Download Localization of retinitis pigmentosa 2 to cilia is regulated by Importin 2

718
Research Article
Localization of retinitis pigmentosa 2 to cilia is
regulated by Importin 2
Toby W. Hurd1,*, Shuling Fan1,* and Ben L. Margolis1,2,‡
1
Department of Internal Medicine, University of Michigan, Ann Arbor, MI 48109, USA
Department of Biological Chemistry, University of Michigan, Ann Arbor, MI 48109, USA
2
*These authors contributed equally to this work
‡
Author for correspondence ([email protected])
Journal of Cell Science
Accepted 25 October 2010
Journal of Cell Science 124, 718-726
© 2011. Published by The Company of Biologists Ltd
doi:10.1242/jcs.070839
Summary
Ciliopathies represent a newly emerging group of human diseases that share a common etiology resulting from dysfunction of the
cilium or centrosome. The gene encoding the retinitis pigmentosa 2 protein (RP2) is mutated in X-linked retinitis pigmentosa. RP2
localizes to the ciliary base and this requires the dual acylation of the N-terminus, but the precise mechanism by which RP2 is
trafficked to the cilia is unknown. Here we have characterized an interaction between RP2 and Importin 2 (transportin-1), a member
of the Importin- family that regulates nuclear–cytoplasmic shuttling. We demonstrate that Importin 2 is necessary for localization
of RP2 to the primary cilium because ablation of Importin 2 by shRNA blocks entry both of endogenous and exogenous RP2 to the
cilium. Furthermore, we identify two distinct binding sites of RP2, which interact independently with Importin 2. One binding site
is a nuclear localization signal (NLS)-like sequence that is located at the N-terminus of RP2 and the other is an M9-like sequence
within the tubulin folding cofactor C (TBCC) domain. Mutation of the NLS-like consensus sequence did not abolish localization of
RP2 to cilia, suggesting that the sequence is not essential for RP2 ciliary targeting. Interestingly, we found that several missense
mutations that cause human disease fall within the M9-like sequence of RP2 and these mutations block entry of RP2 into the cilium,
as well as its interaction with Importin 2. Together, this work further highlights a role of Importin 2 in regulation of the entry of
RP2 and other proteins into the ciliary compartment.
Key words: Cilia, Importin, RP2, Retinitis pigmentosa
Introduction
Cilia are highly evolutionarily conserved organelles. They can
mediate many different sensory functions, which are dependent on
the cell type on which they are located (Berbari et al., 2009;
Silverman and Leroux, 2009). In addition, cilia function in motility
at the level of both single-celled and multicellular organisms.
Despite this array of functions, the basic structure of cilia appears
to be highly conserved irrespective of cell type (Pedersen et al.,
2008; Rosenbaum and Witman, 2002). Cilia are nucleated from a
mother centriole/basal body and contain a ring of nine microtubule
doublets that run along the length of the cilium, known as the
ciliary axoneme. Some motile cilia also have a central microtubule
pair to facilitate cilia bending. Early studies in the biflagellate
Chlamydomonas reinhardtii identified two intraflagellar transport
(IFT) protein complexes (Complex A and Complex B), which
appear to traffic large protein aggregates or rafts from the basal
body to the distal tip of the cilium and back (Rosenbaum and
Witman, 2002). It has been demonstrated that these IFT proteins
are highly conserved evolutionarily and many are required for cilia
formation, whereas others function peripherally to direct specific
cargo into the cilia.
Despite the growing understanding of IFT function and cilia
assembly, there is much to learn about the regulation of ciliary
trafficking. Specifically, the primary cilium is believed to be
biochemically distinct from other subcellular compartments with
respect to protein and lipid content. Thus the cilia are able to
specifically segregate cilia proteins from non-cilia proteins
(Rosenbaum and Witman, 2002). This segregation via
compartmentalization provides a mechanism that allows for
exquisite regulation of cilia-mediated signaling (Pazour and
Bloodgood, 2008). How the cilia regulate import and export of
proteins is still unclear, but it might involve an electron-dense area
at the base of the cilia, which is known as the transition zone. This
area might function in a similar fashion to the tight junctions of
epithelia, which specifically segregate apical from basolateral
plasma membrane domains by preventing the admixture by lateral
diffusion of membrane proteins from each domain. The cilia
transition zone might also function in a similar manner to the
nuclear pore complex (NPC), a multi-protein complex that allows
the regulated import and export of specific proteins between the
cytosol and nucleus (Rosenbaum and Witman, 2002).
Indeed, a role for a transport mechanism to cilia and centrosomes
that appears to use the nuclear transport system involving Ran and
Importin proteins has recently been uncovered. One study focused
on cilia and pericentrosomal trafficking by a splice variant of the
Crumbs3 polarity protein (Fan et al., 2007). Another more recent
work focused on Kif17, a motor that is important in IFT (Dishinger
et al., 2010). These cilia-targeted proteins appear to use classical
nuclear localization signals to mediate their transport. However,
several other protein motifs have also been identified that facilitate
trafficking of proteins to the cilia (Pazour and Bloodgood, 2008).
Most notably, a VxPx motif has been identified in Polycystin-2,
CNGB1b and Rhodopsin, and is necessary for localization of these
proteins to cilia (Geng et al., 2006; Jenkins et al., 2006; Mazelova
et al., 2009). Recent work in the retina identified that this motif
might be bound by the small GTPase Arf4 (ADP-ribosylation
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Importin 2 and RP2
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factor 4). Furthermore, in conjunction with Rab11, FIP3 and
ASAP1 (Arf-GAP with SH3 domain, ANK repeat and PHdomain-containing protein 1), Arf 4 appears to regulate the
budding of cilia-destined vesicles from the trans-Golgi (Mazelova
et al., 2009).
In this paper, we focus on mechanisms that mediate the
trafficking of retinitis pigmentosa 2 (RP2, also known as protein
XRP2) to the cilia. RP2 is an X-linked gene that, when mutated,
gives rise to retinitis pigmentosa (Hardcastle et al., 1999). The RP2
gene encodes a 350 residue polypeptide that appears to be
ubiquitously expressed. Previous work has demonstrated that RP2
is membrane-associated by dual acylation of the N-terminus,
namely myristoylation at Gly2 and palmitoylation at Cys3 (Chapple
et al., 2002). The N-terminal half of RP2 has both sequence and
structural similarity to Tubulin cofactor C (TBCC), a protein that
is involved in the formation of -tubulin–-tubulin heterodimers.
Further work has shown that this domain binds and has GAP
activity towards the small G-protein Arl3 (Veltel et al., 2008).
Most recently RP2 has been demonstrated to traffic to the ciliary
base and proposed to have a role in Golgi-to-cilia membrane
delivery through its regulation of Arl3 (Evans et al., 2010). To
examine the mechanism by which RP2 traffics to cilia, we
performed large-scale purification of RP2 and associated proteins
from renal epithelia. Using this approach, we identified Importin
2 as a binding partner of RP2 that appears to regulate RP2
trafficking into the cilia. This work adds further support to the
recent findings that Importin proteins have a role in cilia trafficking.
Results
RP2 localizes to primary cilium in epithelia
It has been demonstrated that RP2 localizes to ciliary basal bodies
in several different cell types (Evans et al., 2010). Our laboratory
has studied RP2 localization in MDCK cells. We consistently
found both endogenous RP2 (Fig. 1A) and exogenous EGFPtagged RP2 (Fig. 1B) localized to primary cilia in fully polarized
MDCK cells. Confocal microscope images demonstrated that beside
its ciliary localization, the exogenously expressed RP2–EGFP also
localized to both apical and basal membranes in polarized MDCK
cells (Fig. 1C).
RP2 forms a complex with Importin-
To elucidate the mechanism by which RP2 is trafficked to the
cilium, a proteomic approach was adopted to identify new
interacting proteins. MDCK cells stably expressing either empty
vector (pQCXIP) or FLAG-tagged RP2 were generated (RP2–
FLAG). Tandem mass spectrometry (MS) analysis of proteins that
co-immunoprecipitated with RP2–FLAG identified several proteins
including Importin 2 (Transportin-1), Importin 3 and -tubulin
(Fig. 2A and supplementary material Table S1). These interactions
were confirmed by western blotting of RP2–FLAG
immunoprecipitates. All three endogenous proteins, Importin 2,
Importin 3 and -tubulin, co-immunoprecipitated with RP2–
FLAG (Fig. 2B). We have previously demonstrated that Importin
1 interacts with another cilia protein, Crumbs3–CLP1, so we
analyzed whether Importin 1 also interacted with RP2 (Fan et al.,
2007). Indeed, endogenous Importin 1 also efficiently coimmunoprecipitated with RP2–FLAG (Fig. 2B). Next, we examined
whether these Importin proteins could co-immunoprecipitate
endogenous RP2. MDCK cells were generated stably expressing
Myc-tagged Importin 1, Importin 2 and Importin 3. Using
these cells, we demonstrated that immunoprecipitation of the
Fig. 1. RP2 localizes to primary cilium in epithelia. (A)Endogenous RP2
localizes to primary cilia in MDCK cells. MDCK cells were grown on
Transwell filters for 7 days after confluence. Cells were then fixed and
immunostained with rabbit anti-RP2 antibody (green) and mouse antipolyglutamylated tubulin (red) and nuclei were stained with DAPI (blue).
(B)RP2–EGFP localizes to primary cilia. Fully polarized MDCK cells stably
expressing RP2–EGFP (green) were stained with polyglutamylated tubulin
(red, top panel) or g-tubulin (red, bottom panel). (C)RP2–EGFP localizes to
both apical and basolateral membranes. Fully polarized RP2–EGFP MDCK
cells were fixed and stained with DAPI to indicate nuclei. Representative zprojections are shown. Scale bars: 10m.
Importin proteins with an anti-Myc antibody, but not a control
anti-HA antibody, could efficiently co-immunoprecipitate
endogenous RP2 (Fig. 2C). To further test these interactions, RP2–
FLAG and the Myc-tagged Importin proteins were transiently
transfected into HEK293 cells and anti-FLAG immunoprecipitation
performed. Interestingly, in this cell type, although none of the
Importin proteins co-immunoprecipitated in the absence of RP2–
FLAG, only Importin 2 co-immunoprecipitated with RP2–FLAG
(Fig. 2D), suggesting that it is the major Importin  binding partner
of RP2. Furthermore, GST–Importin-2 was able to precipitate
endogenous RP2 from MDCK cell lysates (see below).
Importin 2 localizes to centrosome and cilium
We examined the subcellular localization of Importin 2 in MDCK
cells. When MDCK cells stably expressing Myc-tagged Importin
2 were grown on Transwell filters for 7 days after confluence,
Myc–Importin-2 was distributed in a punctate fashion along the
length of the ciliary axoneme (Fig. 3A). This was similar to what
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Fig. 2. RP2 forms a complex with Importin proteins. (A)Large-scale
protein purification of RP2-interacting proteins. MDCK cells stably expressing
either empty vector (pQCXIP) or RP2–FLAG were lysed and
immunoprecipitated with immobilized FLAG antibody. Immunoprecipitates
were extensively washed and separated by Bis-Tris PAGE. Resulting gels were
silver stained and bands of interest excised and submitted for MS/MS analysis.
(B)RP2–FLAG was immunoprecipitated from stable MDCK cells and the
immunoprecipitates were subjected to Bis-Tris PAGE and blotted with the
indicated antibodies. (C)MDCK cells stably expressing Myc-tagged Importin
1, Importin 2 or Importin 3 were immunoprecipitated with either anti-HA
(control) or anti-Myc antibody. Immunoprecipitates were subjected to Bis-Tris
PAGE and western blotted with the antibodies indicated. (D)HEK293 cells
were transiently transfected with RP2–FLAG and Myc-tagged Importin 1,
Importin 2 or Importin 3, respectively. Then cells were lysed and
immunoprecipitated with anti-FLAG antibody and immunoprecipitates were
subjected to Bis-Tris PAGE and blotted with the indicated antibodies.
was observed in Odora cells (Dishinger et al., 2010). Next, we
examined the localization of Myc-tagged Importin 2 in
subconfluent MDCK cells (1 day) and fully polarized MDCK cells
(7 days). In these cells, Myc-tagged Importin 2 appeared to be
almost exclusively localized to the nucleus (Fig. 3B, top) at
subconfluence. However, as the cells became fully polarized, a
redistribution of Myc–Importin-2 to a subapical region was
observed (Fig. 3B, bottom). It is important to note that we used
0.1% Triton X-100 to detect cilia staining of Importin 2, whereas
to detect nuclear and cell body staining, we used 0.25% Triton X100.
We have previously shown that Importin 1 and the polarity
protein Crumbs3–CLP1 interact and colocalize at centrosomes in
mitotic cells and in primary cilium in fully polarized MDCK cells
(Fan et al., 2007). Accordingly, we next examined whether Importin
Fig. 3. Importin 2 localizes to primary cilia and centrosomes in
epithelia. (A)Importin 2 localizes to cilia. MDCK cells stably expressing
Myc-tagged Importin 2 were grown on Transwell filters for 7 days after
confluence, fixed with 4% paraformaldehyde, permeabilized with 0.1%
Triton X-100 and stained with anti-Myc antibody (green) and antipolyglutamylated tubulin antibody (red). (B)Importin 2 translocates from
nuclear to subapical regions during polarization in MDCK cells. MDCK cells
stably expressing Myc–Importin 2 were seeded onto Transwell filters and
grown for the times indicated (1 day or 7 days). Cells were then fixed,
permeabilized with 0.25% Triton X-100 and immunostained with antibodies
against Myc (green) and ZO-1 (red); nuclei were stained with DAPI (blue).
Images were captured by the confocal microscope to assemble X-Y and Zstack images. (C)Importin 2 co-sediments with purified centrosomes.
Centrosomes were isolated from fully polarized MDCK cells and separated
on a discontinuous sucrose gradient. Gradients were fractionated and
fractions subjected to Bis-Tris PAGE and western blots probed with the
indicated antibodies. Lysate (1%) is shown on the right. Actin control in the
bottom panel indicates centrosomal fractions have minimal contamination by
the cytoskeleton. (D)Importin 2 localizes near centrosomes in confluent
MDCK cells. MDCK cells were grown for 7 days after confluence on
Transwell filters. Cells were then fixed with acetone and stained with
antibodies against pericentrin (red) and Importin 2 (green). Nuclei were
visualized with DAPI (blue). Shown are X-Y and Z projections, respectively.
(E)GFP–Importin-2 localizes to spindle poles. MDCK cells stably
expressing GFP–Importin-2 (green) were seeded at low confluence on
coverslips. Cells were then fixed with acetone and centrosomes stained with
anti-g-tubulin (red); nuclei are stained with DAPI (blue). (F)RP2–EGFP
localizes to a pericentrosomal region during cell division. MDCK cells stably
expressing RP2–EGFP (green) were treated as in E. Scale bars: 10m.
Importin 2 and RP2
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2 is also associated with centrosomal fractions. We purified
centrosomes from confluent MDCK cells and examined whether
Importin 2 would co-sediment with the centrosome marker gtubulin using a discontinuous sucrose gradient. Indeed, in the high
density g-tubulin-positive fractions, enrichment of endogenous
Importin 2 was observed (Fig. 3C). We used actin as a control to
show that this high-density g-tubulin fraction was not contaminated
by cytoskeletal components. In addition, when we looked at the
localization of endogenous Importin 2 in confluent MDCK cells,
it appeared that a subfraction localized to centrosomes (Fig. 3D).
We next examined the localization of RP2 and Importin 2 in
mitotic cells. A fraction of exogenously expressed EGFP–Importin2 was enriched at the spindle poles of mitotic cells, as shown by
colocalization with the centrosome marker g-tubulin (Fig. 3E). In
addition, EGFP-tagged RP2 was localized to a pericentrosomal
region (Fig. 3F), similarly to our previously reported data for
Crumbs3–CLP1 and Importin 1 (Fan et al., 2007).
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RP2 interacts with the C-terminus of Importin 2
GST fused full-length Importin 2 was able to pull down
endogenous RP2 (Fig. 4A). We examined which region of Importin
2 was involved in the interaction with RP2. Importin  proteins
share a similar domain architecture with an N-terminal Ran-binding
domain (RBD) followed by HEAT and Armadillo repeats (Fig.
4C). Because the RBD mediates binding of the GTP-bound form
of the small GTPase Ran to Importin  proteins, we next tested
whether constitutively active Ran (Ran Q69L) could precipitate a
complex of Importin 2 and RP2. Importin 2 was efficiently
precipitated by WT and Q69L Ran, but no RP2 was present in the
complex (Fig. 4B). This is consistent with the ability of Ran-GTP
to dissociate cargo from Importin . To examine which region of
Importin 2 is necessary for the interaction with RP2, a series of
Importin 2 N-terminal truncations were generated (Fig. 4C). As
previously demonstrated, full-length (FL) Myc–Importin-2 was
co-immunoprecipitated by RP2–FLAG (Fig. 4D). However,
deletion of either the first 320 residues of Importin 2,
encompassing the RBD and first set of HEAT repeats, or the first
546 residues did not abolish this interaction (Fig. 4D). These data
suggest that the C-terminal domain of Importin 2 is sufficient to
bind RP2, as has been shown with other cargoes (Stewart, 2007).
Knockdown of Importin 2 blocks RP2 ciliary targeting
Next, we wanted to test the roles of Importin 2 in targeting of RP2
to the cilium. We used two shRNA targeting sequences to knock
down Importin 2 and test its role in the ciliary targeting of
endogenous RP2 and RP2–EGFP in MDCK cells. Both of the
shRNAs sufficiently reduced Importin 2 without affecting
expression of Importin 1 and RP2 (Fig. 5A). Cells without Importin
2 grew normally, but RP2 did not travel efficiently to the cilia when
Importin 2 expression levels were reduced (Fig. 5B). We next
generated stably expressing RP2–EGFP cells by co-infecting RP2–
EGFP into control shRNA and Importin 2 shRNA cells. After
selection with G418, expression levels of Importin 2 were still
suppressed, whereas expression of RP2–EGFP was similar in control
and Importin 2 shRNA cells (supplementary material Fig. S1A,B).
RP2–EGFP ciliary targeting was markedly reduced in cells in which
Importin 2 was knocked down (Fig. 5C). This lack of cilia targeting
in knockdown cells was not related to cilia length or RP2–EGFP
expression (supplementary material Fig. S1C). These results are
quantified in Fig. 5D, indicating significant impairment of RP2
ciliary targeting after knockdown of Importin 2.
Fig. 4. RP2 interacts with the C-terminus of Importin 2. (A)GST–
Importin 2 precipitates endogenous RP2. MDCK lysates were subjected to
GST pull-down with either GST alone or GST–Importin-2 and subjected to
Bis-Tris PAGE. Western blots were probed with indicated antibodies.
(B)Ran does not precipitate RP2. MDCK lysates were subjected to GST
pull-down with GST alone, wild-type (WT) Ran, dominant-negative (T24N)
Ran or constitutively active (Q69L) Ran. Precipitates were analyzed as in A.
(C)Schematic diagram showing domain organization of Importin 2 (RBD,
Ran-binding domain; HEAT HEAT repeat) and N-terminal truncations.
(D)The C-terminus of Importin 2 is sufficient to interact with RP2.
HEK293 cells were transiently transfected with RP2–FLAG and either Myctagged Importin 2 full-length (FL), Importin 2 lacking the N-terminal 320
residues (321-END) or the N-terminal 546 residues (547-END). Cells were
lysed and subjected to FLAG immunoprecipitation. Immunoprecipitates
were subjected to Bis-Tris PAGE and blots probed with the antibodies
indicated.
Binding sites on RP2 for Importin 2
Importin complexes regulate the trafficking of proteins between
the cytosol and the nucleus. Proteins destined for the nucleus
contain a nuclear localization signal (NLS) consisting of either a
single cluster (monopartite) or two clusters (bipartite) of basic
amino acid residues. Examination of the sequence of RP2 revealed
a highly conserved basic cluster at the N-terminus resembling the
consensus K-R/K-x-K/R monopartite NLS (Fig. 6A). Following
this putative NLS is a string of acidic residues that have previously
been shown to improve the efficacy of other NLS sequences (Fig.
6A). This type of NLS sequence can bind Importin- proteins, but
also can bind directly to Importin- proteins (Jkel and Grlich,
1998). To test whether this sequence mediates the interaction
between RP2 and Importin 2, a series of expression constructs
were generated driving expression of the N-terminus of RP2 fused
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Fig. 5. Loss of Importin 2 inhibits RP2 ciliary trafficking.
(A)Stable knockdown of endogenous Importin 2. MDCK
cells were infected with retrovirus directing expression of two
different canine Importin 2 shRNAs or a control shRNA
(luciferase shRNA) and stable pools were selected. Lysates
were subjected to Bis-Tris PAGE and blotted with the
antibodies indicated. (B)Knockdown of Importin 2
significantly reduces ciliary localization of endogenous RP2.
Control shRNA (top panel) and Importin 2 shRNA (bottom
panel) infected MDCK cells were grown for 7 days after
confluence on Transwell filters, fixed and stained with
antibodies against RP2 (green) and polyglutamylated tubulin
(red). (C)Ablation of Importin 2 blocks ciliary localization
of exogenous RP2-EGFP. MDCK cells infected with control
shRNA (upper panel) and Importin 2 shRNA-2 (bottom
panel) were stably transfected with RP2–EGFP (green) and
treated as in B. M-1 represents high magnification of RP2EGFP in control shRNA cells and M-2 represents RP2–EGFP
in Importin 2 shRNA cells. (D)Quantification of results from
experiments depicted in C. 100 ciliated RP2–EGFP cells each
from control shRNA, Importin 2 shRNA1 and shRNA2 were
evaluated. Results represent the mean of three individual
experiments, shown as mean ± s.d. Scale bars: 10m.
to EGFP. GST–Importin-2 pull down from HEK293 lysates
transfected with these constructs revealed that the first 15 residues
of RP2 are sufficient to mediate the interaction with Importin 2
(Fig. 6B). In addition, the RP2–EGFP 1–15 fusion protein is
efficiently trafficked to the primary cilium in MDCK cells
(supplementary material Fig. S2A).
It has been previously demonstrated that targeting of RP2 to the
plasma membrane requires myristoylation and palmitoylation of
Gly2 and Cys3, respectively (Chapple et al., 2002). Furthermore,
lipid modification is also required for targeting of RP2 to the basal
body (Evans et al., 2010). Thus we examined whether these
modifications are also necessary for the interaction with Importin
2. When HEK293 cells were co-transfected with Myc-tagged
Importin 2 with WT, G2A point mutant (which blocks
myristoylation and palmitoylation) or the C3S point mutant (which
blocks only palmitoylation) and a R118H mutant (which blocks
interaction with the small GTPase Arl3) (Khnel et al., 2006) of
RP2–FLAG, both WT and R118H were efficiently coimmunoprecipitated with Myc–Importin-2. However, neither the
G2A nor C3S mutants co-immunoprecipitated with Importin 2
(supplementary material Fig. S2B). These results suggest that
membrane association of RP2 is a prerequisite for formation of an
RP2–Importin-2 complex.
We further examined the RP2 sites required for interaction with
Importin 2. MDCK cells were transfected with the following
FLAG-tagged RP2 mutants: G2A, an N-terminal deletion of the
first 20 amino acids, the NLS mutant that converted KRRK to
AAAA, R118H and another mutation in the C-terminus, R282W
(Thiselton et al., 2000). Removal of the first 20 amino acids or
lipid modification (G2A) severely impaired binding to Importin 2
(Fig. 6C) as expected from the results previously obtained in
HEK293 cells (supplementary material Fig. S2B).
At this point, we thought that Importin 2 would bind RP2
through the N-terminal NLS sequence. Surprisingly, we found that
mutation of this polybasic NLS sequence (KRRK to AAAA) without
mutating the sites of lipid modification did not abolish Importin 2
binding (Fig. 6C). This indicated that RP2 must have an alternative
Importin 2 binding site. Another binding site for Importin 2 is
the M9 sequence found in heteronuclear ribonucleoprotein A1
(hnRNPA1) (Pollard et al., 1996). Indeed, we found an M9-corelike sequence in RP2 (Fig. 6D), which appeared to be essential for
Importin 2 binding to RP2 because point mutants of the M9
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Importin 2 and RP2
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Fig. 6. Two distinct binding sites of RP2 interact independently with Importin 2. (A) The N-terminus of RP2 contains a nuclear localization signal (NLS)-like
sequence. Blue, basic residues; red, acidic residues. (B)The first 15 residues of RP2 can interact with Importin 2. HEK293 cells were transiently transfected with
EGFP alone, full-length RP2–EGFP or residues 1–15, 1–25 or 1–50 of RP2 fused to EGFP. Cell lysates were subjected to precipitation with GST–Importin-2.
Precipitates were subjected to Bis-Tris PAGE and blots were probed with indicated antibodies. (C)Wild-type and mutant FLAG-tagged RP2 constructs were
transfected into MDCK cells. Lysates were than immunoprecipitated with anti-FLAG antibodies and lysates (input) or FLAG immunoprecipitates were resolved on
Bis-Tris gels. After transfer to nitrocellulose, the blots were probed with antibodies against Importin 2 (ImpB2) and FLAG. (D)RP2 contains an M9-like
sequence. Alignment of a sequence from RP2 and the classic M9 sequence found in hnRNPA1 and the consensus M9 sequence that can mediate binding of
Importin 2. The alignment and terminology is from Bogerd and colleagues (Bogerd et al., 1999) with J representing a hydrophilic amino acid and Z, a
hydrophobic amino acid. (E)Point mutations of the M9 consensus (C86Y, P95L or C108G) abolish the interaction between Importin 2 and RP2. FLAG-tagged
RP2 with various mutations were transfected into HEK293 cells with Myc-tagged Importin 2. After immunoprecipitation with anti-FLAG antibody, proteins were
resolved on a Bis-Tris gel and transferred to nitrocellulose. Blotting was then performed with anti-FLAG or anti-Myc antibodies.
consensus residues of RP2 (C86Y, P95L or C108G) abolished the
interaction with Importin 2 (Fig. 6E). Interestingly, these residues
have been found to be mutated in patients with retinitis pigmentosa
(Sharon et al., 2000). This suggests that the M9 sequence is part of
the main Importin 2 binding site in RP2 because mutation in this
site severely reduced binding and the polybasic NLS binding site
was unable to compensate for this.
The interaction between Importin 2 and M9 sequence of
RP2 is crucial for RP2 ciliary targeting
The results of the binding experiments correlated well when
trafficking of RP2 to the cilia was studied. As shown in Fig. 7A,
mutation of the polybasic NLS region did not block Importin 2
binding and did not prevent targeting to the cilia. By contrast,
mutations within the M9 sequence blocked both Importin 2
binding and cilia targeting (Fig. 7B).
Discussion
Mutation of RP2 leads to retinitis pigmentosa but the exact role of
RP2 in photoreceptors is unclear (Evans et al., 2006). Recent work
has demonstrated that RP2 binds to the small G protein ARL3
(Bartolini et al., 2002) and has GAP activity towards this GTPase
(Veltel et al., 2008). A recent study (Evans et al., 2010)
demonstrated that RP2 localizes to the basal body and is involved
in Golgi to ciliary base trafficking. In addition to basal body
staining, we found that RP2 localized throughout the cilia in renal
epithelial cells.
In this manuscript, we identified the binding of RP2 to Importin
2 and found that this binding appears important in regulating the
targeting of RP2 to cilia. Importin proteins have been classically
thought to regulate nuclear traffic (for a review, see Stewart, 2007).
Importin 1 uses Importin- proteins as adaptors to bind to
polybasic NLS sequences on proteins to be imported into the
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Fig. 7. The interaction between Importin 2 and M9 sequence of RP2 is
crucial for RP2 ciliary targeting. (A)Mutation of NLS region of RP2 does
not prevent RP2 ciliary targeting. MDCK cells were transfected with RP2–
EGFP (green) with the NLS motif mutated (KRRK–AAAA). Cilia are stained
with anti-polyglutamylated tubulin (red). (B)Point mutations in the M9
sequence, either C89Y or P95L of RP2–EGFP blocks ciliary targeting. MDCK
cells expressing EGFP–RP2 with point mutation of the M9 sequence, C89Y
(top panel) or P95L (bottom panel) were grown for 7 days after confluence on
Transwell filters. Cilia are stained with anti-polyglutamylated tubulin (red),
whereas EGFP–RP2 is green. Scale bars: 10m.
nucleus. Ran GTP releases the cargo as well as Importin- proteins
from Importin 1 within the nucleus (for a review, see Cook et al.,
2007). Importin 2 (also known as transportin-1) does not use
Importin- proteins but rather directly interacts with cargoes,
releasing them upon binding Ran GTP (Pollard et al., 1996;
Bonifaci et al., 1997; Siomi et al., 1997). Importin 2 usually binds
a different sequence than Importin 1, such as the M9-like
sequences we describe in RP2 (Michael et al., 1995; Weighardt et
al., 1995; Bogerd et al., 1999). Mutation of this M9 sequence in
RP2 impaired binding of RP2 to Importin 2 and impaired RP2
trafficking to the cilia. We also identified a polybasic NLS sequence
in the N-terminus of RP2. However, this polybasic sequence could
not support Importin 2 binding in full-length RP2 when the M9
sequence was mutated. We did observe trafficking to the cilia of
the smaller 1–15 GFP–RP2 fragment, which contains only the
lipid modified sites and the polybasic NLS. The trafficking of this
fragment might be different than that of the full-length RP2 because
of its smaller size, which allows easier ciliary entry, or more
efficient interaction of the polybasic NLS in this smaller fragment
with Importin proteins.
The binding and trafficking data reported here define additional
roles for Ran and its binding partners aside from nuclear trafficking.
Indeed, studies have already identified additional cellular roles for
Ran and its binding partners. One of the best described additional
roles is the trafficking and regulation of proteins that are crucial
for spindle assembly during cell division (for a review, see
Ciciarello et al., 2007; Clarke and Zhang, 2008; Kalab and Heald,
2008). In this process, Importin proteins bind and inhibit factors
such as HURP and TPX2, which are essential for microtubule
spindle growth, and this inhibition is relieved by Ran-GTP (Gruss
et al., 2001; Nachury et al., 2001; Wiese et al., 2001; Silljé et al.,
2006). Although these roles have been ascribed for Importin 1
and its Importin- binding partners, Importin 2 might also have
a role in mitotic spindle assembly (Lau et al., 2009). In this report
and other recent reports, we also suggest a further role for Importin
proteins in centrosomal and cilia targeting (Dishinger et al., 2010;
Fan et al., 2007). In the first of these studies, we demonstrated the
trafficking of a Crumbs3 splice isoform to the cilia and
pericentrosomal region (Fan et al., 2007). We described a role for
Ran and Importin 1 in this trafficking. More recently, Verhey and
co-workers have demonstrated a role for Importin 2 in the
trafficking of Kif17 to the cilia through an NLS sequence in the Cterminus (Dishinger et al., 2010). These data fit well with the
hypothesis that Ran and Importin proteins control cilia traffic,
possibly via the previously hypothesized ciliary pore at the base of
the cilia (Rosenbaum and Witman, 2002). Others have noted the
similarity between IFT proteins and nuclear pore proteins (Jekely
and Arendt, 2006) and the similarities of nuclear and cilia pores
(Christensen et al., 2007).
Although the trafficking of RP2 to the cilia appeared to be
regulated by Importin 2, we found that RP2 also bound to Importin
1 and Importin 3. Knockdown of Importin 1 affects cell
division, thus it is not possible to detect its affect on cilia trafficking.
Our preliminary data with knockdown of Importin 3 showed no
effect on RP2 trafficking (our unpublished observations). Binding
to Importin 2 was the strongest and was detected in three different
assays, so this might explain why cilia defects were seen primarily
with knockdown of Importin 2. We also found that the sites of
myristoylation and palmitoylation on the N-terminus of RP2 are
important for the binding of RP2 to Importin proteins. Recent
studies from Cheetham and co-workers have also found that lipid
modification is important for targeting to the cilia base (Evans et
al., 2010). We have observed similar defects in cilia trafficking of
RP2 mutated at the lipid modification sites in polarized epithelial
cells (Hurd et al., 2010). In fact, although mutation of the NLS
sequences did not affect binding of Importin 2 to RP2 we found
that mutation of the myristoylation site did impair Importin 2
binding. This is somewhat consistent with our results with
Crumbs3–CLP1 where we have found that the transmembrane
domain is essential for binding of Importin proteins (Fan et al.,
2007) (our unpublished observations). These results suggest that
membrane attachment is essential for Importin binding to these
targets, but it is not clear whether it is directly involved or is
essential for other cofactors. In this regard, it is interesting to note
that some Importin proteins have been described to be membrane
associated (Hachet et al., 2004).
It is also interesting to note that although RP2 could bind
Importin proteins, we could not detect RP2 in the nucleus of
epithelial cells despite trying several conditions, including UV
stress, previously reported to send RP2 to the nucleus in HeLa and
RPE cells (Yoon et al., 2006). If we removed the lipid anchor by
mutating Gly2 to alanine, RP2 moved to the nucleus instead of the
cilia (Hurd et al., 2010). Whether this involves binding to other
Importin proteins is unclear, because this mutation did block
Journal of Cell Science
Importin 2 and RP2
binding of Importin 2 to RP2. The role of lipid modification in
protein trafficking to the cilia and flagella has been described
(Emmer et al., 2009; Hurd and Margolis, 2009; Saric et al., 2009).
Recently Pazour and colleagues have described a cilia-trafficking
motif in the Fibrocystin gene (Follit et al., 2010). They identified
a sequence that contains lipid modification sites, as well as a
polybasic region, to be necessary for targeting a fragment of this
protein to the cilia. However, in their data they did not invoke the
Ran–Importin system, but rather describe interactions of this
sequence with Rab8.
In summary, our data indicate an important role for the Importin
proteins in cilia trafficking. Ran and Importin proteins have been
found consistently in cilia and centrosomal proteomes (Andersen
et al., 2003; Liu et al., 2007; Pazour et al., 2005), and combined
with studies such as those presented here, leave little doubt that the
Ran–Importin system is localized to the cilia and centrosome.
However, further work is required to understand their complete
role. As previously described, the Ran–Importin system could
work similarly to its role in nuclear trafficking by releasing cargo
in a Ran-dependent fashion from Importin proteins at the base of
the cilia. However the Ran–Importin system might have other
roles in the cilia that need to be considered. One is the finding that
Importin proteins have important roles in microtubule growth and
might have a role in building the cilia (Carazo-Salas et al., 2001).
Our results upon knockdown of Importin 2 showed, however,
that cilia formed normally. Another possible role is trafficking of
proteins from the cilia to the nucleus. There is an expanding
description of signaling proteins that travel from the cilia to the
nucleus, and Importin proteins in the cilia might have a role in this
process. Proteins travel from the cilia to the nucleus in several
signaling pathways (Haycraft et al., 2005; Huangfu et al., 2003;
Lal et al., 2008), suggesting that Importin proteins could also
function to traffic proteins out of the cilia. Regardless, it seems
likely that the role of the Ran–Importin system in cilia function is
a rich area for future study.
Materials and Methods
Antibodies
Rabbit anti-RP2 antibody was generated by immunization with recombinant human
RP2 (Cocalico Biologicals, Reamstown, PA). Antibody was affinity purified with
immobilized recombinant human HIS-RP2. Mouse anti-FLAG, anti-acetylated
tubulin, anti--tubulin, anti-polyglutamylated tubulin, anti-g-tubulin and rabbit antiImportin-2 antibodies were purchased from Sigma. Rabbit anti-EGFP was purchased
from Invitrogen. Rabbit anti-Myc was purchased from Bethyl labs and mouse antiImportin-1 was from Affinity Bioreagent. Rabbit anti-pericentrin was purchased
from Covance. Mouse anti-Importin-3 was purchased from Abnova. Mouse antiRan was purchased from BD Biosciences.
Constructs
All constructs were generated by PCR from full-length ESTs (Mouse Importin 1,
2 and 3; Open Biosystems, Huntsville, AL). RP2–EGFP and RP2–FLAG
expression constructs were generated by cloning the human RP2 open reading frame
into pEGFP-N1 and pQCXIP respectively. For GST fusions, open-reading frames
were cloned into unique restriction sites of pGEX-4T3 (GE Healthcare). Myc-tagged
Importin proteins and Importin-2–EGFP were cloned into the multiple cloning sites
of pQCXIH or pEGFP-N1, respectively (Clontech). For recombinant protein
expression, ORFs of Importin and Ran were cloned into pGEX4T3 (GE Healthcare).
Cell culture
MDCKII and HEK293 cells were grown in DMEM supplemented with 10% FBS,
100 U/ml penicillin, 100 g/ml streptomycin sulfate and 2 mM L-glutamine. For
generation of stable MDCK cell lines, HEK293T cells were transiently transfected
with pGAG/POL and pVSVG plus either pQCXIH for stable protein expression or
pSIREN-RetroQ for shRNA expression (Clontech). Virus-containing medium was
collected, filtered though a 0.4 m filter and mixed with an equal volume of fresh
culture medium. Polybrene (Clontech) was added to 4 g/ml and virus was added to
MDCKII cells. 12 hours later, medium was replaced. 48 hours after infection, stable
expressing pools were selected using medium containing 200 g/ml Hygromycin or
725
5 g/ml Puromycin, respectively (Invitrogen, Carlsbad, CA). All transient transfection
experiments were performed using Fugene 6 reagent (Roche).
Immunofluorescence and confocal microscopy
We performed immunofluorescence as described previously (Fan et al., 2007). In
general, cells were fixed with 4% paraformaldehyde for 15 minutes. After brief
washing, 0.25% Triton X-100 in PBS was used to permeabilize the cells in most
conditions. We used 0.1% Triton X-100 in PBS to permeabilize cells for ciliary
staining. For g-tubulin and pericentrin staining, we used cold acetone to fix the cells
for 4 minutes. These procedures were followed by 3% goat serum in PBS blocking
for 30 minutes. Primary antibodies and secondary antibodies were diluted and
incubated as previously described (Fan et al., 2007). All images were obtained using
either inverted fluorescence microscope (Nikon Eclipse TE2000U) or a meta laserscanning confocal microscope (LSM 510; Carl Zeiss MicroImaging).
Protein knockdown
Canine Importin 2 sequences GCAGTGCCTTTGCTACCTTAG and
GCATGTTAAGCCTTGTATAGC were selected as Importin 2 shRNA targeting
sequences. These sequences were placed into the Clontech shRNA Sequence Designer
Tool and the selected oligos were annealed and ligated in pSIREN-RetroQ (Clontech)
to encode the shRNA. Retroviral shRNA infection was as above.
Centrosome purification
Centrosomes were isolated using a modified protocol (Zhou et al., 2006). In brief,
MDCKII cells were grown for 6 days after confluence before incubation with 10
g/ml nocodazole and 5 g/ml cytochalasin B at 37°C for 90 minutes. Cells were
washed with PBS, 8% sucrose in 0.1 PBS, and finally 8% sucrose. Cells were
lysed (1 mM HEPES, pH 7.2, 0.5% NP-40, 0.5 mM MgCl2, 0.1% 2-mercaptoethanol,
and protease inhibitors) with shaking at 4°C for 10 minutes. Lysates were cleared by
centrifugation at 2500 g for 10 minutes and supernatant was filtered through a 40
m nylon mesh. HEPES (pH 7.2) was added to a final concentration of 10 mM and
DNaseI added to 2 U/ml followed by incubation on ice for 30 minutes. Lysates were
underlaid with 60% sucrose solution (60% wt/wt sucrose in 10 mM PIPES, pH 7.2,
0.1% Triton X-100 and 0.1% 2-mercaptoethanol) and spun at 25,000 g for 45
minutes to sediment centrosomes into the sucrose cushion. The sucrose cushion was
collected and diluted with lysis buffer to a final concentration of sucrose of 30%.
The centrosome preparation was then loaded onto a discontinuous sucrose gradient
(0.5 ml of 70%, 0.3 ml of 50% and 0.3 ml of 40% sucrose solution) and spun at
120,000 g for 60 minutes. Fractions (0.15 ml each) were collected and diluted with
1 ml of 10 mM PIPES, pH 7.2 and sedimented by centrifugation at 16,000 g for 15
minutes. For centrosomal protein analysis, the supernatant was discarded and the
pelleted protein resuspended in LDS sample buffer (Invitrogen).
Immunoprecipitation and western blotting
HEK293 and MDCK cells were rinse twice in ice-cold PBS and lysed in Triton lysis
buffer [1% Triton X-100 (v/v), 20 mM Tris-HCl, pH 7.4, 150 mM NaCl, 1.5 mM
MgCl2, 10% (v/v) glycerol] followed by centrifugation. For anti-FLAG
immunoprecipitations, FLAG M2 beads (Sigma) were added to cleared lysates and
incubated overnight at 4°C. For Myc and HA immunoprecipitations, cleared lysates
were incubated overnight at 4°C with 2 g of appropriate antibody plus 20 l
Protein-A–Sepharose (Invitrogen). Immunoprecipitations were washed four times
with lysis buffer and resuspended in LDS sample buffer (Invitrogen). Samples were
subsequently run on 4–12% Bis-Tris Novex gels (Invitrogen) and transferred to
PVDF. Membranes were blocked with 5% BSA-TBS and primary antibodies
incubated for 1 hour in blocking buffer. Secondary antibodies were added in 5%
non-fat milk in TBS with 0.05% Tween20 for 1 hour. For GST pull-down experiments,
4 g glutathione Sepharose4B immobilized GST protein was incubated with cleared
lysate overnight at 4°C. Beads were then washed four times with lysis buffer
and resuspended in LDS sampled buffer. For MS/MS analysis FLAG
immunoprecipitations were performed as described above and run on Bis-Tris gels.
Gels were subsequently washed and stained with Imperial Blue colloidal Coomassie
(Pierce) and positive bands excised. MS/MS analysis was performed by the Michigan
Proteome Consortium.
We thank Anand Swaroop, Hemant Khanna, Kristen Verhey and
Jeff Martens for support and helpful discussions. The work was
supported by a grant from the PKD Foundation and by NIH DK84725.
This work utilized the Microscopy Core of the Michigan Diabetes
Research and Training Center funded by DK020572 from the National
Institute of Diabetes and Digestive and Kidney Diseases. Deposited in
PMC for release after 12 months.
Supplementary material available online at
http://jcs.biologists.org/cgi/content/full/124/5/718/DC1
726
Journal of Cell Science 124 (5)
Journal of Cell Science
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