of short-rib polydactyly syndrome. Sci Rep. 2016; 6: 34232.
Perrault I, Saunier S, Hanein S, et al. Mainzer-Saldino syndrome is a ciliopathy caused by IFT140 mutations. Am J Hum Genet. 2012; 90: 864–870.
Schmidts M, Frank V, Eisenberger T, et al. Combined NGS approaches identify mutations in the intraflagellar transport gene IFT140 in skeletal ciliopathies with early progressive kidney disease. Hum Mutat. 2013; 34: 714–724.
Dharmat R, Liu W, Ge Z, et al. IFT81 as a candidate gene for nonsyndromic retinal degeneration. Invest Ophthalmol Vis Sci. 2017; 58: 2483–2490.
Brazelton WJ, Amundsen CD, Silflow CD, Lefebvre PA. The bld1 mutation identifies the Chlamydomonas osm-6 homolog as a gene required for flagellar assembly. Curr Biol. 2001; 11: 1591–1594.
Richey EA, Qin H. Dissecting the sequential assembly and localization of intraflagellar transport particle complex B in Chlamydomonas. PLoS One. 2012; 7: e43118.
Rosenbaum JL, Cole DG, Diener DR. Intraflagellar transport: the eyes have it. J Cell Biol. 1999; 144: 385–388.
Tsujikawa M, Malicki J. Intraflagellar transport genes are essential for differentiation and survival of vertebrate sensory neurons. Neuron. 2004; 42: 703–716.
Figure 1
 
Systemic evaluations of patient YWH-IV:1. (A, B) Fundus photos of patient YWH-IV:1 suggest waxy optic disk and attenuated retinal vessels. Sporadic pigment deposits are also noticed in the midperipheral retina (indicated by arrow). (C, D) OCT examinations of patient YWH-IV:1 reveal loss of ellipsoid in bilateral macular region. (E) Both scotopic and photopic ERG responses are diminished. (F) Patient YWH-IV:1 presents small chest with short ribs and micromelic limbs. (G) Sandal gap is found in her right foot, between her fourth and fifth phalanges. (H, I) Dental dysplasia is identified. Panoramic dental X-ray suggests that one left mandibular premolar, one right mandibular premolar, and one right mandibular incisor are missing (I).
Figure 1
 
Systemic evaluations of patient YWH-IV:1. (A, B) Fundus photos of patient YWH-IV:1 suggest waxy optic disk and attenuated retinal vessels. Sporadic pigment deposits are also noticed in the midperipheral retina (indicated by arrow). (C, D) OCT examinations of patient YWH-IV:1 reveal loss of ellipsoid in bilateral macular region. (E) Both scotopic and photopic ERG responses are diminished. (F) Patient YWH-IV:1 presents small chest with short ribs and micromelic limbs. (G) Sandal gap is found in her right foot, between her fourth and fifth phalanges. (H, I) Dental dysplasia is identified. Panoramic dental X-ray suggests that one left mandibular premolar, one right mandibular premolar, and one right mandibular incisor are missing (I).
Figure 2
 
Family pedigree and genetic annotation of the identified mutation. (A) Pedigree of family YWH. +: wild-type allele; MU: IFT52 c.556A>G (p.T186A). (B) Sequence chromatograms of the identified mutation. (C) Orthologous protein sequence alignment of IFT52 from humans (H. sapiens), chimpanzees (P. troglodytes), dogs (C. lupus), cows (B. taurus), pigs (S. scrofa), mice (M. musculus), chickens (G. gallus), zebrafish (D. rerio), and fruit flies (D. melanogaster). Conserved residues are shaded. The mutated residue 186 is boxed and indicated. (D) Schematic representation of the relative linear location of the identified IFT52 mutation in context of genome structure (upper) and protein structure (below). Regions of IFT52 protein that interact directly with another three IFT-B components (IFT88, IFT70, and IFT46) are indicated. (E) Crystal structural analysis of the wild-type and mutant IFT52 protein. A novel hydrogen bond between residues 186 and 192 is generated by the mutation. (F) Expression of Ift52 in multiple murine tissues including heart, liver, spleen, lung, kidney, brain, muscle, stomach, intestines, colorectum, neural retina, RPE, and optic nerve is demonstrated. A 243-bp PCR product of the murine Ift52 (top panel) is detected in all tested tissues. PCR product of the murine Rplp0 (109 bp) is analyzed in parallel as a loading control.
Figure 2
 
Family pedigree and genetic annotation of the identified mutation. (A) Pedigree of family YWH. +: wild-type allele; MU: IFT52 c.556A>G (p.T186A). (B) Sequence chromatograms of the identified mutation. (C) Orthologous protein sequence alignment of IFT52 from humans (H. sapiens), chimpanzees (P. troglodytes), dogs (C. lupus), cows (B. taurus), pigs (S. scrofa), mice (M. musculus), chickens (G. gallus), zebrafish (D. rerio), and fruit flies (D. melanogaster). Conserved residues are shaded. The mutated residue 186 is boxed and indicated. (D) Schematic representation of the relative linear location of the identified IFT52 mutation in context of genome structure (upper) and protein structure (below). Regions of IFT52 protein that interact directly with another three IFT-B components (IFT88, IFT70, and IFT46) are indicated. (E) Crystal structural analysis of the wild-type and mutant IFT52 protein. A novel hydrogen bond between residues 186 and 192 is generated by the mutation. (F) Expression of Ift52 in multiple murine tissues including heart, liver, spleen, lung, kidney, brain, muscle, stomach, intestines, colorectum, neural retina, RPE, and optic nerve is demonstrated. A 243-bp PCR product of the murine Ift52 (top panel) is detected in all tested tissues. PCR product of the murine Rplp0 (109 bp) is analyzed in parallel as a loading control.
Figure 3
 
IFT52 p.T186A mutation interrupts IFT52 protein stability. (A) IFT52 mRNA expressions are remarkably decreased in hTERT-RPE1 cells transfected with IFT52 siRNA-1/IFT52 siRNA-2/IFT52 siRNA-3 compared to cells transfected with scramble siRNA. (B) Immunofluorescence of FLAG (green, primary antibody: anti-mouse FLAG; secondary antibody: Alexa Fluor 488 Goat Anti-Mouse IgG H+L) and IFT52 (red, primary antibody: anti-rabbit IFT52; secondary antibody: Alexa Fluor 594 Donkey Anti-Rabbit IgG H+L) suggests declined IFT52 protein expression in hTERT-RPE1 cells transfected with IFT52 siRNA compared to cells transfected with scramble siRNA and decreased IFT52 and FLAG intensity in cells transfected with IFT52 siRNA plus Ac-IFT52T186A compared to cells transfected with IFT52 siRNA plus Ac-IFT52WT. (CE) Immunoblotting also implies reduced IFT52 protein expression in hTERT-RPE1 cells transfected with IFT52 siRNA plus Ac-IFT52T186A compared to cells transfected with empty vector or with IFT52 siRNA plus Ac-IFT52WT (C, D). FLAG expression is also declined in the IFT52 siRNA plus Ac-IFT52T186A transfected group compared to the IFT52 siRNA plus Ac-IFT52WT transfected group. *P < 0.05, **P < 0.01, ***P < 0.001.
Figure 3
 
IFT52 p.T186A mutation interrupts IFT52 protein stability. (A) IFT52 mRNA expressions are remarkably decreased in hTERT-RPE1 cells transfected with IFT52 siRNA-1/IFT52 siRNA-2/IFT52 siRNA-3 compared to cells transfected with scramble siRNA. (B) Immunofluorescence of FLAG (green, primary antibody: anti-mouse FLAG; secondary antibody: Alexa Fluor 488 Goat Anti-Mouse IgG H+L) and IFT52 (red, primary antibody: anti-rabbit IFT52; secondary antibody: Alexa Fluor 594 Donkey Anti-Rabbit IgG H+L) suggests declined IFT52 protein expression in hTERT-RPE1 cells transfected with IFT52 siRNA compared to cells transfected with scramble siRNA and decreased IFT52 and FLAG intensity in cells transfected with IFT52 siRNA plus Ac-IFT52T186A compared to cells transfected with IFT52 siRNA plus Ac-IFT52WT. (CE) Immunoblotting also implies reduced IFT52 protein expression in hTERT-RPE1 cells transfected with IFT52 siRNA plus Ac-IFT52T186A compared to cells transfected with empty vector or with IFT52 siRNA plus Ac-IFT52WT (C, D). FLAG expression is also declined in the IFT52 siRNA plus Ac-IFT52T186A transfected group compared to the IFT52 siRNA plus Ac-IFT52WT transfected group. *P < 0.05, **P < 0.01, ***P < 0.001.
Figure 4
 
IFT52 p.T186A mutation impairs cilia elongation in hTERT-RPE1 cells. (A) Analysis of percentage of ciliated cells in hTERT-RPE1 cells transfected with scramble siRNA (n = 226) and IFT52 siRNA (n = 166). (B) Analysis of primary cilia length in untransfected hTERT-RPE1 cells (Ctrl; n = 48), cells transfected with scramble siRNA (n = 28), cells transfected with IFT52 siRNA (n = 54), cells transfected with IFT52 siRNA and Ac-IFT52WT (n = 51), and cells transfected with IFT52 siRNA and Ac-IFT52T186A (n = 78). (C, D) Immunofluorescence of IFT52 ([C] green, primary antibody: anti-rabbit IFT52; secondary antibody: Alexa Fluor 488 Goat anti-Rabbit IgG H+L), ARL13B ([D] green, primary antibody: anti-rabbit ARL13B; secondary antibody: Alexa Fluor 488 Goat anti-Rabbit IgG H+L), and Ac-α-tubulin (C, D) red, primary antibody: anti-mouse Ac-α-tubulin; secondary antibody: Alexa Fluor 594 Donkey Anti-Mouse IgG H+L shows reduced cilia length in hTERT-RPE1 cells transfected with IFT52 siRNA compared to untreated cells (Ctrl) and cells transfected with scramble siRNA. Scale bar: 10 μm. (E, F) Immunofluorescence of FLAG ([E], green, primary antibody: anti-rabbit FLAG; secondary antibody: Alexa Fluor 488 Goat Anti-Rabbit IgG H+L; [F] green, primary antibody: anti-mouse FLAG; secondary antibody: Alexa Fluor 488 Goat Anti-Mouse IgG H+L), Ac-α-tubulin ([E] red, primary antibody: anti-mouse Ac-α-tubulin; secondary antibody: Alexa Fluor 594 Donkey Anti-Mouse IgG H+L), and ARL13B ([F] red, primary antibody: anti-rabbit ARL13B; secondary antibody: Alexa Fluor 594 Donkey Anti-Rabbit IgG H+L) indicates reduced cilia length in hTERT-RPE1 cells transfected with IFT52 siRNA plus Ac-IFT52T186A compared to cells transfected with IFT52 siRNA plus Ac-IFT52WT. Scale bar: 10 μm. Error bars: SEM. ***P < 0.001.
Figure 4
 
IFT52 p.T186A mutation impairs cilia elongation in hTERT-RPE1 cells. (A) Analysis of percentage of ciliated cells in hTERT-RPE1 cells transfected with scramble siRNA (n = 226) and IFT52 siRNA (n = 166). (B) Analysis of primary cilia length in untransfected hTERT-RPE1 cells (Ctrl; n = 48), cells transfected with scramble siRNA (n = 28), cells transfected with IFT52 siRNA (n = 54), cells transfected with IFT52 siRNA and Ac-IFT52WT (n = 51), and cells transfected with IFT52 siRNA and Ac-IFT52T186A (n = 78). (C, D) Immunofluorescence of IFT52 ([C] green, primary antibody: anti-rabbit IFT52; secondary antibody: Alexa Fluor 488 Goat anti-Rabbit IgG H+L), ARL13B ([D] green, primary antibody: anti-rabbit ARL13B; secondary antibody: Alexa Fluor 488 Goat anti-Rabbit IgG H+L), and Ac-α-tubulin (C, D) red, primary antibody: anti-mouse Ac-α-tubulin; secondary antibody: Alexa Fluor 594 Donkey Anti-Mouse IgG H+L shows reduced cilia length in hTERT-RPE1 cells transfected with IFT52 siRNA compared to untreated cells (Ctrl) and cells transfected with scramble siRNA. Scale bar: 10 μm. (E, F) Immunofluorescence of FLAG ([E], green, primary antibody: anti-rabbit FLAG; secondary antibody: Alexa Fluor 488 Goat Anti-Rabbit IgG H+L; [F] green, primary antibody: anti-mouse FLAG; secondary antibody: Alexa Fluor 488 Goat Anti-Mouse IgG H+L), Ac-α-tubulin ([E] red, primary antibody: anti-mouse Ac-α-tubulin; secondary antibody: Alexa Fluor 594 Donkey Anti-Mouse IgG H+L), and ARL13B ([F] red, primary antibody: anti-rabbit ARL13B; secondary antibody: Alexa Fluor 594 Donkey Anti-Rabbit IgG H+L) indicates reduced cilia length in hTERT-RPE1 cells transfected with IFT52 siRNA plus Ac-IFT52T186A compared to cells transfected with IFT52 siRNA plus Ac-IFT52WT. Scale bar: 10 μm. Error bars: SEM. ***P < 0.001.
Table
 
Summary of Reported Identified IFT52 Mutations
Table
 
Summary of Reported Identified IFT52 Mutations
Supplement 1
Supplement 2
Supplement 3
×
×

This PDF is available to Subscribers Only

Sign in or purchase a subscription to access this content. ×

You must be signed into an individual account to use this feature.

×