The Rockefeller University Press $30.00
J. Cell Biol. Vol. 190 No. 5 927–940
Correspondence to George B. Witman: George.Witman@umassmed.edu
Abbreviations used in this paper: BAC, bacterial artificial chromosome; BBS,
Bardet-Biedl syndrome; DIC, differential interference contrast; IFT, intraflagellar
transport; LCA, Leber congenital amaurosis.
Mutations that disrupt the assembly, structure, and/or function of
cilia or flagella result in cilia-related disorders termed ciliopathies
(Badano et al., 2006; Fliegauf et al., 2007). Mutations in CEP290
cause ciliopathies that exhibit a range of severity (Helou et al.,
2007). CEP290 mutations are a common cause of Leber congeni-
tal amaurosis (LCA; den Hollander et al., 2006; Sundaresan et al.,
2009), in which blindness results from degeneration of the retina
but other organ systems are often unaffected. Other CEP290 mu-
tations cause Meckel syndrome (Baala et al., 2007; Frank et al.,
2008), a perinatal-lethal disease in which essentially all tissues
with cilia are affected. Of intermediate severity is Joubert syn-
drome, in which brain development is affected and which
often also presents with ocular and renal manifestations (Sayer
et al., 2006; Valente et al., 2006; Brancati et al., 2007).
The described subcellular localization of CEP290 (also
known as nephrocystin-6/NPHP6) is consistent with a role for
CEP290 in cilia. In mammalian cells, CEP290 has been localized
to centrosomes/basal bodies (Andersen et al., 2003; Chang et al.,
2006; Sayer et al., 2006; Valente et al., 2006; Tsang et al., 2008),
pericentriolar satellites (Kim et al., 2008), the connecting cilium
of photoreceptors (Chang et al., 2006; Sayer et al., 2006), and the
dendritic knobs of olfactory sensory neurons (McEwen et al.,
2007), and CEP290 was immunoprecipitated with centrosomal
components (Chang et al., 2006; McEwen et al., 2007; Tsang
et al., 2008). CEP290 has a nuclear localization signal and, in ad-
dition to its cytoplasmic localization, has been observed in the
nucleus (Guo et al., 2004; Sayer et al., 2006).
The exact role of CEP290 is unclear, but all available evi-
dence suggests that CEP290 is somehow involved in ciliary
function. Morpholino knockdown of CEP290 in zebrafish re-
sulted in retinal, renal, and cerebellar phenotypes that are indic-
ative of defects in cilia and are commonly seen in humans with
Joubert syndrome (Sayer et al., 2006). Defects in the localization
of several ciliary proteins were found in photoreceptors and
a Chlamydomonas reinhardtii mutant in which most of the
CEP290 gene is deleted. Immunoelectron microscopy in-
dicated that CEP290 is located in the flagellar transition
zone in close association with the prominent microtubule–
membrane links there. Ultrastructural analysis revealed
defects in these microtubule–membrane connectors,
resulting in loss of attachment of the flagellar membrane
to the transition zone microtubules. Biochemical analysis
utations in human CEP290 cause cilia-related
disorders that range in severity from isolated
blindness to perinatal lethality. Here, we describe
of isolated flagella revealed that the mutant flagella have
abnormal protein content, including abnormal levels
of intraflagellar transport proteins and proteins asso-
ciated with ciliopathies. Experiments with dikaryons showed
that CEP290 at the transition zone is dynamic and under-
goes rapid turnover. The results indicate that CEP290
is required to form microtubule–membrane linkers that
tether the flagellar membrane to the transition zone
microtubules, and is essential for controlling flagellar
CEP290 tethers flagellar transition zone
microtubules to the membrane and regulates
flagellar protein content
Branch Craige,1 Che-Chia Tsao,2 Dennis R. Diener,2 Yuqing Hou,1 Karl-Ferdinand Lechtreck,1 Joel L. Rosenbaum,2
and George B. Witman1
1Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA 01655
2Department of Molecular, Cellular and Developmental Biology, Yale University, New Haven, CT 06520
© 2010 Craige et al. This article is distributed under the terms of an Attribution–
Noncommercial–Share Alike–No Mirror Sites license for the first six months after the pub-
lication date (see http://www.rupress.org/terms). After six months it is available under a
Creative Commons License (Attribution–Noncommercial–Share Alike 3.0 Unported license,
as described at http://creativecommons.org/licenses/by-nc-sa/3.0/).
T H E J O U R N A L O F C E L L B I O L O G Y
JCB • VOLUME 190 • NUMBER 5 • 2010 928
The Y168 mutant was backcrossed twice to wild-type cells
(the F2 progeny with the deletion is henceforth referred to as
cep290), and each time the mutant phenotype cosegregated with
the deletion. The mutant phenotype was rescued by transforma-
tion with genomic DNA encoding either untagged or HA-tagged
CEP290 and no other gene (Fig. 1, A and B; and Videos 1–4),
which confirmed that the mutant phenotype is caused by the dele-
tion of CEP290. Antibodies generated against a peptide consist-
ing of the last 14 amino acids in the C terminus of C. reinhardtii
CEP290 recognized a single band of the predicted molecular mass
(275 kD) that was present in Western blots of whole-cell lysates
of wild-type and rescued cells, but absent in cep290 cells, which
confirmed that the antibody is specific and that CEP290 is absent
in the mutant (Fig. 1 E).
CEP290 is located in the transition zone
Immunofluorescence microscopy using the antibody directed
against CEP290 showed CEP290 immunoreactivity at the base
of each flagellum of wild-type cells (Fig. 2 A, a–c; and Fig. 2 B);
the label was missing from cep290 mutant cells (Fig. 2 A, d–f),
indicating that the labeling was specific. This localization pat-
tern was confirmed by using the CEP290-HA rescued strain
and antibodies to HA (see Fig. 6). Co-labeling with antibodies
to polyglutamylated tubulin, which label basal bodies and fla-
gellar axonemes but fail to label the transition zone of flagella
(Lechtreck and Geimer, 2000), revealed that the CEP290 signal
was located in the region that lacked polyglutamylated tubulin
labeling, indicating that CEP290 is located in the transition zone
(Fig. 2 C). CEP290 remained associated with the transition zone
after deflagellation (Fig. S1), a finding that is consistent with its
presence in the C. reinhardtii centriole proteome (which included
the transition zone; Keller et al., 2005) and its absence in the
C. reinhardtii flagellar proteome (which lacked the transition
zone; Pazour et al., 2005). CEP290 was undetectable in West-
ern blots of isolated flagella (Fig. S2), which confirmed that
CEP290 does not enter the flagellar compartment distal to the
cep290 mutant cells have defects in the
structures that bridge the transition zone
microtubules and membrane
To determine if loss of CEP290 resulted in ultrastructural defects,
we examined cep290 mutant cells by EM. We observed no defects
in basal bodies. In longitudinal sections through the transition
zone, the wedge-shaped structures that extend between the dou-
blet microtubules and membrane in wild-type cells (Ringo, 1967)
were missing or collapsed onto the doublets in the mutant cells
(Fig. 3, A and B). In cross sections through transition zones, the
Y-shaped connectors that extend from the transition zone doublet
microtubules to the flagellar membrane were often absent in the
mutant cells (Fig. 3, C and D). Image averaging (Fig. 3, C and D,
insets) revealed remnants of Y connectors on the mutant doublets,
including a small electron density possibly corresponding to the
base of the Y connector. In short flagella, we occasionally observed
bulges in the flagellar membrane filled with electron-dense material
(Fig. 3, E–G). In rare instances, we observed defects in axonemal
microtubules (Fig. 3 F); however, the vast majority of sections
olfactory sensory neurons of the rd16 mouse, which harbors
a hypomorphic in-frame deletion in Cep290 and displays early-
onset retinal degeneration and anosmia (Chang et al., 2006;
McEwen et al., 2007). In addition, RNAi knockdown of CEP290
in cultured mammalian cells decreased the percentage of cells
displaying primary cilia (Graser et al., 2007; Tsang et al., 2008).
CEP290 binds to and activates the transcription factor ATF4,
a protein implicated in renal cyst formation (Sayer et al., 2006).
Collectively, the data suggest that CEP290 is involved in ciliary
assembly and the expression, targeting, or transport of ciliary
proteins, but why CEP290 deficiency causes defects in cilia
To elucidate the molecular role of CEP290, we used
Chlamydomonas reinhardtii, which has numerous character-
istics that make it an ideal model organism for studying the
function of ciliary and basal body components. Human and
C. reinhardtii CEP290 are highly conserved (Basic Local
Alignment Search Tool [BLAST] E = 5 e27); Keller et al.
(2005) previously identified the C. reinhardtii CEP290 ortho-
logue by proteomic analysis of isolated centrioles. Our results
show that C. reinhardtii CEP290 is localized to the transition
zone between the flagellar basal body and axoneme, where it is
required for the assembly of the microtubule–membrane link-
ers characteristic of this poorly understood region. Loss of the
protein results in defects in flagellar composition, including alter-
ation of the normal balance of intraflagellar transport (IFT) com-
plexes A and B and abnormal levels of the membrane-associated
proteins BBS4 and PKD2, the human homologues of which are
involved in ciliopathies. The results also show that CEP290 is
highly dynamic at the transition zone, which suggests that it
may be involved in signaling between the cilium and cell body,
and which has clinical implications for CEP290 gene therapy in
the retina to treat LCA due to defects in CEP290. We propose
that CEP290 is part of a complex that links the membrane to
the microtubules in the transition zone and regulates entry of
proteins into the ciliary compartment.
Identification of a C. reinhardtii
To understand how CEP290 deficiency affects flagellar func-
tion, a library of C. reinhardtii insertional mutants was screened
using PCR with primers complementary to sequences in the
5 and 3 regions of the C. reinhardtii CEP290 homologue (pre-
viously designated POC3; Keller et al., 2005). In one strain
(Y168), primers directed toward the 3 end of CEP290 failed
to amplify a product, which indicated that this region is deleted
(Fig. 1 A). Further analysis by PCR revealed an 18.5-kb dele-
tion encompassing all but the first 2–4 exons (out of a total of
37 exons) of CEP290. The mutant cells were mostly palmel-
loid (i.e., failed to hatch from the mother cell wall after mitosis;
Fig. 1 B). Release from the mother cell wall by treatment with
autolysin enzyme (Harris, 2009) demonstrated that the cells have
very short/stumpy flagella, some of which begin to elongate over
time (Fig. 1 C). Occasionally, bulges were present in the short or
stumpy mutant flagella (Fig. 1 D).
929CEP290 function in the flagellar transition zone • Craige et al.
Figure 1. Identification of a C. reinhardtii CEP290 mutant. (A) Schematic of the region of C. reinhardtii chromosome 3 containing the CEP290 (POC3)
locus. CEP290 is flanked by the FAP61 gene and by gene models encoding predicted proteins nos.182839 and 144014 in Joint Genome Institute
version 4.0 of the Chlamydomonas genome (http://genome.jgi-psf.org/Chlre4/Chlre4.home.html). Arrows indicate the positions of PCR primer pairs; plus
and minus symbols indicate whether a PCR product was generated using genomic DNA of strain Y168 as template (all primer pairs amplified a product
when wild-type genomic DNA was used as template). Rescuing constructs consisted of either untagged or HA-tagged genomic DNA fragments containing
only the CEP290 gene. (B) 30-frame image averages of videos (acquired at 30 frames per second) depicting the mutant phenotype (cep290) and rescue
of the phenotype (cep290::CEP290 and cep290::CEP290-HA). Motile cells are seen as long meandering tracks, whereas nonmotile (palmelloid) cells
appear as bright foci (cep290). See Videos 1–4. Bar, 10 µm. (C) Palmelloid cep290 mutant cells were released from the mother cell wall by treatment
with autolysin, then fixed at the indicated time points after the addition of autolysin and processed for immunofluorescence using antibodies to acety-
lated tubulin. The mutant cells have stumpy flagella that partially elongate over time. Bar, 10 µm. (D) cep290 mutant cells were hatched with autolysin,
immobilized in 0.5% agar, and imaged at the indicated time points (minutes) after the addition of autolysin. The arrowhead marks a flagellum that began
to elongate then formed a bulge. The arrow marks a flagellum that failed to elongate and formed a bulge. Bar, 1 µm. (E) Western blot of whole cells.
A peptide antibody generated against the C-terminal 14 amino acids of C. reinhardtii CEP290 is specific and confirms the absence of CEP290 in the
mutant strain and expression of CEP290 in the rescued strains. F1-ATPase is a mitochondrial protein used as a loading control. The positions of standard
proteins and their molecular masses in kD are indicated.
JCB • VOLUME 190 • NUMBER 5 • 2010 930
In contrast, in the mutant, the transition zone membrane was lost
after detergent extraction (Fig. 3 J). The data indicate that loss of
CEP290 causes defects in the microtubule–membrane connec-
tions in the transition zone, which results in loss of attachment
of the flagellar membrane to the transition zone.
CEP290 localizes to the transition zone
Single- and double-label immunogold EM was used to localize
CEP290 more precisely within the transition zone. Using the anti-
body that recognizes the C-terminal 14 amino acids of CEP290,
gold particles exclusively labeled the transition zone of detergent-
extracted cytoskeletons of wild-type cells, with most labeling be-
tween the doublet microtubules and the membrane (Fig. 4, A–C;
through axonemes revealed normal outer doublets, radial spokes,
dynein arms, and central pair microtubules.
To determine if the disruption of the transition zone struc-
tures altered the normal relationship between the transition zone
microtubules and membrane, we measured the distance between
the transition zone cylinder and the membrane in wild-type and
mutant cells. In cep290 mutant cells, the distance was greater
and more variable (Fig. 3 H). To determine if this reflects a func-
tional detachment of the membrane from the microtubules, we
examined the association of the transition zone membrane with
the underlying microtubules in detergent-extracted cytoskel-
etons. In wild-type cells, the transition zone membrane remains
intact and associated with the transition zone microtubules
after detergent extraction (Fig. 3 I; Kamiya and Witman, 1984).
Figure 2. C. reinhardtii CEP290 is located in the transition
zone. (A) Wild-type and cep290 mutant cells were fixed and
processed for immunofluorescence using the indicated anti-
bodies. (B and C) Wild-type cells were detergent-extracted,
and the resulting cytoskeletons were processed for immuno-
fluorescence using the indicated antibodies. Co-labeling
with antibodies to CEP290 and acetylated tubulin revealed
CEP290 localization at the base of each flagellum (A and B).
(C) Co-labeling with polyglutamylated tubulin indicated that
CEP290 is located within the transition zone (arrowheads;
Lechtreck and Geimer, 2000). Also see Fig. S1. Bars, 1 µm.
931CEP290 function in the flagellar transition zone • Craige et al.
the Y connectors and the area immediately surrounding the Y
connectors (Figs. 4 C and S4). These experiments confirm that
CEP290 is located in the transition zone, and suggest that the
middle of CEP290 is closely associated with the microtubule–
membrane connectors in the center of that region, whereas the
C terminus of the protein is slightly more dispersed. These results
are consistent with our ultrastructural findings that these connec-
tors are missing or altered in the mutant.
cep290 mutant flagella have an abnormal
An important advantage of C. reinhardtii is that the flagella
can be isolated, so that the effect of loss of CEP290 on flagellar
protein content can be assessed biochemically. Flagella were
Fig. S3, and Fig. S4). In longitudinal sections, gold particles were
distributed throughout the length of the transition zone, with the
highest concentration in the proximal part of the transition zone.
In cross sections, most gold particles were located in the space
between the Y-shaped connectors, with a few over the connectors.
No labeling was observed with cytoskeletons of cep290 mutant
cells treated in an identical manner (Fig. S3, A–D). Using anti-
bodies to the HA epitope, which is located near the middle of
CEP290-HA (Fig. 4 D), and cells that were rescued with the
tagged protein, we again observed a concentration of gold particles
between the transition zone doublet microtubules and mem-
brane. In this case, they were centered longitudinally in the
transition zone, close to or over the wedge-shaped structures
(Figs. 4 B and S4). In cross sections, the gold particles labeled
Figure 3. Loss of CEP290 causes defects in the microtubule–membrane connections within the transition zone. (A and B) Longitudinal sections through the
transition zone (brackets) reveal that the electron-dense wedge-shaped structures (arrowheads in A) between the transition zone microtubules and flagellar
membrane are missing in the cep290 mutant. (C and D) Cross sections reveal that the Y-shaped connectors (arrowheads in C) that bridge the transition
zone microtubules with the membrane are missing from most cep290 doublet microtubules. Insets in C and D are image averages of 30 transition zone
doublet microtubules. (E–G) Some of the mutant flagella have bulges filled with electron-dense material (arrowheads in E). The flagellum shown in F also has
defects in axonemal doublets and the central pair microtubules; however, such defects are unusual in the mutant, as most axonemal cross sections appear
normal. (H) Histogram depicting the distribution of the distances between the transition zone cylinder and the flagellar membrane in wild-type and cep290
mutant cells; the difference in the distributions is significant at P = 0.0025 for a Student’s t test and P = 0.0009 for a Welch’s t test. (I and J) Cells were
detergent-extracted, then fixed and processed for EM. The detergent-resistant membrane that remains attached to the wild-type transition zone (arrows in I)
is no longer attached to the mutant transition zone. Bar, 100 nm.
JCB • VOLUME 190 • NUMBER 5 • 2010 932
and EF1-, were abnormally increased, whereas other pro-
teins, including flagellar adenylate kinase, FAP5 (conserved
uncharacterized flagellar associated protein; Pazour et al.,
2005), and FAP12 (triacylglycerol lipase), were missing or re-
duced in the mutant flagella (Fig. 5 B). Therefore, loss of
CEP290 affects the flagellar levels of a large number of pro-
teins, including membrane proteins and proteins associated
isolated from wild-type and cep290 mutant cells, and their pro-
teins were compared in Western blots and silver-stained gels.
The mutant flagella had increased amounts of IFT complex B
proteins and BBS4 relative to wild-type flagella, and decreased
amounts of the IFT complex A protein IFT139 and polycystin-2
(Fig. 5 A). Comparison of wild-type and mutant flagella in
silver-stained gels, followed by mass spectrometry to identify
selected bands, revealed that some proteins, including EF-3
Figure 4. Immunogold localization of CEP290. (A) Wild-type detergent-extracted cytoskeletons were incubated with CEP290 antibody and gold-
conjugated secondary antibody, then fixed, embedded, and sectioned. The locations of 34 gold particles from 30 sections are indicated by black dots
superimposed on the EM. For simplicity, the black dots are depicted on only one side of the transition zone; no bias for either side of the transition zone
was observed. A selection of original micrographs is shown in Fig. S3. (B and C) cep290::CEP290-HA cytoskeletons were double-labeled with antibodies
to CEP290 (12-nm gold) and HA (6-nm gold). As in A, the locations of the particles from several longitudinal sections (B) and cross sections (C) are repre-
sented by black dots superimposed on a single EM. A selection of original micrographs is shown in Fig. S4. Bars, 100 nm. (D) Schematic of HA-tagged
CEP290 depicting the locations of the HA and C-terminal epitopes.
933CEP290 function in the flagellar transition zone • Craige et al.
(Fig. 5 C). However, using high-contrast differential inter-
ference (DIC) microscopy to observe IFT in live cells, we
found that cep290 mutant cells had normal anterograde IFT,
and only a slight decrease in the velocity and frequency of
retrograde IFT (Fig. 5, D and E; and Videos 5 and 6). More-
over, our biochemical analysis indicated that the flagellar
level of the IFT motor subunit D1bLIC was normal (Fig. 5 A).
Therefore, loss of CEP290 has little effect on IFT motility
As noted above, some cep290 mutant flagella have
bulges; such bulges in cilia and flagella are usually caused
by an accumulation of IFT proteins and defects in retrograde
IFT (Pazour et al., 1998, 1999; Piperno et al., 1998; Porter
et al., 1999; Iomini et al., 2001, 2009; Hou et al., 2004; Tran
et al., 2008). To determine if this is the case in the cep290
mutant, we localized IFT complex A and B proteins in the
mutant cells by immunofluorescence microscopy, and indeed
found a concentration of IFT proteins in the flagellar bulges
Figure 5. cep290 mutant flagella have an abnormal protein composition. (A) Western blots of isolated flagella show an accumulation of IFT complex
B proteins (IFT20, IFT46, and IFT81) and BBS4, and a reduction in the IFT complex A protein IFT139 and polycystin-2 (PKD2). The arrowhead marks
an anti–FMG-1-immunoreactive band that is absent in the mutant flagella. (B) Silver-stained gel of isolated wild-type (WT) and mutant (cep290) flagella
demonstrating that some proteins are abnormally present (arrows) and others are missing or reduced (arrowheads) in the mutant flagella. Proteins identi-
fied by mass spectrometry include (1) flagellar adenylate kinase and FAP5, (2) FAP12, (3) EF-3, and (4) EF-11. The positions of standard proteins and
their molecular masses in kD are indicated. (C) Immunofluorescence indicates that bulges (arrowheads) in the cep290 mutant flagella contain IFT proteins.
(D) DIC imaging and kymographs of IFT in living cells. (D, a and b) Still images of the videos (Videos 5 and 6) used to create the kymographs (a and b).
Bars, 1 µm. (E) Velocities and frequencies of IFT in wild-type and cep290 mutant flagella. Data are represented as mean ± SD (error bars); P-values were
determined using a Student’s t test.
JCB • VOLUME 190 • NUMBER 5 • 2010 934
CEP290 is a component of the
microtubule–membrane linkages within the
The transition zone is a structurally complex and highly con-
served structure that lies between the basal body and the flagellar
axoneme. The ultrastructure of the C. reinhardtii basal body and
transition zone has been well characterized (Ringo, 1967;
Johnson and Porter, 1968; Cavalier-Smith, 1974; Weiss et al.,
1977; Gaffal, 1988; O’Toole et al., 2003; Geimer and Melkonian,
2004). At the distal end of the basal body, the triplet microtubules
of the basal body transition into doublets, marking the beginning
of the transition zone. Prominent features of the transition zone
are structures that bridge the doublet microtubules to the flagellar
membrane. In cross sections, these appear as thin Y-shaped con-
nectors (Fig. 3 C); in longitudinal sections, wedge-shaped elec-
tron densities are observed to slant proximally and distally from
the doublet microtubules to the membrane (Fig. 3 A; Ringo, 1967;
Gilula and Satir, 1972; Dentler, 2009a). The central pair micro-
tubules begin at the distal end of the transition zone, marking
the beginning of the axoneme.
Although the structure of the transition zone has been de-
scribed in detail, very little is known regarding the function of this
short and unique region of the flagellum (Dutcher, 2009). In addi-
tion to its role (along with the basal body) in templating the axo-
neme, evidence suggests that the transition zone is the site for
docking of the IFT particles (Deane et al., 2001). The structure
and location of the transition zone make it well poised to act as a
flagellar “pore,” regulating the import and export of proteins into
and out of the flagellum (Rosenbaum and Witman, 2002). The
transition zone also has been proposed to be a site for enrichment
of Ca2+ channels and pumps (Tamm, 1988). Related to this, Ca2+-
dependent flagellar abscission in C. reinhardtii occurs immedi-
ately distal to the transition zone (Quarmby, 2009).
Both immunofluorescence microscopy and immuno-EM
indicated that CEP290 is located at the transition zone; the
immuno-EM showed that the protein is located near the periph-
ery of the transition zone, between the outer doublets and the
transition zone membrane. Comparison of the labeling patterns
obtained for epitopes at the C terminus versus the central portion
of CEP290 indicated that the middle of the protein is closely
associated with the wedge-shaped structures seen in longitudinal
sections and the Y-shaped connectors seen in cross sections,
whereas the C terminus is more dispersed. CEP290 is certainly
large enough to account for this distribution of gold particles.
In silico analysis of the CEP290 amino acid sequence predicts
a high density of coiled-coiled domains throughout the length
of the protein. If CEP290 forms a large coiled-coiled structure,
it could be as long as 380 nm (Fraser and MacRae, 1973) and
potentially span the length of the transition zone.
Consistent with the localization of CEP290, the cep290 mu-
tant cells have striking defects in the structures located between
the transition zone doublet microtubules and the flagellar mem-
brane. Specifically, most of the Y connectors seen in cross section
were missing, and the wedge-shaped structures seen in longitudi-
nal section were missing or collapsed back onto the doublets. It is
CEP290 can incorporate into
preassembled wild-type or mutant
Currently, the only successful approach for treating LCA is
somatic gene therapy to replace the defective gene (Bainbridge
et al., 2008; Cideciyan et al., 2008; Hauswirth et al., 2008;
Maguire et al., 2008). Because the target in the case of CEP290
would be photoreceptor cells, which are not renewed during the
life of the organism, this approach would require that CEP290 ex-
pressed by a gene therapy vector be incorporated into preassembled
ciliary cytoskeletons that lack endogenous CEP290 or have cop-
ies of the mutant protein. Although no information is available on
whether transition zone components undergo turnover, this can be
tested in C. reinhardtii using dikaryon rescue. During mating of
C. reinhardtii, two gametic cells of opposite mating types come into
contact and fuse, resulting in a single quadriflagellated, binucleated
cell or dikaryon (Harris, 2009). When a wild-type cell is mated to
a null mutant, the protein missing in the mutant may be supplied
by the wild-type cell, and its ability to be incorporated into the
mutant flagellar cytoskeleton then observed. Similarly, when a cell
expressing a tagged protein is mated to a wild-type cell, the abil-
ity of the tagged protein to displace the endogenous protein in the
flagellar cytoskeleton of the untagged partner can be tracked.
We used this technique first to address whether wild-type
CEP290 can incorporate into preassembled cep290 mutant tran-
sition zones; i.e., transition zones lacking CEP290. Wild-type and
cep290 mutant gametes were mated, fixed at various time points,
and processed for immunofluorescence using antibodies to acety-
lated tubulin and CEP290. After 20 min, most of the quadri-
flagellates displayed CEP290 signal on a single pair of transition
zones, which must have derived from the wild-type cell; however
some of the quadriflagellates at this time point displayed a pair of
transition zones with a stronger CEP290 signal and a pair with a
weaker signal (presumably the mutant transition zones; Fig. 6 A,
a–h). Over the next 40 min, the relative strength of the CEP290
signal on the dimmer pair increased, whereas that on the brighter
pair decreased, so that by 60 min all four transition zones had
similar signal intensities (Fig. 6 A, i–p). This indicates that wild-
type CEP290 can assemble onto previously formed CEP290-
deficient transition zones. Moreover, the fact that the brightness
of the labeled and unlabeled transition zones equilibrated with
time suggested that CEP290 in the transition zone undergoes
relatively rapid exchange with CEP290 in the cytoplasm.
Next, to determine if new CEP290 can replace CEP290 pre-
viously assembled into wild-type transition zones, we mated wild-
type gametes to cep290::CEP290-HA gametes, which express
only HA-tagged CEP290. The resulting dikaryons were then pro-
cessed at various times for immunofluorescence microscopy using
antibodies to the HA tag and acetylated tubulin (Fig. 6 B). Initially,
HA-tagged CEP290 was present only on the two transition zones
derived from the HA-tagged parent. After 20 min, CEP290-HA
could be observed on all four transition zones, with one pair brighter
than the other. Equilibration of the HA signal on all four transition
zones occurred by 60 min. Therefore, newly supplied CEP290 can
replace CEP290 previously assembled into transition zones. These
results confirm that CEP290 is a dynamic component of the transi-
tion zone and rapidly equilibrates with cytoplasmic CEP290.
CEP290 function in the flagellar transition zone • Craige et al.
Figure 6. CEP290 is dynamic and can incorporate into preassembled mutant or wild-type transition zones. (A) Wild-type gametic cells were mated with
cep290 mutant gametic cells, and the fused, quadriflagellated cells were detergent-extracted, fixed, and processed for immunofluorescence at the indicated
time points after the two cell types were mixed for mating. The quadriflagellated cytoskeletons were double labeled with antibodies to CEP290 and acety-
lated tubulin. Arrowheads mark the dimmer pair of transition zones. (B) Wild-type gametes were mated to cep290::CEP290-HA gametes and processed
as in A except that CEP290-HA was visualized using anti-HA. Bars, 1 µm.
JCB • VOLUME 190 • NUMBER 5 • 2010 936
complex A and/or the accumulation of complex B in the mutant
flagella, as similar decreases in retrograde IFT velocities have
been observed in complex A mutants, which also have increased
complex B and decreased complex A in their flagella (Piperno
et al., 1998; Iomini et al., 2001, 2009). In any case, it seems un-
likely that a defect in IFT motility per se is the cause of the abnor-
mal balance of IFT complexes A and B in the cep290 flagella.
It should be noted that other C. reinhardtii mutants accumulate
IFT proteins in bulges at the flagellar tip yet display normal ve-
locities and frequencies of IFT (Tam et al., 2003; Dentler, 2005).
We propose that the abnormal balance of IFT particles and
the abnormal protein composition of cep290 mutant flagella re-
sult from loss of the microtubule–membrane connectors, with
concomitant defects in the remodeling, cargo loading/unloading,
or “quality control” of IFT particles as they pass through the tran-
sition zone. CEP290 and the membrane connectors within the
transition zone could provide a platform for IFT particles as they
are loaded and unloaded at the base of the flagellum, a checkpoint
that admits only proteins with flagellar targeting sequences, or a
“gate” through which only properly assembled IFT particles can
pass. Loss of the gate may allow an excess of IFT complex
B proteins to enter the flagellum, where they accumulate without
the proper complement of IFT complex A, which may be neces-
sary to couple them to the retrograde IFT motor for export from
the flagellum (Hao and Scholey, 2009). Alternatively, the mem-
brane connectors and/or CEP290 might be required for the effi-
cient entry of IFT complex A into the flagellum; defects in
complex A trafficking could also result in the accumulation of
IFT complex B, as is seen in C. reinhardtii strains with mutations
in IFT complex A components (Piperno et al., 1998; Iomini et al.,
2001, 2009). Because IFT is necessary for flagellar assembly and
maintenance, such defects could in turn lead to the many other
abnormalities in flagellar protein content that we observed. Previ-
ous studies reported mislocalization of some ciliary components
in mice harboring a hypomorphic mutation in Cep290 (Chang et al.,
2006; McEwen et al., 2007). Here, we demonstrate that loss of
CEP290 affects the normal levels of IFT particles in the flagel-
lum, which could provide mechanistic insight into why defects in
CEP290 affect ciliary assembly and the localization of other cili-
Loss of CEP290 results in abnormal
flagellar levels of ciliary disease proteins
Among the abnormalities in cep290 mutant flagella is a de-
creased level of polycystin-2, an important observation given
the prevalence of cystic kidneys in patients with CEP290 mu-
tations. Polycystin-2 is a six-pass transmembrane protein that
localizes to cilia and forms a cation-permeable channel; muta-
tions in the human gene (PKD2) are the cause of autosomal dom-
inant polycystic kidney disease type 2 (OMIM no. 613095). Our
results demonstrate that CEP290 is required for normal levels of
polycystin-2 in flagella, and suggest that cystic kidney disease in
patients with CEP290 mutations could result from defects in the
ciliary levels of polycystin-2.
Many of the disease symptoms caused by CEP290 muta-
tions are also present in patients with Bardet-Biedl syndrome
(BBS; OMIM no. 209900), and a homozygous mutation in
not yet clear if the Y connector and wedges are one and the same
structure viewed from two different angles, but their locations and
the fact that both are severely affected in the cep290 mutant sug-
gest a close relationship. CEP290 is likely to be an essential com-
ponent of the wedges, because immuno-gold EM showed that the
central part of CEP290 is localized to this structure, and the struc-
ture is missing when CEP290 is missing. CEP290 antibodies also
labeled the Y connectors; however, CEP290 is unlikely to be the
only component that comprises them, because Y connectors are
seen, albeit rarely, in the null mutant.
Loss of CEP290 and concomitant loss of the transition zone
connectors results in loss of attachment of the membrane to the
microtubules in the transition zone. Although C. reinhardtii mu-
tants with pleiotropic defects in basal bodies and transition zones
have been described previously (Goodenough and St. Clair, 1975;
Huang et al., 1982; Jarvik and Suhan, 1991; Dutcher and Trabuco,
1998; Piasecki et al., 2008; Piasecki and Silflow, 2009), this is the
first study of a mutation that specifically compromises the integ-
rity of the transition zone microtubule–membrane connectors.
The results demonstrate that the connectors physically tether the
transition zone membrane to the underlying microtubules, a func-
tion likely to be important for controlling entry of proteins into
CEP290 is important for flagellar assembly
and protein content
The cep290 mutant cells are palmelloid, which is a phenotype
commonly seen in C. reinhardtii mutants with defects in flagellar
assembly (Harris, 2009). Indeed, when cep290 cells are released by
digesting the mother cell wall with autolysin, the freshly released
cells display very short, stumpy flagella. For reasons not yet under-
stood, the flagella of many cep290 cells then slowly elongate to
various lengths. In mammalian cell culture models, CEP290 siRNA
decreases the percentage of cells displaying primary cilia (Graser
et al., 2007; Tsang et al., 2008). Collectively, these data indicate a
conserved requirement for CEP290 in ciliogenesis.
SDS-PAGE revealed numerous differences in the flagellar
proteins of wild type versus the cep290 mutant; some of these
were identified by mass spectrometry. Notably, cep290 flagella
have reduced amounts of flagellar adenylate kinase; of FAP12,
which is predicted to be a triacylglycerol lipase; and of FAP5,
which is a conserved flagellar-associated protein of unknown
function (Pazour et al., 2005). Loss of flagellar adenylate kinase
could affect ATP production in the flagellum, whereas loss of
triacylglycerol lipase could affect flagellar membrane lipid com-
position. Thus, either of these deficiencies could affect flagellar
assembly. EF-3 and EF-1 were elevated in cep290 flagella; the
functions of these proteins in flagella are unknown.
The flagella of cep290 mutant cells occasionally have
bulges filled with a finely divided material that includes IFT pro-
teins, and Western blot analysis of isolated flagella revealed that
the mutant flagella accumulate IFT complex B proteins and have
decreased levels of the IFT complex A component IFT139. When
cep290 cells were compared with wild-type cells, there was no
significant difference in anterograde IFT, but retrograde IFT was
slightly reduced both in velocity and frequency. The decrease in
retrograde IFT velocity may be related to the decrease in IFT
937CEP290 function in the flagellar transition zone • Craige et al.
In summary, our results show that CEP290 is a transition
zone protein localized to and required for the assembly of the
unique microtubule–membrane connectors in this region. Loss
of CEP290 and these structures results in an imbalance of IFT
complexes in the flagellum, and leads to numerous other abnor-
malities in the composition of flagella, including abnormal levels
of flagellar membrane proteins and proteins associated with
ciliopathies. Similar changes in ciliary protein composition are
almost certain to be the underlying cause of the human pathol-
ogies resulting from CEP290 mutation.
Materials and methods
Strains and cell culture
Strains were grown in liquid M medium I (Sager and Granick, 1953), with
2.2 mM KH2PO4 and 1.71 mM K2HPO4, aeration by 5% CO2, and a light/
dark cycle of 14/10 h (Witman, 1986). For some experiments, cells were
cultured without aeration or treated with autolysin in order to induce flagel-
lation. The following C. reinhardtii strains were used: 137c (CC-125; nit1,
nit2, mt+), g1 (nit1, agg1, mt+), CC-124 (nit1, nit2, agg1, mt), Y168
(nit1::NIT1, agg1, cep290, mt+), F1P4 (nit1::NIT1, agg1, cep290, mt),
cep290 (nit1::NIT1, agg1, cep290, mt+), cep290::CEP290 (nit1::NIT1,
agg1, cep290::CEP290, mt), and cep290::CEP290-HA (nit1::NIT1, agg1,
A synthetic peptide consisting of the last 14 amino acids of C. reinhardtii
CEP290 with cysteine added to the N terminus for conjugation to KLH
(C-ADGAGPSGQRRGGR) was used to generate rabbit polyclonal antibodies
(GenScript). The resulting antiserum was affinity purified on an agarose
support (Thermo Fisher Scientific) according to the manufacturer’s instruc-
tions. Additional antibodies used in this study were: rat high-affinity anti-HA
(Roche), anti-acetylated tubulin (Sigma-Aldrich), anti–-tubulin (Sigma-
Aldrich), anti-polyglutamylated tubulin (Wolff et al., 1992), anti–-F1-
ATPase (Agrisera), anti-BBS4 (Lechtreck et al., 2009), anti-IC2 (King et al.,
1985), anti-FMG1 (provided by R. Bloodgood, University of Virginia, Char-
lottesville, VA; Bloodgood et al., 1986), anti-PKD2 “loop” (provided by
K. Huang, Yale University, New Haven, CT; Huang et al., 2007), anti-D1bLIC
(Hou et al., 2004), anti-IFT20 (Hou et al., 2007), anti-IFT46 (provided by
H. Qin, Texas A&M University, College Station, TX; Hou et al., 2007), and
anti-IFT81 and anti-IFT139 (provided by D. Cole, University of Idaho, Mos-
cow, ID; Cole et al., 1998).
RT-PCR screen and backcrossing
A library of insertional mutants was screened by real-time PCR using the
QuantiTect SYBR Green PCR kit (QIAGEN) and primer pairs (see Table S1)
specific for the 5 and 3 ends of C. reinhardtii CEP290. Additional primer
pairs were used to map the deletion in strain Y168. Y168 was backcrossed
to wild type (CC-124) and the Y168 mutant phenotype cosegregated with
the deletion. A mutant progeny (F1P4) from this cross was then backcrossed
to the Y168 parent strain (g1 [Pazour et al., 1995], used as the wild-type
control strain for this study) and the mutant phenotype again cosegregated
with the deletion. A mutant progeny from the second backcross, strain
cep290, was used for the remainder of the study.
Cloning the CEP290 gene and rescue of the cep290 mutant
First, a cassette conferring phleomycin resistance (Stevens et al., 1996;
Lumbreras et al., 1998) was cloned into the HindIII site in pNEB193 (New
England Biolabs, Inc.) to make pBC1. Second, a bacterial artificial chromo-
some (BAC) containing C. reinhardtii CEP290 (BAC 29K10, Clemson Uni-
versity Genomics Institute) was digested with AscI and StuI to obtain a 9.3-kb
fragment that was cloned into pBC1 cut with AscI and PmeI, yielding pBC2.
Third, the BAC was digested with ApoI and AscI, and a 6-kb fragment was
purified and cloned into pBC2 also cut with ApoI and AscI, yielding pBC3.
To introduce a triple HA tag, p3×HA (Silflow et al., 2001) was digested with
SmaI and NaeI, and the resulting fragment containing the 3×HA sequence
was cloned into pBC3 by partially digesting pBC3 with ScaI, yielding pBC4.
All constructs were verified by sequencing. 5 µg of pBC3 or pBC4 plasmid
linearized with SspI were used to transform the F1P4 CEP290 mutant strain
or the cep290 strain, respectively, using glass bead agitation (Kindle, 1990).
Rescued transformants were identified visually by screening for wild-type
CEP290 (E1903X) was found in a BBS patient that also car-
ried a complex heterozygous mutation in MKS3 (Leitch et al.,
2008). Flagella of the cep290 mutant have increased levels of
BBS4, which is a component of the BBSome (Nachury et al.,
2007) and is trafficked in the flagellum by a subset of IFT par-
ticles (Lechtreck et al., 2009). This raises the possibility that
the amount of BBSomes in the flagellum is determined by the
same factors that control the amount of complex B, which is
also increased in cep290 flagella. In any case, the BBSome is
required for the removal of specific signaling proteins (e.g.,
phospholipase D) from flagella (Lechtreck et al., 2009), and
the phenotypic overlap between BBS and CEP290-associated
ciliopathies could be explained, at least in part, by the accu-
mulation of BBS4 in cep290 mutant cilia. It is unknown if
CEP290 interacts directly with the BBSome, but both CEP290
and BBS4 interact with the pericentriolar protein PCM-1 (Kim
et al., 2004; Kim et al., 2008), and CEP290 depletion by siRNA
shifted the sedimentation profile of BBS4 in sucrose gradients
(Kim et al., 2008).
Finally, given our finding that CEP290 is a component of
the transition zone, it is relevant that the connecting cilia of
mammalian photoreceptor cells are thought to be greatly elon-
gated transition zones (Besharse and Horst, 1990) and contain
microtubule–membrane Y connectors analogous to those seen in
the C. reinhardtii transition zone. Mutations in human CEP290
almost always are associated with blindness, regardless of whether
or not other organs are affected, so CEP290 may be especially
important in the function of the connecting cilium, through which
huge amounts of precisely sorted IFT-dependent cargo must
pass each day to maintain the photoreceptor outer segment.
CEP290 is dynamically associated with the
LCA caused by mutations in RPE65 has been treated by gene
therapy, with some individuals experiencing improvement in
visual acuity (Bainbridge et al., 2008; Cideciyan et al., 2008;
Hauswirth et al., 2008; Maguire et al., 2008). It has been estimated
that 20% of LCA cases are caused by mutations in CEP290
(den Hollander et al., 2006), making CEP290 an important target
for gene therapy in these patients. We demonstrated that CEP290
is dynamic and can incorporate into preassembled CEP290-
deficient and wild-type transition zones, a property that will be
essential for successful CEP290 gene therapy in the retina.
Our results indicate that CEP290 shuttles between the tran-
sition zone and the cytoplasm. In mammalian cells, CEP290 re-
distributes to the cytosol during mitosis (Sayer et al., 2006), a
time in the cell cycle when cilia are not present. In addition,
CEP290 contains a putative nuclear localization signal (Chang
et al., 2006; Sayer et al., 2006) that is conserved in C. reinhardtii,
CEP290 has been detected in the nucleus (Guo et al., 2004; Sayer
et al., 2006), and CEP290 has been shown to activate the tran-
scription factor ATF4 in a luciferase reporter assay (Sayer et al.,
2006). Thus, CEP290 may also be involved in signal transduc-
tion. The location of CEP290 at the transition zone and its dy-
namic properties would make it ideally suited for monitoring
flagellar status and relaying information about the flagellum back
to the cell body or nucleus.
JCB • VOLUME 190 • NUMBER 5 • 2010 938
embedded in epon, and thin-sectioned. Sections were stained with lead
citrate and imaged on an electron microscope (CM10; Philips). Images
were acquired using a digital camera (Gatan). For image averages, 30
wild-type or mutant cross-sections were aligned and superimposed over
each other with 10% transparency using Photoshop (Adobe). For analysis
of detergent-resistant membranes (Kamiya and Witman, 1984), cytoskele-
tons were prepared as described above (Immunofluorescence microscopy)
then fixed and processed for EM as described.
For immuno-EM, cytoskeletons were prepared as above (Immuno-
fluorescence microscopy) and fixed with 3% paraformaldehyde in MT
buffer (30 mM Hepes, 5 mM EGTA, 15 mM KCl, and 5 mM MgSO4, pH 7.2)
on ice for 20 min. After blocking in 2% BSA with 0.1% cold fish skin gelatin
and 0.05% Tween-20 in PBS for 1 h, the samples were labeled with pri-
mary antibody (1:25 rabbit anti-CEP290 and 1:50 rat anti-HA monoclonal
3F10) and gold-conjugated secondary antibody (1:20 goat anti–rabbit
12-nm gold conjugant and 1:20 goat anti–rat 6-nm gold conjugant) for
90 min each, then postfixed with 2.5% glutaraldehyde with 0.1% tannic
acid for 1 h, followed by 1% OsO4 at 4°C for 1 h. Sections were stained
with 2% uranyl acetate in methanol for 10 min.
Flagella isolation, SDS-PAGE, Western blotting, and mass spectrometry
To induce cep290 cells to hatch and assemble flagella, cells were cultured
in minimal medium in the absence of aeration. For comparisons, control
wild-type cells were cultured under identical conditions. Flagella were har-
vested using dibucaine (Witman et al., 1972; Witman, 1986) and resus-
pended in HMDEK supplemented with Complete Protease Inhibitor Cocktail
(Roche). Protein concentration of isolated flagella was determined using the
RC-DC Protein Assay (BioRad Laboratories). Equal amounts of wild-type and
mutant flagellar protein were analyzed by SDS-PAGE and silver staining
(BioRad Laboratories) or Western blotting. Silver-stained bands of interest
were excised and identified by mass spectrometry as described previously
(Lechtreck et al., 2009). For Western blots of whole-cell lysates, mid-log
phase cells were harvested by centrifugation, resuspended in 5× denatur-
ing sample buffer (50 mM Tris, pH 8.0, 160 mM DTT, 5 mM EDTA, pH 8.0,
50% sucrose, and 5% SDS), heated at 75°C for 5 min, then passed through
a 26-gauge needle three times.
Gamete generation and mating, crude autolysin production,
and dikaryon formation
To generate gametes, 137c (mt+) and CC124 (mt) cells were plated onto
TAP medium plates with 1.5% agar (Harris, 2009), grown for 5–7 d in
cycled light, then placed in dim cycled light for 2–4 wk. The cells were
then scraped into 10 ml of nitrogen-free medium (M-N) and agitated on
an orbital shaker overnight with continuous light in a 125-ml flask. The
next morning, the cells were gently collected by centrifugation, resuspended
in 10 ml of M-N diluted 1:5 with water + 10 mM Hepes, pH 7.4, and
subjected to 4 h of orbital rotation under constant light. For production
of crude autolysin, gametes of the two cell types were mixed for 5 min
and centrifuged, and the supernatant was filtered through a 0.8/0.2-µm sy-
ringe filter. Autolysin was either used immediately or flash-frozen with liquid
nitrogen and stored at 80°C. To mate cells for backcrossing, gametic cells
were treated for 1–2 h with autolysin to hatch cep290 gametes, and then
mixed for 1–2 h. The zygotes were then plated onto M medium plates with
4% agar and cultured for 24 h under constant light. Progeny from tetrads
were isolated as described previously (Pazour et al., 1999; Hou et al.,
2004), and real-time PCR was used to identify the progeny harboring the
deletion in CEP290. For experiments involving dikaryons, gametes of op-
posite mating types were mixed for various time points. The cells were then
detergent-extracted and processed for immunofluorescence as described
above (Immunofluorescence microscopy).
Mid-log phase cells were concentrated twofold and the pH was quickly
lowered to 4.5 by the addition of 0.5 N acetic acid. After 45 s, the pH was
neutralized to 7.0 by the addition of 0.5 N KOH. The cells were then fixed
and processed for immunofluorescence microscopy at various time points.
Online supplemental material
Fig. S1 shows that CEP290 remains at the transition zone after flagellar
abscission and during flagellar regeneration. Fig. S2 shows that CEP290
is not present in isolated flagella. Fig. S3 shows a representative selection
of EMs that were used to generate Fig. 4 A. Fig. S4 shows a represen-
tative selection of EMs used to generate Fig. 4 (B and C). Videos 1–4
show the motility of wild-type, the cep290 mutant, the cep290 mutant
following rescue with untagged CEP290, and the cep290 mutant follow-
ing rescue with HA-tagged CEP290, respectively. Videos 5 and 6 show the
motility, and the presence of the rescuing construct was confirmed by PCR
and Western blotting.
Live cell imaging and videos
Cells were cultured as described above, placed into an 8-well slide, and
sealed with Vaseline and a coverslip. The cells were visualized by dark-
field phase contrast using a microscope (Axioskop II plus; Carl Zeiss, Inc.)
equipped with a 10× Plan-Neofluar objective lens. Images were captured
at 30 frames/second with a UNIQ UP-610 digital charge-coupled device
camera, and image sequences were output as AVI files and processed
using ImageJ software, then converted into MOV files using QuickTime Pro
(Apple). To document motility (Fig. 1 B), the WalkingAverage function in
ImageJ was used to average 30 frames. For DIC imaging of IFT (summa-
rized in Dentler et al., 2009b), live cells were immobilized in 1% agarose,
sealed in an eight-well slide as described above, and observed using an
inverted microscope (Ti U; Nikon) equipped with a 60× 1.49 NA objec-
tive lens, a 1.4 NA oil condenser, a 1.5× relay lens, and DIC optics.
Illumination was provided by a Lumen 200 (Prior Scientific) light source,
and light was filtered through UV, green band pass, and heat-absorbing
filters. Images were captured at 24 frames per second with a Clara Interline
camera (Andor) and NIS-Elements software (Nikon). Image sequences were
cropped, image brightness and contrast was adjusted in ImageJ, and the
resulting image stacks were exported as AVI files using ImageJ, then con-
verted into MOV files using QuickTime Pro. Kymographs were generated
in ImageJ by drawing a line that included the entire width of the flagellum
(width 11) and the MultipleKymograph ImageJ plugin with a line width of 1.
IFT velocities and frequencies were calculated by measuring the slope and
number of the particle tracks in the kymographs, respectively.
Cells were cultured as described above and harvested by centrifugation.
In some experiments, cells were treated with autolysin for 1–2 h before pro-
cessing for immunofluorescence to induce the growth of flagella. For immuno-
fluorescence of whole cells, cells were allowed to adhere for 5 min to
polyethylenimine-coated coverslips, excess cells were wicked off, and the
coverslips were submerged in 20°C methanol for 20 min. The slips were
then air-dried, rehydrated, and blocked for 30 min at RT with blocking solu-
tion (6% fish skin gelatin, 1% BSA, and 0.05% Tween-20 in PBS, pH 7.0),
incubated with primary antibodies (either 30–60 min at RT or 4°C over-
night), washed 3× with blocking solution, incubated with secondary
antibodies (Alexa Fluor–conjugated IgG; Invitrogen) for 30 min at RT,
washed 3× with PBS, then mounted with ProLong Antifade Gold (Invitrogen).
For immunofluorescence of isolated cytoskeletons, cells were harvested by
centrifugation, resuspended in autolysin for 1–2 h, washed once with HMDEK
(30 mM Hepes, pH 7.4, 5 mM MgSO4, 1 mM DTT, 0.5 mM EGTA, and
25 mM KCl), resuspended in HMDEK, and chilled on ice. Then, an equal
volume of ice-cold HMDEK + 2% NP-40 Alternative (EMD) + 2× Complete
Protease Inhibitor (Roche) was added. The cells were lysed on ice for 15 min
then added to an equal volume of RT HMDEK + 4% paraformaldehyde, and
the detergent-extracted cytoskeletons were adhered onto poly-L-lysine–
coated coverslips for 15 min. The slips were rinsed twice with PBS, then air-
dried. The slips were then blocked, incubated with antibody, and mounted
as described above. Samples were imaged using a microscope (Axioskop
II plus) with a 100× Plan-Apochromat 1.4 NA objective equipped with a
DIC prism. Images were captured with a digital charge-coupled device
camera (MrM) and Axiovision software (both from Carl Zeiss, Inc.). When
applicable, Photoshop (Adobe) was used to make equal brightness and/or
contrast adjustments for samples under comparison.
For ultrastructural analysis, cells were harvested by centrifugation, gently
resuspended in conditioned medium, and then fixed by addition of an
equal volume of 2% glutaraldehyde in conditioned medium. After 15 min
at RT, the cells were pelleted, the primary fixative was removed, and the
cells were gently resuspended in secondary fixative (1% glutaraldehyde
and 100 mM sodium cacodylate, pH 7.2) and incubated for 2 h at RT.
The cells were then pelleted and washed three times for 15 min each
with 100 mM sodium cacodylate, pH 7.2. After the final wash, 50 µl of
cell pellet was removed and added to a microcentrifuge tube containing
50 µl of 2% agarose dissolved in 100 mM sodium cacodylate, pH 7.2.
The cell/agarose suspension was solidified on ice, and the solidified cell/
agarose block was removed by cutting off the bottom of the microcentri-
fuge tube. The cell/agarose block was then cut into 1 mm2 pieces, post-
fixed for 1 h on ice with 1% OsO4 in 50 mM sodium cacodylate, washed
three times with ice-cold water, and stained overnight at 4°C in freshly
prepared 1% uranyl acetate in water. The samples were then dehydrated,
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We thank Drs. R. Bloodgood (University of Virginia, Charlottesville, VA), D. Cole
(University of Idaho, Moscow, ID), B. Eddé (Centre National de la Recherche
Scientifique, Montpellier, France) K. Huang (Yale University, New Haven, CT),
and H. Qin (Texas A&M University, College Station, TX) for kindly providing
antibodies used in this study. We are grateful to D. M. Sanderson (University of
Massachusetts Medical School) for the loan of microscopy equipment.
This research was supported by National Institutes of Health grants
(GM030626 to G.B. Witman, GM087848 to B. Craige, and GM014642
to J.L. Rosenbaum), the Grousbeck Family Foundation (to G.B. Witman and
J.L. Rosenbaum), and the Robert W. Booth Endowment (to G.B. Witman).
G.B. Witman is a member of the University of Massachusetts Medical School
Diabetes and Endocrinology Research Center (DERC; DK32520), and core
resources used in this research were supported by a DERC grant (DK32520).
Submitted: 16 June 2010
Accepted: 11 August 2010
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