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© 2016. Published by The Company of Biologists Ltd.
Gene-targeted CEP164-deficient cells show a ciliation defect with intact
DNA repair capacity
Owen M. Daly1, David Gaboriau1,†, Kadin Karakaya2, Sinéad King1, Tiago J. Dantas1,¶,
Pierce Lalor3, Peter Dockery3, Alwin Krämer2 and Ciaran G. Morrison1,*
1Centre for Chromosome Biology, School of Natural Sciences and 3Anatomy, School of
Medicine, National University of Ireland Galway, Galway, Ireland; 2Clinical Cooperation Unit
Molecular Hematology/Oncology, German Cancer Research Center (DKFZ) and
Department of Internal Medicine V, University of Heidelberg, Im Neuenheimer Feld 280,
69120, Heidelberg, Germany
Summary statement: Knockout of the CEP164 ciliopathy gene ablates ciliogenesis but
causes no increase in sensitivity to DNA damage induced by ionising or ultraviolet
irradiation.
Keywords: Primary cilium; Centrosome amplification; DNA damage response; DNA repair;
CEP164; ciliopathy
†Current address: Facility for Imaging by Light Microscopy, Sir Alexander Fleming
Building, Imperial College London, UK.
¶Current address: Department of Pathology and Cell Biology, Columbia University,
New York, USA
*Correspondence to: Ciaran.Morrison@nuigalway.ie
JCS Advance Online Article. Posted on 10 March 2016
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Abstract
Primary cilia are microtubule structures that extend from the distal end of the mature,
mother centriole. CEP164 is a component of the distal appendages carried by the
mother centriole that is required for primary cilium formation. Recent data have
implicated CEP164 as a ciliopathy gene and suggest that CEP164 plays some roles
in the DNA damage response (DDR). We used reverse genetics to test the role of
CEP164 in the DDR. We found that conditional depletion of CEP164 in chicken
DT40 cells using an auxin-inducible degron led to no increase in sensitivity to DNA
damage induced by ionising or ultraviolet irradiation. Disruption of CEP164 in human
retinal pigmented epithelial cells blocked primary cilium formation but did not affect
cellular proliferation or cellular responses to ionising or ultraviolet irradiation.
Furthermore, we observed no localisation of CEP164 to the nucleus using
immunofluorescence microscopy and analysis of multiple tagged forms of CEP164.
Our data suggest that CEP164 is not required in the DDR.
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Introduction
Primary cilia are membrane-enclosed, microtubule-based organelles that extend like
antennae from the surface of most mammalian cell types to sense and transduce
various extracellular signals. They arise from the basal body, a template provided
when the mature, mother centriole docks to the plasma membrane (Goetz and
Anderson, 2010). Centrioles display structural polarity, with the proximal ends
containing microtubule triplets, which taper to doublets at the distal ends. The distal
ends of mature centrioles carry two sets of appendages, which anchor cytoplasmic
microtubules and which allow the docking of the mother centriole to the cell
membrane during the formation of the primary cilium (Goetz and Anderson, 2010).
The cilium core, the axoneme, consists of 9 microtubule doublets that extend from
the basal body.
In mammalian cells, cilium formation is closely regulated and linked to the cell cycle,
as cilia must be resorbed to allow the basal body to act as a centrosome and to
organise the mitotic spindle. Cellular quiescence, a temporary exit from the cell cycle
that can be induced by the removal of growth factors, facilitates ciliogenesis
(Kobayashi and Dynlacht, 2011). Current models associate primary cilia with cell
cycle exit and reduced proliferation, although the underlying mechanisms of such a
link are not well defined (Goto et al., 2013).
CEP164 encodes a centriolar appendage protein that is required for ciliogenesis
(Graser et al., 2007, Schmidt et al., 2012). It has also been implicated in modulating
the DNA damage response (DDR), particularly CHK1 (Sivasubramaniam et al., 2008).
CEP164 was initially identified in a proteomic analysis of the centrosome and later,
as a component of the distal appendages whose depletion by siRNA treatment
caused a marked reduction in primary cilium formation (Andersen et al., 2003,
Graser et al., 2007, Schmidt et al., 2012). Immuno- electron microscopy (EM)
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demonstrated the localisation of CEP164 to the distal end of the mother centriole
(Graser et al., 2007). Dual PALM/STORM imaging localised CEP164 in a ring
around the centriole barrel with a periodic enrichment of the signal within the ring
(Sillibourne et al., 2011) and stimulated emission depletion microscopy found that the
enriched CEP164 signal corresponds to nine symmetrically-arranged clusters around
the centriole, indicative of its association with each of the nine distal appendages
(Lau et al., 2012).
Recent data have indicated CEP164 mutations in nephronophthisis-related ciliopathy
(as NPHP15), a rare recessive degenerative disease of the kidney, retina and brain,
suggesting a link between ciliopathy and a DDR role of CEP164 (Chaki et al., 2012).
We set out to explore the mechanisms that link ciliary dysfunction with DDR defects,
using gene targeting to ablate CEP164 function.
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Results and Discussion
To analyse the roles of CEP164 in DNA repair, we used gene targeting in chicken
DT40 cells to insert a tag that combined GFP with an auxin-inducible degron (AID;
(Nishimura et al., 2009)) into the CEP164 locus of cells that stably expressed the
TIR1 E3 ligase component (Supplementary Figure 1A, B). As shown in Figure 1A,
AID-GFP-tagged CEP164 localised to the centrosome, although we observed no
localisation of CEP164 to the nucleus, even after UV irradiation of cells to levels that
induced robust formation of gamma-H2AX foci (Figure 1B). Upon addition of auxin,
AID-GFP-tagged CEP164 was depleted within 1 h (Figure 1C; Supplementary Figure
1C, D). CEP164-deficient cells showed doubling times of 8.3 h (clone 1) and 8.3 h
(clone 2), compared with control times of 8.4 h and 8.4 h for each clone, respectively,
and 8.3 h for wild-type cells. We observed no difference in sensitivity to ionising
radiation (IR) or UV treatment between CEP164-deficient and wild-type cells (Figure
1D, E). In keeping with this observation, IR-induced centrosome amplification, a
potential readout for the DDR (Bourke et al., 2007), occurred to the same levels in
both CEP164-deficient and wild-type cells (Supplementary Figure 1E). These data
show that CEP164 plays a limited role, if any, in nuclear responses to IR or UV-
induced DNA damage in DT40 cells.
Next, we cloned human CEP164 and expressed N- and C-terminally GFP- and
FLAG-tagged versions in human cell lines. As shown in Figure 2A-2D, we
consistently observed a centrosomal localisation for recombinant, overexpressed
CEP164, but saw no nuclear signal. Immunofluorescence microscopy with
previously-published anti-CEP164 antibodies also detected centrosomal, but not
nuclear signals (Figure 2E, 2F). Next, we generated a monoclonal antibody to
CEP164. As shown in Supplementary Figure 2A, monoclonal antibody 1F3G10
generated against amino acids 6-296 of CEP164 recognised a protein of
approximately 200 kDa in 3 human cell lines. We next confirmed 1F3G10 specificity
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by siRNA-depleting CEP164 from RPE1 cells. After CEP164 depletion, the 1F3G10
signal disappeared, with no effect on a GAPDH control (Supplementary Figure 2B).
In immunofluorescence microscopy experiments, 1F3G10 detected a signal that
partly co-localised with ninein, a component of the subdistal appendages, but
localised adjacent to CEP135, a centriole proximal end component (Supplementary
Figure 2C), consistent with the known localisation of CEP164 at the distal
appendages (Graser et al., 2007). While we conclude from these experiments that
the 1F3G10 monoclonal antibody is specific for CEP164, there was no signal seen in
the nucleus of U2OS or RPE1 cells.
Previous data have indicated a role for CEP164 in primary ciliogenesis (Cajanek and
Nigg, 2014, Graser et al., 2007, Schmidt et al., 2012, Chaki et al., 2012). Despite the
feasibility of inducing ciliogenesis in DT40 cells (Prosser and Morrison, 2015), we
preferred to examine the roles of CEP164 in a cell line with high levels of primary
ciliation. Thus, we used CRISPR-Cas9 technology to disrupt CEP164 in hTERT-
RPE1 cells, which show high levels of primary cilium formation upon serum
starvation. We used a guide RNA designed to direct DNA double-strand breaks in
exon 9 (the 7th coding exon) of the human CEP164 locus and selected clones that
had lost CEP164 expression by immunoblot analysis (Figure 3A). Sequence
analysis demonstrated that CEP164-deficient clones had incurred mutations in the
CEP164 locus that led to premature stop codons being transcribed in-frame with the
gene (Supplementary Figure 3A). Immunofluorescence microscopy confirmed that
these clones no longer expressed CEP164, although they still carried intact
centrioles (Figure 3B). These cells proliferated as rapidly as wild-type cells, with
doubling times of 24.1 h (clone 1) and 23.6 h (clone 2), compared with 23.5 h for
wild-type cells. We saw no alteration in cell cycle distribution in the absence of
CEP164 (Supplementary Figure 3B). Strikingly, CEP164-deficient cells showed a
complete absence of primary ciliation capacity that was rescued by transgenic
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expression of CEP164 (Figure 3C, 3D). Transmission EM analysis of the CEP164
null cells revealed no obvious structural defects in centriole structures, based on the
dimensions of the centriole barrels (Supplementary Figure 3C). 16 vesicles were
identified in proximity to the centrioles in 7 CEP164 null cells, but no docking was
observed, consistent with a defect at the vesicle docking stage in cilium formation
seen in siRNA knockdown experiments (Schmidt et al., 2012) (Figure 3E). Thus, our
disruption of the CEP164 locus confirms the findings made on CEP164’s roles in
primary ciliogenesis with siRNA experiments (Cajanek and Nigg, 2014, Graser et al.,
2007, Schmidt et al., 2012).
We next tested whether CEP164 deficiency impacted on cells’ ability to withstand
UV-induced DNA damage. A clonogenic survival assay showed that CEP164-
deficient RPE1 cells were no more sensitive than wild-type cells (Figure 4A). In a
positive control experiment, CETN2 null RPE1 cells (Prosser and Morrison, 2015)
showed an increased UV sensitivity, as had centrin-deficient chicken DT40 cells
(Dantas et al., 2011). Furthermore, we observed no localisation of CEP164 to
nuclear DNA damage foci of γ-H2AX after IR or UV treatment in RPE1 or HeLa cells
with either of two antibodies in immunofluorescence experiments (Figure 4B, C).
Taken together, these data indicate no defect in the response to DNA damage in
CEP164-deficient hTERT-RPE1 cells.
The results in the two models we have explored do not support a role for CEP164 in
the DDR. We did not see a proliferative decline, such as that described in IMCD3
cells after siRNA knockdown of CEP164 (Chaki et al., 2012) or an acceleration of cell
cycle progression, as has been described after siRNA knockdown of CEP164 in
RPE-FUCCI cells (Slaats et al., 2014). In our null lines, we observed no elevated
sensitivity to IR or UV irradiation in the absence of CEP164, which contrasts with the
phenotypes of UV sensitivity and loss of the G2-to-M checkpoint reported with siRNA
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knockdown of CEP164 in HeLa cells (Sivasubramaniam et al., 2008, Pan and Lee,
2009). These discrepancies have potential implications for understanding how
CEP164 mutations cause disease.
There are clear technical differences in the approaches that we have used and those
previously reported. An obvious possibility is that the gradual or partial depletion
imposed by siRNA treatment may lead to cellular responses different to those seen
with the loss of a protein, although our degron-mediated experiment might have been
expected to address this. Another possibility is that off-target effects of the siRNA
treatments resulted in more marked phenotypes. While the proliferative decline and
cell cycle defects in IMCD3 cells were rescued by transgenic expression of human
CEP164 (Chaki et al., 2012, Slaats et al., 2014), it is worth noting that rescues for the
UV sensitivity and checkpoint defects seen in CEP164 knockdown cells were not
performed (Sivasubramaniam et al., 2008, Pan and Lee, 2009), so that the specificity
of these RNAi phenotypes cannot be assessed.
We have not seen any significant nuclear localisation of CEP164 during the normal
cell cycle or after DNA damage in 1 chicken and 3 human cell lines, using 3 different
antibodies and multiple, differently-tagged versions of transgenically-expressed
CEP164. Similarly to published results (Graser et al., 2007, Schmidt et al., 2012),
our experiments have detected only cytosolic or centrosomal signals, in contrast to
the predominantly nuclear signals reported with those antibodies generated in the
original study that implicated CEP164 in the DDR (Sivasubramaniam et al., 2008).
Tagging experiments and several antibodies used in a recently-published study
showed predominantly cytosolic or centrosomal CEP164 signals, although these
authors also observed nuclear signals using the original CEP164 antibodies (Chaki
et al., 2012). Controls for the specificity of the nuclear immunofluorescence signals
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seen with these reagents after CEP164 knockdown or depletion have not been
detailed (Sivasubramaniam et al., 2008, Pan and Lee, 2009).
We have performed our DNA damage sensitivity and localisation analyses in cell
lines from different tissues. Thus, while we cannot exclude the possibility that
CEP164 contributes to the DDR in certain cell types, this does not appear to be a
general activity of the protein. Our data, which support a marked defect in primary
cilium formation, but normal levels of DNA repair capacity in the absence of CEP164,
suggest that the principal cellular defect associated with CEP164 deficiency is the
inability to undertake primary ciliogenesis.
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Materials and Methods
Cell culture
Chicken DT40 cells were cultured as previously described (Takata et al., 1998).
hTERT-RPE1 cells were cultured as previously described (Prosser and Morrison,
2015). HeLa and U2OS cells were obtained from ATCC and cultured in DMEM
(Lonza or PAA/ GE Healthcare), supplemented with 10% FCS (Lonza or Biochrom).
Jurkat cells were from the European Collection of Animal Cell Cultures and were
grown in RPMI with 10% FCS (Lonza). Auxin (Sigma-Aldrich) was prepared at 0.5 M
in ethanol. IR treatments used a 137Cs source (Mainance Engineering). For UV-C
irradiation, cells were irradiated using an NU-6 254-nm UV-C lamp at 23 J/m2/min
(Benda). DT40 clonogenic survival assays were performed as previously described
(Takata et al., 1998), with 500M auxin added to the medium of cells 24 h prior to
irradiation where a degron-tagged protein was to be depleted, and retained in the
methylcellulose medium used for clonogenesis. For UV clonogenic survival assays
in hTERT-RPE1, cells were counted before being serially diluted and plated in 10 cm
dishes. The cells in each dish were allowed to adhere for 6 h before the medium
was siphoned off and they were irradiated. Conditioned medium (filtered media taken
from 50% confluent cells) was used to replenish the dishes before incubation.
Cloning
For targeting the chicken CEP164 locus, 5′ and 3’ homology arms and probe
sequence were amplified from DT40 genomic DNA with KOD polymerase
(Novagen/Merck) using the following primers:
5’ arm: 5’-gacgtcCAGACAACAAGCTAGGATATGTACCT-3’ and 5’-
ccgcggGTACCGGTACACTTTAATTTGTCTGT-3’
3’ arm: 5’-agatctAAGGTGGGACTTGGTGTTTTCAGCC-3’ and 5’-
cctaggTTTGGGTTTCAGTGCCATCCCGTG-3’
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5’ probe: 5’-CTTCTGATTTCAGTCCTGCGTGTT-3’ and 5’-
CAGACATTAAATACAAGTCCCCTCC-3’
The probe for Southern analysis was labeled with digoxigenin using the PCR DIG
Probe Synthesis Kit (Roche). AID-encoding sequence (Eykelenboom et al., 2013)
was subcloned into pEGFP-N1 (BD Biosciences/ Clontech) and a TIR1-9myc
plasmid (pJE108 (Eykelenboom et al., 2013)) was stably cloned into DT40 cells to
control the degron.
For cloning human CEP164 cDNA, hTERT-RPE1 RNA was extracted using TRI
reagent (Invitrogen). Reverse transcription was performed using SuperScript First-
Strand (Invitrogen) and PCR with KOD Hot Start. cDNAs were cloned into pGEM-T
Easy (Promega), sequenced and then subcloned into pEGFP-N1, pEGFP-C1 (BD
Biosciences/ Clontech) or pCMV8 Tag 4A (Agilent Technologies, Santa Clara, CA,
USA). The primers used to amplify human CEP164 cDNA (isoform 1,
NP_055771.4 ) were as follows: 5′-aagcttATGGCTGGACGACCCCTCCGCA-3’ and
5’-gtcgacCAGAAGCGATACACCYYCACTC-3’. Isoform 2 (UniProtKB - Q9UPV0
(CE164_HUMAN)) was cloned by mutating CEP164 cDNA isoform 1 using the
QuickChange Lightning Site-Directed Mutagenesis Kit (#210518, Agilent) with the
following primers:
Deletion of GGAG: 5'-AGCAGTCCAAAGGCCTGGAAGGTTATCTCCTC-3' and 5'-
GAGGAGATAACCTTCCAGGCCTTTGGACTGCT-3'; Deletion of
GTGAGTGGTGGCGGCAGCAGAGGATCGACTCAA: 5'-
CCCCGCCTCACCCCCCGAGTCTCA-3' and 5'-
TGAGACTCGGGGGGTGAGGCGGGG-3'; Insertion of GGAGAGGTACCAT: 5'-
AGCAGTCCAAAGGCCTGGAGGAGAGGTACCATAGGTTATCTCCTC-3' and 5'-
GAGGAGATAACCTATGGTACCTCTCCTCCAGGCCTTTGGACTGCT-3'; Insertion
of TCGACTCAA:
5'-CCCCGCCTCACCTCGACTCAACCCCGAGTCTCA-3'; 5'-
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TGAGACTCGGGGTTGAGTCGAGGTGAGGCGGGG-3'.
CRISPR/Cas9 targeting of CEP164 in hTERT-RPE1 cells
Primers targeting exon 9 (Mali et al., 2013) were cloned into pX330-U6-
Chimeric_BB-CBh-hSpCas9 (plasmid 43330; Addgene (Cong et al., 2013)): 5′-
CACCGCTGTTGGCAAAGGGCGACA-3′ and 5′-
AAACTGTCGCCCTTTGCCCACAGC-3′. Transfections used Lipofectamine 2000
(Invitrogen). Genomic PCR products obtained with the diagnostic primer pair, 5′-
CTGGGTGATTGATAACCATTGGG -3’ and 5′-CGCAAATGAAGCTCCTGACTCAGT
-3′ were cloned into pGEM-T-Easy and sequenced.
Monoclonal antibody generation
cDNA sequence encoding CEP164 amino acids 6-296 was cloned into pGEX-4T1
(GE Healthcare) and the bacterially-expressed GST fusion product was purified over
a glutathione column prior to thrombin cleavage. Purified CEP164 protein fragment
was used for hybridoma generation (Dundee Cell Products). Individual supernatants
were screened by immunoblot and microscopy and then concentrated antibody was
purified from the best-performing 1F3G10 supernatant (Proteogenix).
RNA-mediated interference
Cells were transfected with 50 nmol custom siRNA targeting CEP164 from Qiagen
CAGGUGACAUUUACUAUUUCA or Silencer Select siRNA targeting GAPDH
UGGUUUACAUGUUCCAAUATT using Oligofectamine (Invitrogen).
Immunofluorescence microscopy
Cells were fixed for analysis as previously described (Prosser and Morrison, 2015).
Donkey and goat secondary antibodies were labeled with Cy3, Alexa Fluor 488 or
Alexa Fluor 594 (Jackson ImmunoResearch or Molecular Probes). Rabbit polyclonal
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antibodies used were as follows: γ-tubulin (T3559, Sigma) γ-H2AX (Ab2893, Abcam),
CEP135 (1420 739 (Bird and Hyman, 2008)), CEP164 (HPA037606, Sigma),
CEP164 (R171 (Graser et al., 2007)), detyrosinated α-tubulin (Ab48389, Abcam) and
ninein (ab4447, Abcam). Mouse monoclonal antibodies used were γ-tubulin (GTU88,
Sigma or TU-30/11–465-C100, Exbio), γ-H2AX (JBW301, Upstate), CEP164
(1F3G10) and centrin (20H5, Millipore). Images of DT40 and hTERT-RPE1 cells
were captured on an IX71 microscope (Olympus) with a 100× oil objective, NA 1.35,
using Volocity software (PerkinElmer), and are presented as maximum intensity
projections of Z-stacks after deconvolution. Alternatively, images of U2OS cells were
captured and processed using an Axiovert 200 M microscope equipped with a Plan-
Apochromat 63x, NA 1.4 objective and AxioVision software (Carl Zeiss Microscopy)
and are presented as single sections.
Electron microscopy
hTERT-RPE1 cells were serum-starved in 0.1% FCS for 24h prior to harvest. Cell
pellets were prepared for transmission EM and imaged with an H-7000 Electron
Microscope (Hitachi) as described (Prosser and Morrison, 2015).
Immunoblotting
Whole-cell extracts were prepared using RIPA buffer (50mM Tris-HCl pH7.4, 1% NP-
40, 0.25% sodium deoxycholate, 150mM NaCl, 1mM EDTA and protease inhibitor
cocktail). Immunoblot analyses used the following primary antibodies: α-tubulin
(B512, Sigma), CEP164 (IF3G10), GFP (11814460001, Roche) and GAPDH (14C10,
Cell Signalling).
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Flow cytometry
Cells were fixed in 70% ice-cold ethanol overnight at 4°C, washed twice in PBS, and
incubated in 40 µg/ml propidium iodide and 200 µg/ml RNase A in PBS for 1 h.
Cytometry was performed on a FACSCanto (BD).
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Competing interests
No competing interests declared.
Author contributions
Project conception and direction: O.M.D., K.K., A.K., C.G.M. Data analysis: O.M.D.,
D.G., K.K., S.K., T.J.D, P.D., A.K., C.G.M. Experimental work: O.M.D., T.J.D, K.K.
(cell biology); D.G., S.K. (monoclonal antibody); P.L. (EM). Writing the paper: O.M.D.
and C.G.M.
Funding
We acknowledge the National Biophotonics and Imaging Platform Ireland and the
NCBES Flow Cytometry core facility, which were supported by Irish Government
Programme for Research in Third-Level Institutions cycles 4 and 5. This work was
funded by Science Foundation Ireland Principal Investigator award 10/IN.1/B2972
and European Commission SEC-2009-4.3-02, project 242361 ‘BOOSTER’.
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Figures
Figure 1 Wild-type DNA damage sensitivity after AID-mediated depletion of
CEP164
A. Centrosomal localisation of AID-GFP-CEP164 in chicken DT40s (green). Co-
staining was for γ-tubulin (red). DNA was labelled with DAPI (blue). Scale bar, 2 µm.
B. Absence of nuclear AID-GFP-tagged CEP164 signal. Cells were treated with 10
J/ m2 UV irradiation 1 h prior to fixation and staining for γ-H2AX (red) and DNA (blue).
Scale bar, 2 µm.
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C. Auxin-mediated depletion of AID-GFP-CEP164. Immunoblot shows total cell
extracts from cells of the indicated genotype before and 24 h after treatment with 500
µM auxin. α-tubulin was used as a loading control.
D., E. Clonogenic survival assay of cells of the indicated genotype after (D.) IR or
(E.) UV irradiation. Curves show mean + s.d. of three independent experiments.
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Figure 2 Centrosomal, but not nuclear, localisation of CEP164 in human cells
Localisation of the indicated transiently-transfected tagged CEP164 isoform (green)
in A. hTERT-RPE1 cells and B-D. U2OS cells. Co-staining was for CEP135 or γ-
tubulin (red). Iso., isoform.
E. Immunofluorescence localisation of CEP164 (green) in U2OS cells using rabbit
polyclonal antibodies (Graser et al., 2007). Co-staining was with antibodies for γ-
tubulin (red).
F. Immunofluorescence localisation of CEP164 (red) in hTERT-RPE1 cells using
rabbit polyclonal antibodies (Sigma). Co-staining was for γ-tubulin (green). DNA
was labelled with DAPI (blue). Scale bars, 5 µm.
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Figure 3 Absence of primary ciliation in CEP164 null hTERT-RPE1 cells
A. Immunoblot analysis of CRISPR-disrupted CEP164 null clones.
B. Absence of CEP164 signal in CEP164 null cells. Cells were stained with mouse
monoclonal 1F3G10 (m) or polyclonal rabbit (r) antibodies to CEP164 then co-
stained with antibodies to CEP135 or centrin2 and for DNA (blue). Scale bar, 5 µm.
C. Immunofluorescence microscopy analysis of primary cilia in cells of the indicated
genotype. After 72 h serum starvation, cells were fixed and stained for detyrosinated
tubulin (green) and centrin2 (red). DNA was visualised with DAPI (blue). Scale bar,
5 µm.
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D. Quantitation of primary ciliation frequency in wild-type, CEP164 null cells and
CEP164 nulls that were stably transfected with CEP164. Cilia were quantitated by
microscopy of detyrosinated tubulin and bar graph indicates the mean + s.d. of three
independent experiments in which at least 100 cells were counted. AS,
asynchronous; SS, serum-starved.
E. TEM analysis of ciliogenesis in cells of the indicated genotype. Panel 1 shows an
assembled primary cilium in wild-type cells; Panel 2 a docked ciliary vesicle; Panels
3 and 4 show mother centrioles in proximity to vesicles without any docking in serum-
starved CEP164-deficient cells. Scale bars, 500 nm.
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Figure 4 Intact DDR in CEP164 null hTERT-RPE1 cells
A. Clonogenic survival assay of cells of the indicated genotype after UV irradiation.
Curves show mean + s.d. of three independent experiments.
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B., C. Absence of nuclear CEP164 signals after DNA damage. hTERT-RPE1 (B.) or
HeLa (C.) cells were treated with 10 Gy IR or 20 J/m2 UV 1h prior to fixation and
staining for CEP164 (green), γ-H2AX (red) and DNA (blue). Scale bar, 5 µm.