CEP41 is mutated in Joubert syndrome and is required for tubulin glutamylation at the cilium.

Abstract

Tubulin glutamylation is a post-translational modification that occurs predominantly in the ciliary axoneme and has been suggested to be important for ciliary function. However, its relationship to disorders of the primary cilium, termed ciliopathies, has not been explored. Here we mapped a new locus for Joubert syndrome (JBTS), which we have designated as JBTS15, and identified causative mutations in CEP41, which encodes a 41-kDa centrosomal protein. We show that CEP41 is localized to the basal body and primary cilia, and regulates ciliary entry of TTLL6, an evolutionarily conserved polyglutamylase enzyme. Depletion of CEP41 causes ciliopathy-related phenotypes in zebrafish and mice and results in glutamylation defects in the ciliary axoneme. Our data identify CEP41 mutations as a cause of JBTS and implicate tubulin post-translational modification in the pathogenesis of human ciliary dysfunction.

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CEP41 is mutated in Joubert syndrome and is required for
tubulin glutamylation at the cilium
Ji Eun Lee1, Jennifer L. Silhavy1, Maha S. Zaki2, Jana Schroth1, Stephanie L. Bielas1, Sarah
E. Marsh1, Jesus Olvera1, Francesco Brancati3,4, Miriam Iannicelli3, Koji Ikegami12, Andrew
M. Schlossman1, Barry Merriman5, Tania Attié-Bitach6, Clare V. Logan7, Ian A. Glass8,
Andrew Cluckey9, Carrie M. Louie1, Jeong Ho Lee1, Hilary R. Raynes10, Isabelle Rapin10,
Ignacio P. Castroviejo11, Mitsutoshi Setou12, Clara Barbot13, Eugen Boltshauser14, Stanley
F. Nelson5, Friedhelm Hildebrandt9, Colin A. Johnson7, Daniel A. Doherty8, Enza Maria
Valente3,15, and Joseph G. Gleeson1
1Neurogenetics Laboratory, Institute for Genomic Medicine, Department of Neurosciences and
Pediatrics, Howard Hughes Medical Institute, University of California, San Diego, CA 92093, USA
2Clinical Genetics Department, Human Genetics and Genome Research Division, National
Research Centre, El-Tahrir St., Dokki, Giza, 12311 Cairo, Egypt 3Casa Sollievo della Sofferenza
Hospital, CSS-Mendel Laboratory, 71013 San Giovanni Rotondo (FG), Italy 4Medical Genetics
Unit, Department of Biopathology and Diagnostic Imaging, Tor Vergata University, 00133 Rome,
Italy 5Department of Human Genetics, Pathology and Laboratory Medicine, David Geffen School
of Medicine, University of California, Los Angeles CA 90095, USA 6Département de Génétique,
INSERM U781, Hôpital Necker-Enfants Malades, Université Paris Descartes, Paris, France
7Section of Ophthalmology and Neurosciences, Wellcome Trust Brenner Building, Leeds Institute
of Molecular Medicine, St James’s University Hospital, Leeds, LS9 7TF, UK 8Divisions of
Developmental Medicine and Genetic Medicine, Department of Pediatrics, University of
Washington Seattle Children’s Hospital, 4800 Sand Point Way NE, Seattle, WA 98105, USA
9Department of Pediatrics and Communicable Diseases, Howard Hughes Medical Institute,
University of Michigan, Ann Arbor, Michigan, USA 10Saul R. Korey Department of Neurology,
Department of Pediatrics, and Rose F. Kennedy Intellectual and Developmental Disabilities
Research Center, Albert Einstein College of Medicine, Bronx, N. Y. 10461, USA 11Pediatric
Neurology Service, University Hospital La Paz, Paseo de la Castellana 261, Madrid 28046, Spain
12Department of Cell Biology and Anatomy, Hamamatsu University School of Medicine, 1-20-1
Handayama, Hamamatsu 431-3192, Japan 13Serviço de Neuropediatria, Hospital de Crianças
Maria Pia, Rua da Boavista, 827, 4050-111 Porto, Portugal 14Deparmtent of Pediatric Neurology,
University Children’s Hospital of Zürich, 75, CH 8032, Switzerland 15Department of Medical and
Surgical Pediatric Sciences, University of Messina, 98125 Messina, Italy
Abstract
Tubulin glutamylation is a post-translational modification (PTM) occurring predominantly on
ciliary axonemal tubulin and has been suggested to be important for ciliary function 1,2. However,
Correspondence should be addressed to J.G.G. (jogleeson@ucsd.edu).
AUTHOR CONTRIBUTIONS
J.E.L. M.S.Z. and J.G.G. designed the study and experiments with substantial contributions from B.M. and S.F.N. helped fine
mapping, J.L.S., S.L.B., J.O., F.B., M.I., A.M.S., T.A.-B., C.V.L., I.A.G., A.C., F.H., C.A.J., D.A.D., and E.M.V. performed genetic
screening, and J.E.L., J.L.S., J.S., J.O., F.B., M.I., T.A.-B., I.A.G., D.A.D., C.M.L., and J.H.L. performed mutation analysis. M.S.Z.,
S.E.M., H.R.R., I.R., I.P.C., E.B. and E.M.V. identified and recruited patients, K.I. and M.S. shared critical reagents, and J.S. helped
genotyping of mutant mice. J.E.L. performed microscopy, biochemical assays, zebrafish and mouse experiments. J.E.L. and J.G.G.
interpreted the data and wrote the manuscript.
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its relationship to disorders of the primary cilium, termed ‘ciliopathies’, has not been explored.
Here, in Joubert syndrome (JBTS) 3, we identify the JBTS15 locus and the responsible gene as
CEP41, encoding a centrosomal protein of 41 KDa 4. We show that CEP41 is localized to the
basal body/primary cilium, and regulates the ciliary entry of TTLL6, an evolutionarily conserved
polyglutamylase enzyme 5. Depletion of CEP41 causes ciliopathy-related phenotypes in zebrafish
and mouse, and induces cilia axonemal glutamylation defects. Our data identify loss of CEP41 as
a cause of JBTS ciliopathy and highlight involvement of tubulin PTM in pathogenesis of the
ciliopathy spectrum.
Joubert syndrome (OMIM 213300) is characterized by cerebellar hypoplasia, and
neurological features including ataxia, psychomotor delay and oculomotor apraxia with a
pathognomonic “molar tooth sign” on brain imaging. JBTS is frequently accompanied by
various multiorgan signs and symptoms including retinal dystrophy, nephronophthisis, liver
fibrosis and polydactyly, conditions associated with disorders of the ciliopathy spectrum of
diseases that include Meckel-Gruber Syndrome (MKS), Bardet-Biedl Syndrome (BBS) and
Nephronophthisis (NPHP). Though several causative genes have been found for these
disorders, they account for less than 50% of cases 6,7. We recruited a consanguineous two-
branch Egyptian family (MTI-429) with five affected members (Fig. 1a–b, Table 1). We
excluded linkage to previously identified JBTS loci using a panel of highly informative
markers. Analysis of the family using whole genome Illumina 5K SNP Linkage chip Ver. IV
scan identified a 5 Mbp region of linkage on chromosome 7q31.33–32.3, with a peak
multipoint LOD score of 3.71, thus defining the JBTS15 locus. Haplotype analysis
suggested a candidate interval between rs766240 and rs4728251 delineating the peak of
highest significance (Supplementary Fig. 1a–b).
In order to further narrow the interval, we re-analyzed MTI-429 with the denser Affymetrix
250K NspI SNP array by applying a linkage-free IBD model 8. The combination of the 5K
SNP and 250K SNP linkage analyses generated a 2.8 Mbp IBD interval between
rs17165226 and rs2971773 containing 26 genes (Fig. 1c). Direct sequence analyses of
candidate genes within the interval led to the identification of a homozygous c.33+2T>G
base change from reference sequence NM_018718, which was predicted to abolish the
consensus splice donor site from exon 1 of the CEP41 gene (Fig. 1d, Supplementary Fig.
2a). To confirm a mutation-specific splicing defect, we evaluated CEP41 transcripts from
MTI-429 primary patient fibroblasts (MTI-429-IV-1 and -IV-6). The RT-PCR result showed
an absence of mature CEP41 mRNA products in both affected patient cells, likely attributed
to nonsense mediated decay (Fig. 1e).
We next screened an additional 832 ciliopathy patients: 720 of JBTS and 112 of MKS
(many of whom were excluded for mutations in known ciliopathy genes) by directly
sequencing CEP41 and found two additional consanguineous families with homozygous
mutations: c.97+3_5delGAG in an Egyptian JBTS family (MTI-1491) and c.423-2A>C in a
Portuguese JBTS family (COR-98) (Fig. 1b–d, Supplementary Fig. 2a). These mutations
were predicted to abolish the consensus splice donor site from exon 2 and the splice acceptor
site from exon 7, respectively. Moreover, we confirmed that the mutation in MTI-1491 led
to skipping of exon 2, thereby generating a premature stop in exon 3 (Supplementary Fig.
2b). Interestingly, in addition to the JBTS patients, the MTI-1491 family included one
individual that was consistent with a phenotype of BBS and lacked the pathognomonic
“molar tooth sign”, and this patient was heterozygous for the c.97+3_5delGAG mutation
(Supplementary Fig. 2b–d), suggesting CEP41 may modify other ciliopathy conditions.
From our cohort screen, we further identified heterozygous CEP41 mutations (c.83C>A, c.
107T>C, c.265C>G, c.536G>A, c.1078C>T), which altered highly conserved amino acid
residues among vertebrates or led to a premature stop codon from five different families
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(Fig. 1d, Supplementary Fig. 2e, Supplementary Table 1). Each of these CEP41-mutated
patients was additionally sequenced at the known JBTS genes, and in four there was an
additional heterozygous potentially deleterious variant. It was notable that all homozygous
mutations in CEP41 were splice site mutations and identified only in JBTS patients while
heterozygous variants were present in several ciliopathies including BBS and MKS. Our
findings suggest that constitutive disruptions of CEP41 result in JBTS, but that it may also
serve as a modifier in the broader class of ciliopathies.
The CEP41 gene has been poorly characterized except for its expression analysis in human
organs including brain, testis and kidney 9. CEP41 encodes for Centrosomal Protein 41
KDa, predicted to contain two coiled-coils and a rhodanese-like domain (RHOD), which is
structurally related to the catalytic subunit of the Cdc25 class of phosphatases 10. However,
we found that it lacks phosphatase activity in the in vitro para-Nitrophenylphosphate (pNPP)
phosphatase assay (not shown). The RHOD domain therefore may be an enzymatically
inactive version like some described RHOD domains in other proteins 10 functioning in
protein interactions.
We next examined CEP41 gene expression at the mRNA level and CEP41 protein
subcellular localization. In zebrafish, cep41 was expressed in the various ciliary organs
including Kupffer’s vesicle (KV), ear and heart as well as brain and kidney, regions
predominantly affected in JBTS (Fig. 2a, Supplementary Fig. 3). In several ciliated cell lines
such as mouse inner medullary collecting duct (IMCD3) cells and human retinal pigment
epithelial (hTERT-RPE1) cells, endogenous CEP41 was predominantly noted at the
centrioles and cilia (Fig. 2b). The cilia-associated expression/localization of CEP41
prompted us to assess a possible role in cilia-related function. Accordingly, we performed
knockdown experiments using translation blocking morpholino anti-sense oligonucleotides
(MOs) in zebrafish. In cep41 MOs-injected embryos (morphants), we observed peripheral
heart edema and tail defects along with ciliopathy-related phenotypes including
hydrocephalus, abnormal ear otolith formation and smaller eyes 11–14 (Supplementary Fig.
4a–b). We further found that injection of cep41 MOs induced a decrease in production of the
protein with a dose-dependent phenotypic severity in the embryos (Supplementary Fig. 4b–
c).
Cilia of KV, a structure that corresponds to the mammalian embryonic node, are essential to
mediate lateral asymmetry 15. Accordingly, the defect of left/right (L/R) asymmetry in the
heart, a well-established ciliopathy phenotype in mammals, is also shown in zebrafish 16–18.
To examine whether cep41 depletion results in the phenotype, we injected cep41 MOs into
Tg(myl7:egfp) zebrafish, a myocardium-specific transgenic reporter line, and found heart
asymmetry defects such as inversion or failure to develop asymmetry of the ventricle and
atrium (Fig. 2c). We also generated a Cep41 knockout mouse line using a genetrap strategy
(Supplementary Fig. 5a–c) and characterized its phenotype at E10–13. The homozygous
Cep41Gt/Gt embryos showed a range of phenotypes: malformed hindbrain, exencephaly,
brain hemorrhage, dilated pericardial sac and lethality as well as unexpected normal
development in some homozygous mutants (Fig. 2d, Supplementary Fig. 5d–e,
Supplementary Table 2). Although exencephaly, dilated heart and embryonic lethality
suggest possible ciliary roles of Cep41 in mouse 16,19,20, the phenotypic variability
including normal development suggests the presence of extragenic phenotypic modifiers.
We next investigated a genetic rescue using human CEP41 in zebrafish cep41 morphants,
and found partial rescue of the cilia-associated morphant phenotype (Fig. 2e). The data
suggest a potential evolutionarily conserved role of CEP41 in ciliary function.
In order to explore possible roles of CEP41, we examined primary cultured fibroblasts of
patients (MTI-429-IV-1 and -IV-6). We first tested whether CEP41 mutant cells were
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devoid of CEP41 protein. Consistent with RT-PCR results (Fig. 1e), neither patient
fibroblast line tested produced detectable CEP41, suggesting that these mutant cells are
nearly null for CEP41 (Fig. 3a–b). In order to examine the effect of CEP41 loss on cilia
assembly, we induced ciliogenesis using serum starvation-mediated cell cycle arrest in
confluent cells and visualized cilia by co-immunostaining using anti-ARL13B (cilia
marker) 21 and GT335 (centrioles/cilia marker) 22 antibodies. In control fibroblasts, cilia
were evident in 70 % of the total stained cells by 48 hr and 72 hr and nearly all co-stained
with both markers (Fig. 3c, Supplementary Fig. 6). However, in the CEP41 mutant
fibroblasts, cilia were stained positively with ARL13B but not with GT335 (Fig. 3c). We
quantified this effect and found that, whereas the percent of ARL13B-positive ciliated cells
in the mutant fibroblasts was approximately equal to control fibroblasts, the percentage of
GT335-positive ciliated cells was dramatically reduced in mutant fibroblasts
(Supplementary Fig. 6). The GT335 antibody was originally generated to recognize the
glutamylated forms (both mono- and polyglutamylation) of tubulin 23. Therefore, the data
suggest a potential role of CEP41 in regulating tubulin glutamyl posttranslational
modifications (PTMs).
Microtubules are the major structural scaffolds of the ciliary axoneme, composing the 9+0
or 9+2 arrangement, and undergo several PTMs including acetylation, detyrosination,
glycylation and glutamylation (Supplementary Fig. 7). We therefore tested the effect of
CEP41 deficiency on these tubulin PTMs and found no significant defects other than
glutamylation in CEP41 mutant cells (Supplementary Fig. 8). In addition, the result of
immunostaining using PolyE antibody (specific for polyglutamylated tubulins) in mutant
fibroblasts suggested that CEP41 might function in regulating both tubulin mono- and
polyglutamylation (Supplementary Fig. 9). Concurrently, we observed an effect of cep41
deficiency on tubulin glutamylation as well as mildly reduced glycylation of the placode
cilia in zebrafish (Fig. 3d, Supplementary Fig. 10). Forced expression of exogenous CEP41
in the mutant fibroblasts remarkably increased the percentage of cells displaying
glutamylated cilia (Fig. 3e), suggesting that glutamylation of the cilium is dependent on the
expression of CEP41. Furthermore, we found that cell lines from patients with mutations in
other JBTS genes including TMEM216 and INPP5E displayed no such glutamylation defect
(not shown), suggesting the phenotype of tubulin glutamylation phenotype is not a non-
specific consequence of ciliopathy mutation. Together these data suggest that CEP41 is
required for ciliary glutamylation, but not necessary for initial cilia assembly.
Recent studies have shown that defective tubulin PTM is associated with altered ciliary
axonemal structure 24–26. Accordingly, we investigated whether the CEP41-dysfunctional
cilia exhibiting glutamylation defects gave rise to abnormal axonemal structure.
Transmission electron microscopy (TEM) analysis demonstrated that the depletion of cep41
resulted in apparent structural defects in zebrafish renal cilia: specifically A tubules of the
outer doublet microtubules of the axoneme were collapsed and/or duplicated (Fig. 3f,
Supplementary Fig. 11). Previous studies have suggested that ciliary structural disruption
affects ciliary motility 27,28, thus we examined the effect on the cilia of KV and kidney in
zebrafish cep41 morphants and found disabled motility of both cilia (Supplementary Fig. 12,
Supplementary Mov. 1–6). Our data suggest that CEP41 is involved in ciliary structural
formation and motility by playing an essential role in tubulin glutamylation at the cilium.
We next investigated how CEP41 modulates microtubule PTM at the ciliary axoneme. We
noted that only microtubules of ciliary axonemes failed glutamylation, whereas those of
centrioles were properly modified in CEP41 mutant patient cells. (Fig. 3c). Furthermore, the
lack of a Tubulin Tyrosine Ligase (TTL) domain in CEP41, requisite for enzyme activity 5,
implied that it is unlikely to serve as a glutamylase. The main enzymes mediating tubulin
glutamylation are members of the conserved TTL-like (TTLL) family 5. Among several
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identified TTLL factors, we found consistently strong localization of TTLL6 at the basal
body (the organizing structure at the base of cilium, derived from a mother centriole) and the
cilium (Supplementary Fig. 13). In addition, previous studies have suggested that TTLL6
may be involved in ciliary function in several organisms including zebrafish 5,24,29. We
therefore examined the effects of ttll6 deficiency using a ttll6 translational blocking MO 30
in zebrafish and observed ciliopathy-related morphological phenotypes, similar to although
less severe than what we observed following cep41 knockdown (Fig. 4a). Additionally, ttll6
morphants showed A-tubule axonemal defects, similar to those of cep41 morphants (Fig.
4a), besides diverse axonemal structural defects, which were consistent with a previous
report 30 (not shown). These data prompted us to test a possible functional relationship
between CEP41 and TTLL6 by pair-wise co-immunoprecipitation (coIP), and found that the
two proteins were part of a complex (Fig. 4b). Given evidence suggesting a candidate
regulator associated with the transport of a polyglutamylase between the basal body and the
cilium 29, we investigated a possible role for CEP41 in TTLL6 localization. Following
efficient knockdown of Cep41 in IMCD cells using siRNA (Supplementary Fig. 14), we
found localization of TTLL6 restricted mainly to the basal bodies, suggesting a block of
entry of TTLL6 into the cilium (Fig. 4c). These data suggest that CEP41 functions in tubulin
glutamylation by mediating transport of TTLL6 between the basal body and cilium.
Our finding of CEP41 mutations in JBTS patients provides the first evidence directly linking
defective tubulin glutamylation at the cilium to a cause of ciliopathies. Consistent with
previous studies implicating tubulin PTMs in cilia function 23,24,31,32, our data suggest
CEP41-mediated ciliary glutamylation is essential for axonemal formation. Moreover, we
found that CEP41 is required for transport of TTLL6 to modulate tubulin glutamylation,
although it remains to be determined how these molecules co-function. Most likely, these
proteins enter through the ciliary diffusion barrier at the transition zone 33,34 and are then
transported along ciliary microtubules by intraflagellar transport (IFT) motor proteins. Thus
examining whether CEP41 and TTLL6 form a complex with other factors at these locations
will follow as a future study. Because tubulin glycylation is partially affected in cep41
morphants and the phenotype of cep41 morphants is more severe than ttll6 morphants, it is
likely that CEP41 modulates other TTLL protein family members 5,30. Evidence for this
possibility has been described in recent studies suggesting involvement of both tubulin-
glycylation and glutamylation, each regulated by different TTLL-family members, in
maintaining ciliary structure and motility 30,35–37.
METHODS
Research subjects
MTI-429 family and additional families were recruited worldwide based upon the presence
of at least one individual with a neuroradiographically proven ‘molar tooth sign’ associated
with any JBTS or related disorder (JSRD) phenotype. Whenever possible, patients
underwent a full diagnostic protocol as previously reported 38 and a standardized clinical
questionnaire was obtained to assess extent of multi-organ involvement. We used standard
methods to isolate genomic DNA from peripheral blood of the affected and unaffected
family members after obtaining informed consent from all participating. Human subject
research was approved by the Ethics Boards of Leeds (East), CASA Sollievo della
Sofferenza Hospital/CSS-Mendel Institute, Hôpital Necker-Enfants Malades, Human
subjects divisions at the University of Washington, University of Michigan Institutional
Review Board and Human Research Protection Program, University of California, San
Diego.
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Genome-wide screen and fine mapping
A 5K whole genome linkage SNP-scan was performed with family MTI-429 using the
Illumina Linkage IVb mapping panel 39, and analyzed with easyLinkage-Plus software 40,
which runs Allegro version 1.2c in a PC Windows interface to calculate multipoint LOD
scores. Parameters were set to autosomal recessive with full penetrance, and disease allele
frequency of 0.001. Fine mapping on the pedigree was performed with the Affymetrix 250K
Nsp1 SNP array, and data were searched for common shared homozygous intervals from all
affected family members using a custom script implemented in Mathematica (B. Merriman,
unpublished). This script identifies all homozygous intervals longer than 2 Mbp for which
there are no more than 1 % heterozygous calls (to permit potential genotyping errors).
Mutation screening
Mutational screening of CEP41 was performed by direct sequencing of the 11 coding exons
and the adjacent intronic junctions in patients. PCR products obtained were treated with
Exonuclease I (EXO) (Fermentas) and shrimp alkaline phosphatase (SAP) (Promega), and
both strands were sequenced using a BigDye terminator cycle sequencing kit with an
ABI3100 automated sequencer (Applied Biosystems). The primers and optimized PCR
conditions used are depicted (Supplementary Table 3). Segregation of the identified
mutations was investigated in all available family members. All identified mutations in
CEP41 were not encountered in 96 ethnically matched controls (188 chromosomes) upon
direct sequencing.
Bioinformatics
Genetic location is according to the March 2006 Human Genome Browser build hg18. The
ciliary proteome was searched using web-based tools 41,42. Protein sequence conservation
was determined using ClustalW multiple amino acid sequence alignment (see URL section).
Cloning
Full-length human CEP41 was cloned into the TOPO blunt vector, and then shuttled into
EGFP- and HA- containing vectors. Human and zebrafish cep41 open reading frame were
amplified by RT-PCR and cloned into the pCS2+ vector in order to synthesize RNA for
injection into zebrafish embryos. Mouse Cep41 open reading frame was amplified and
cloned into the GST-, EGFP-, and FLAG-conjugated vectors for biochemical assays.
Generation of Cep41 mutant mice
SIGTR ES cell line (AW0157), derived from 129P2/OlaHsd mice carrying a gene trap
insertion 43 in the intron 1 of Cep41, was obtained from the European Conditional Mouse
Mutagenesis Program (EUCOMM) and injected into C57BL/6 recipient blastocysts at the
Mouse Biology Program, UC Davis. High-percentage chimeras (≥ 70 %) were bred to
C57BL/6 for germline transmission. Gene-trapped mice were isolated by PCR for β-
galactosidase-neomycin fusion gene and genotyped by Southern blot analysis hybridizing a
~10 kb (wild-type allele) and a ~8 kb (gene-trapped allele) product (Supplementary Fig. 5).
Zebrafish experiments
AB wild type zebrafish strains were used for in situ hybridization carried out by standard
protocols using a DIG-labeled sense and anti-sense RNA probe for cep41. To knockdown
zebrafish cep41, a translational blocking morpholino antisense oligonucleotide (MO, Gene
Tools Inc) was used: 5′-CATCTTCCAGCAGCAGAGCTTCGGC-3′, diluted to appropriate
concentrations in deionized sterile water, and injected into one-two cell stage embryos,
obtained from natural spawning of zebrafish lines. To rescue the phenotypes in MO-injected
embryos (morphants), RNA transcribed in vitro with the SP6 mMessage mMachine kit
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(Ambion) was co-injected. For characterization of ciliary defects in zebrafish embryos, the
morphological phenotype of either cep41 or ttll6 morphants were observed until 5 days post-
fertilization (dpf) and quantified under bright-field microscopy based upon previously
established criteria 44. For western blot analysis at 1–2 dpf zebrafish control embryos and
cep41 morphants, about 50 embryos with each genotype were deyolked and the embryo
lysates were extracted with RIPA buffer. For immunostaining with GT335, Ac-Tub and
PolyG Abs in whole-mount zebrafish embryos, 3 dpf control embryos and cep41 morphants
were fixed in Dent’s fixative (80 % MeOH: 20 % DMSO) at 4 °C overnight and incubated
with GT335 Ab (1:400), Ac-Tub (1:400) and PolyG (1:300) as primary and with anti-goat-
mouse 594 (1:1000) as secondary in diluted blocking solution (10 % normal goat serum: 0.5
% Tween 20 in PBS).
Cell culture and transfection
IMCD3, hTERT-RPE1, human fibroblast, and human embryonic kidney (HEK293) cells
were grown in appropriate DMEM or MEM media supplemented with 10–20 % fetal bovine
serum (FBS) at 37 °C in 5 % CO2. Healthy human female and male control fibroblast cells
were obtained from ATCC, and patient fibroblasts from skin biopsies were propagated in
culture (≤ 5 passage number). Human fibroblast cells were transfected using the Basic
Nucleofector Kit for Primary Mammalian Fibroblasts (Lonza). Other cells were transfected
at 60–80 % confluency with plasmids or siRNAs using Lipofectamine 2000 (Invitrogen).
The transfected cells were incubated for 24–72 hr with FBS or without FBS, dependent on
the experimental purpose.
Fluorescence microscopy and transmission electron microscopy (TEM)
Images of immunofluorescent stained cells were obtained on a Deltavision RT
Deconvolution microscope (Olympus IX70), under the same parameters for each
experiment. Taken images were edited and analyzed using Adobe Photoshop CS. For
electron microscopy, a standard protocol 45 was used except for one modification that tannic
acid was included in the fixative to enhance the final contrast of the images. Formvar-coated
slot grids (Electron microscopy Sciences) were used for sections (60–70 nm) to maximize
visibility of the tissue and cross section performed to observe cilia axonemal structure.
Live imaging of zebrafish embryos
Zebrafish embryos (12 hpf for cilia in the KV and 2.5 dpf for the renal cilia) were
transferred with embryo media to glass bottom culture dishes (MatTek), and 2.5 dpf
embryos were anesthetized in tricaine solution (~0.016 mg/ml). Images were acquired for 30
sec – 2 min using a Perkin Elmer UltraView Vox Spinning Disk Confocal with EMCCD
Hamamatsu 14 bit 1K × 1K camera, and edited with Volocity imaging software (Perkin
Elmer).
Immunofluorescence and biochemical assay
For immunofluorescence, cells were fixed in 100 % methanol at −20 °C for 10 min. Primary
antibodies used for immunofluorescence are: rabbit anti-CEP41, raised in rabbit to a purified
bacterially expressed protein of GST-fused CEP41, (PRF&L, PA), mouse anti-acetylated-
tubulin (Sigma), mouse GT335 Ab (gift from C. Janke), rabbit PolyE (gift from M.
Gorovsky) and rabbit anti-ARL13B (gift from T. Caspary). We used Alexa 488- or Alexa
594-conjugated secondary antibodies (Molecular Probes) and Hoechst 33342 nuclear dye.
All antibodies were diluted in 4 % normal donkey serum in PBS, primary antibodies were
incubated for overnight at 4 °C and secondary antibodies were applied for 1 hr at room
temperature. For immunoblotting, cells were extracted with RIPA lysis buffer 3 days after
transfection and boiled with SDS sample buffer. Same concentrations of lysates were loaded
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for paired experiments. Primary antibodies used for western blotting were: rabbit anti-GFP
(Roche), mouse anti-Flag (Sigma), and mouse anti-alpha-Tubulin (Sigma). Bound
antibodies were detected using horseradish peroxidase-conjugated secondary antibodies
(Pierce). For co-immunoprecipitation (co-IP), whole cell extracts (WCE) were prepared
from confluent HEK293 cells transiently transfected with 14 μg plasmid DNA in 10 cm
tissue culture dishes. WCE supernatants were processed for IP experiments by using 2 μg
primary antibodies and protein A/G mixed agarose beads (Pierce).
Statistical analysis
The χ2 staticstic was computed manually with P value assigned for 1 degree of freedom in
the characterization of mouse embryonic phenotype (Supplementary Table 2). For other
studies, Student’s two-tailed non-paired t-tests were carried out to determine the statistical
significance of differences between samples. P < 0.05 was considered statistically significant
for all tests.
URLs
Human Genome Browser, http://www.genome.ucsc.edu; ClustalW,
http://www.ebi.ac.uk/Tools/msa/clustalw2/.
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
We thank the Marshfield Clinic Research Foundation, Center for Inherited Disease Research (supported by the US
National Institutes of Health and National Heart, Lung, and Blood Institute) for genotyping support. We thank the
International JSRD Study Group, E. Bertini, and the French Society of Foetal Pathology for patient referrals, J.
Meerloo at the UCSD Microscopy Core (P30NS047101), T. Meerloo, Y. Jones and M. Farquhar and the CMM
Electron Microscopy Core Facility at UCSD, S. Wirth and B. Willis for mutant mouse generation, C. Janke at the
Institute Curie Research Center for GT335 antibody, TTLLs plasmids and technical advice, I. Drummond and N.
Pathak at the Massachusetts General Hospital for ttll6 MO, M. Gorovsky at the Univ. of Rochester for polyE and
polyG antibodies, S. Audollent at the Hôpital Necker-Enfants Malades for technical help, B. Sotak, N. Akizu, A.
Crawford, V. Cantagrel, and E-J. Choi for stimulating scientific discussion and comments. This work was
supported by the US National Institutes of Health R01NS048453 and R01NS052455 (J.G.G.), R01DK068306
(F.H.), R01NS064077 (D.A.D.) the American Heart Association 09POST2250641 (J.E.L.), the Italian Ministry of
Health (Ricerca Finalizzata Malattie Rare and Ricerca Corrente 2011), the Telethon Foundation Italy GGP08145,
and the Pierfranco and Luisa Mariani Foundation (E.M.V.), Grants-in-Aid from MEXT and JSPS, #23117517 and
#23570209 (K.I.), the BDF Newlife, the Medical Research Council (G0700073) and the Sir Jules Thorn Charitable
Trust 09/JTA (C.A.J.), l’Agence National pour la Recherche ANR 07-MRAR-Fetalciliopathies (T.A.-B.), Simons
Foundation Autism Research Initiative (J.G.G.) and the Howard Hughes Medical Institute (F.H. and J.G.G.).
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Figure 1.
Identification of mutations in CEP41 in affected individuals linked to the JBTS15 locus. (a)
Pedigree MTI-429 shows double first cousin marriage with five affected offspring. (b) Axial
brain MRI images from patients with CEP41 mutations in each MTI-429, MTI-1491 (T1-
weighted) and COR-98 (T2-weighted) family, showing the ‘molar tooth sign’ (red arrows).
(c) JBTS15 is 5.5 Mbp located at Chr. 7q31.33–32.3 (red box) defined by rs766240 and
rs4728251. Further narrowing of the interval to 2.8 Mbp based upon denser SNP scan to
encompass rs17165226 and rs2971773 (black arrows). (d) CEP41 genomic organization,
depicting locations of identified base changes including homozygous (red) splice mutations
and heterozygous (blue) missense or nonsense mutations. Capital letters: exon sequences,
small letters: intron sequences, asterisks: point mutations, underline: deletion. (e) RT-PCR
confirmation of the splicing defect of CEP41 in MTI-429 patient fibroblasts. Both CEP41
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mutant cells failed to produce CEP41 mRNA, compared with WT. GAPDH is control.
Green arrows: positions of primers, asterisk: region of splice mutation in MTI-429.
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Figure 2.
CEP41 is expressed in ciliated tissues and its loss recapitulates ciliopathy-related phenotypes
in zebrafish and mouse. (a) Zebrafish cep41 mRNA is expressed ubiquitously at gastrulation
stages (6 hpf), but at later stages, it is specifically expressed in ciliated organs: Kupffer’s
vesicle (red box), inner ear (white boxes), brain (brackets), eyes (arrows), pronephric duct
(black box), and heart (asterisks). A, anterior; D, dorsal; P, posterior; V, ventral. (b) CEP41
is predominantly localized to the basal body (arrowheads) and primary cilium (arrows) in
ciliated IMCD3 and hTERT-RPE1 cells. Insets: co-localization of CEP41 with GT335, a
marker of basal bodies and cilia. Scale bar 5 μm. (c) Knockdown of cep41 by injection of
morpholino oligonucleotide (MO) causes heart asymmetry defects in Tg (myl7:egfp)
zebrafish embryos. The cep41 morphants show either loss of asymmetry (midline) or
inversion of V/A asymmetry (reversed) at 72hpf. *P < 0.01, **P < 0.001. Error bars: s.e.m.
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A, atrium; L, left; R, right; V, ventricle. (d) Murine Cep41 gene-trap shows altered
embryonic morphogenesis. Range of phenotypes of Cep41Gt/Gt embryos includes mild
malformed hindbrain (arrowheads), exencephaly (brackets), hemorrhage in the head
(asterisk), dilated pericardial sac (arrows), failure to rotate and lethality at E10–12. (e)
Injection of human CEP41 RNA into cep41 morphants rescued ciliary phenotypes of
pericardial edema (arrowhead), hydrocephalus (asterisk) and curved tail (arrow), completely
or partially.
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Figure 3.
CEP41 is required for tubulin glutamylation at the ciliary axoneme. (a) Absent CEP41
protein in CEP41 mutant patient cells. (b) Ciliary localization of endogenous CEP41 in
human fibroblasts and loss of the protein in CEP41 mutant cilia. Scale bar 5 μm. (c) WT
cells have both GT335-positive basal bodies (arrowheads) and cilia (arrows, marked by
ARL13B), while mutant cells show staining of GT335 only at the basal bodies. Scale bar 5
μm. (d) Depletion of cep41 causes glutamylation defects in zebrafish olfactory placode cilia
(red arrows). Images of GT335-stainined WT and cep41 morphant embryos were taken in
anterior view (head is up and tail is down, D, dorsal; V, ventral), quantified below. (e)
Exogenous CEP41 expression restores ciliary axoneme glutamylation in CEP41 mutant
cells. Arrows: primary cilia stained for CEP41 and GT335. Insets: merged images at higher
power, quantified below. Scale bar 5 μm. **P < 0.001; error bars = s.e.m. (f) Ultrastructural
analysis of the pronephric ciliary axoneme at 72 hpf zebrafish embryos. Compared to WT,
cep41 morphants have A-tubule specific defects in the outer doublet microtubules. Arrows:
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