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Vol. 18, Issue 10, 3741-3751, October 2007
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*Department of Pathology, National Institute of Infectious Diseases, Shinjuku, Tokyo 162-8640, Japan; Nippi Research Institute of Biomatrix, Toride, Ibaraki 302-0017, Japan; and Japan Institute of Leather Research, Adachi, Tokyo 120-8601, Japan
Submitted December 19, 2006;
Revised June 27, 2007;
Accepted July 6, 2007
Monitoring Editor: Sean Munro
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). The ER is organized into a three-dimensional network of interlinked membranous tubules and cisternae throughout the cell and extends tubular structures toward the cell periphery (Baumann and Walz, 2001; Voeltz et al., 2002). Numerous studies have focused on how motor proteins mediate movement of the ER along MTs and provided evidence for their roles in MT-based ER tubule formation (Allan and Schroer, 1999; Karcher et al., 2002). A protein complex of a motor protein and a cargo receptor is assumed to be required for the transport of ER membranes along MTs (Sheetz, 1999). Although nonmotor MT-binding proteins were shown to be involved in organelle assembly and organization, the underlying molecular mechanism remains unclear. Because only a fraction of the ER in each cell is highly motile (Lee and Chen, 1988), the remaining ER, particularly that in the perinuclear region, is considered to be predominantly stationary. Further "static" association of organelles with MTs must be required for the structural organization and maintenance of the stationary ER. Indeed, accumulating evidence suggests that there are specific nonmotor MT-binding proteins for different organelles. CLIP170 was first identified as a class of cytoplasmic linker proteins that link MTs to endosomes (Pierre et al., 1992). GMAP-210 is a minus-end MT-binding protein that associates with the Golgi apparatus and plays a role in maintaining its structural integrity (Infante et al., 1999). Furthermore, ch-TOG, the human homologue of XMAP215, a previously described MT-associated protein (MAP) in Xenopus eggs, is localized at the ER membrane in interphase cells (Charrasse et al., 1998). In addition to these cytosolic factors, a new class of direct interactions between the ER membrane and MTs, which are mediated by the integral ER membrane protein CLIMP-63, has been demonstrated (Klopfenstein et al., 1998). These interactions are supposed to contribute to the positioning of the ER along MTs.
In the present study, we focused on p180, which is an integral ER membrane protein and a member of the ES/130 family. ES/130 was originally reported as a morphogenesis inducer in the embryonic chicken heart (Rezaee et al., 1993), whereas canine p180 was first identified as a ribosome-binding protein on the rough ER (Savitz and Meyer, 1990). ES130/p180 was reported to be expressed in most human tissues, and especially strongly in secretory tissues (Langley et al., 1998), although its precise role in animal cells has not yet been elucidated. Previously, we demonstrated that human cytomegalovirus (HCMV) specifically binds to human p180 via its N-terminal domain (Ogawa-Goto et al., 2002), suggesting that the interaction between p180 and the virus may facilitate intracellular transport of HCMV capsids. Because an intact MT network is required for efficient transport of HCMV capsids to the nucleus (Ogawa-Goto et al., 2003), we examined whether p180 interacts with the MT network in the host cells. Here, we present data that human p180 has a unique microtubule-binding and -bundling domain that induces acetylated MT bundles and may be involved in maintaining the normal distribution of the ER.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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) were inserted into the pcDNA4 HisMax vector (Invitrogen, Carlsbad, CA) to generate pcDNAp180-54R or pcDNAp180-24R, respectively. pEGFP-C1 and pEGFP-C2 (BD Bioscience Clontech, Palo Alto, CA) were used to construct expression plasmids for truncated forms of human p180. The entire coding sequence of enhanced green fluorescent protein (EGFP) was inserted into pcDNAp180-54R to generate wild-type (WT) p180 fused with GFP (GFP-WT). The signal sequence of calreticulin (Crt) was PCR-amplified from the pEYFP-ER vector (BD Bioscience Clontech) and introduced at the 5'end of GFP-WT to generate GFPer-WT. Various truncation mutants were generated by PCR procedures that introduced premature stop codons or by digestion with restriction enzymes that cut within the p180 coding region. The EGFP coding sequence in all the above-described EGFP-tagged constructs was substituted with EGFP-A208K from GFP-VSVG (kindly provided by Dr. Lippincott-Schwartz, National Institute of Child Health and Human Development) to express monomeric GFP (Zacharias et al., 2002). The pQE31 and pQE32 vectors (Qiagen, Valencia, CA) were used to construct bacterial expression plasmids for (His)6-tagged truncated forms of human p180. For in vitro cosedimentation assays, full-length p180 carrying 24 repeats (24R-p180), rather than 54 repeats, was used because bacterial expression of the latter was unsuccessful.
Cell Culture and Transfection
Chinese hamster ovary (CHO), COS-1, K-1034, and human diploid embryonic lung fibroblast (HEL) cells were cultured in DMEM supplemented with 10% fetal bovine serum (Intergen, New York, NY) and maintained in a humid incubator at 37°C in a 5% CO2 environment. Cells were transiently transfected with 1.5 µg of the p180 constructs or a control plasmid using Fugene-6 (Roche Diagnostics, Mannheim, Germany) or Lipofectamine-plus (Invitrogen), according to the corresponding manufacturer's instructions. Because a well-organized ER structure is most obviously visible after the addition of ascorbic acid (unpublished data), HEL cells were cultured in the presence of 200 µM ascorbic acid for studies of ER morphology. To generate a series of stable transfectants overexpressing human p180, CHO cells were transfected with pcDNAp180-54R and selected with 100 µM zeocin, followed by limited dilution. Concomitantly, control cell lines were selected after transfection with the vehicle plasmid alone. The parental CHO cells contained very little p180, whereas an unknown 160-kDa protein was detected by an anti-p180 C1 antibody (Ogawa-Goto et al., 2002). Among the obtained clones, we used six clones for assays of acetylated tubulin.
Antibodies
Affinity-purified rabbit antibodies against human p180 (C1 and N1) were described previously (Ogawa-Goto et al., 2002). Monoclonal antibodies (mAbs) against the N-terminal domain of human p180 were established by immunization of BALB/c mice with a recombinant (His)6-tagged polypeptide containing amino acids 27–157 by Nippon Biotest (Tokyo, Japan). The established clones, 4H6 (IgG1, kappa chain) and 1E3 (IgG2b, kappa chain), specifically bound to the N-terminal region of p180 in Western blot and ELISA analyses (data not shown). Rabbit polyclonal antibodies against GFP (Clontech), protein disulfide isomerase (PDI) and calnexin (Cnx; Stressgen, San Diego, CA), mAbs against acetylated tubulin (6-11B-1) and 
-tubulin (B-51-2; Sigma-Aldrich, St. Louis, MO), and goat antibodies against Crt and ribophorin II (Santa Cruz Biotechnology, Santa Cruz, CA) were obtained from the respective suppliers.
Immunofluorescence Microscopy
Cultured cells were fixed with paraformaldehyde, permeabilized with 0.1% Triton X-100, and incubated with antibodies as described previously (Ogawa-Goto et al., 2002). The cells were imaged using an LSM410 confocal microscope system with a 63x 1.4 NA Plan-Apochromatic DIC objective lens (Carl Zeiss, Jena, Germany). In some experiments, an FV100 confocal microscope system with a 40x 1.3 NA lens was used (Olympus, Tokyo, Japan). For measurement of the ER area, fluorescence signals of PDI or Crt staining were captured using the LSM410 system. Signals of control and transfected cells were captured by the same acquisition processes and quantified using the public domain software Image J (http://rsb.info.nih.gov/ij). The ER area (in pixels) per cell was profiled and extracted by "particle analysis" with a constant threshold. In a preliminary experiment, the total cell number in the field was simultaneously counted after nuclear staining and indicated that the process could extract appropriate numbers of cells. The average ER area (in pixels) per cell was evaluated in at least 100 cells and compared with that in concomitantly prepared control cells.
Analyses of the Membrane Fraction of p180-transfected Cells
Transiently transfected cells or stable transfectants were washed with phosphate-buffered saline (PBS), scraped and collected by brief centrifugation. The recovered cells were suspended in 50 mM Tris-HCl buffer containing 5 mM MgSO4, 5 mM EGTA, 1 mM EDTA, and Complete proteinase inhibitor mixture (Roche Diagnostics) and homogenized with a glass homogenizer until there were no intact cells in the suspension, as evaluated by light microscopy. The cell lysate was then centrifuged at 1400 x g for 10 min, and the postnuclear supernatant was ultracentrifuged at 100,000 x g for 1 h to sediment the membrane fraction. Soluble proteins in the resulting supernatant were precipitated by adding trichloroacetic acid solution on ice and collected by centrifugation. The samples were analyzed by SDS-PAGE and Western blotting.
In Vitro MT-Binding Assays with Recombinant Polypeptides
Glutathione S-transferase (GST)-fusion proteins of various regions of p180 were prepared as described previously (Ogawa-Goto et al., 2002). Bacterially expressed (His)6-tagged polypeptides were purified using Talon resin (BD Bioscience Clontech) according to the manufacturer's instructions. MTs were polymerized by incubating 5 mg/ml bovine brain tubulin (Cytoskeleton, Denver, CO) in 100PEM buffer (100 mM Pipes, pH 6.8, 2 mM MgCl2, 1 mM EGTA) containing 1 mM GTP and 10% glycerol for 20 min at 35°C. The polymerized MTs were diluted to 0.5 mg/ml, incubated with 20 µM taxol (Sigma-Aldrich) in 100PEM buffer for 10 min, and then mixed with various polypeptides to a final concentration of 2 µM in a total volume of 50–100 µl. The final concentration of NaCl was adjusted to 120 mM unless otherwise described, and the samples were incubated for 30 min. Aliquots of the reaction mixtures were layered onto 100 µl of a cushion buffer (100PEM buffer containing 10 µM taxol and 10% glycerol) and centrifuged in a TLS55 or TLA100 rotor (Beckman, Fullerton, CA) at 30,000 x g for 30 min at 25°C. The resulting supernatants and pellets were analyzed by SDS-PAGE. In some experiments, bundle formation was monitored by the OD at 350 nm in a DU530 spectrophotometer (Beckman).
Chemical Cross-Linking of (His)6-tagged Polypeptides
An aliquot (200 µg/ml) of each (His)6-tagged recombinant polypeptide in 100 mM phosphate buffer (pH 7.4) was cross-linked by adding an equal volume of 20 mM dimethyl pimelimidate (DMP) dihydrochloride (Pierce, Rockford, IL) in 0.2 M triethanolamine (pH 8.2). After a 30-min incubation at room temperature, the reaction was stopped by the addition of 0.5 M Tris-HCl (pH 8.0), and the samples were analyzed by SDS-PAGE.
In Vitro MT-Binding Assays with Full-Length p180
MT-binding assays of immunoprecipitated p180 were carried out as follows. The antibody-antigen complexes were recovered from cell lysates using an anti-C1 or anti-GST antibody and protein A–conjugated magnetic beads (Dynabeads protein A; Dynal Biotech, Oslo, Norway) according to the manufacturer's instructions. After washing, the magnetic beads were blocked with 2% bovine serum albumin in PBS, washed with buffer (50 mM HEPES, pH 7.4, 2 mM EGTA, 100 mM NaCl, 1 mM MgCl2), and incubated with taxol-stabilized MTs in the same buffer for 1 h at room temperature. After extensive washing, the beads were recovered and subjected to SDS-PAGE and Western blot analyses. The presence of MTs in the bead fraction was detected with an anti--tubulin antibody. SDS-PAGE and Western blotting of p180 were carried out as described previously (Ogawa-Goto et al., 2002).
To prepare cell lysates for in vitro cosedimentation assays, cells were lysed in lysis buffer (50 mM Tris-HCl, pH 7.4, 5 mM EDTA, 150 mM NaCl, 1% Nonidet P-40, 1% Triton X-100) containing Complete proteinase inhibitor mixture for 20 min on ice and then centrifuged at 8000 x g for 30 min. The soluble fraction was further clarified at 25,000 x g for 30 min, and the supernatant was diluted 1:2.5 with MT-binding buffer (80 mM PIPES, pH 6.9, 2 mM EGTA, 100 mM NaCl, 1 mM MgCl2) containing Complete proteinase inhibitor mixture, 2 mM DTT, and 10 µM taxol. For cosedimentation assays, in vitro–polymerized MTs (10 µg/assay, as described above) were added to 100 µl of the diluted lysates and incubated for 30 min at room temperature. The samples were centrifuged at 20,500 x g, and the supernatants and pellets were analyzed by SDS-PAGE and Western blotting. In normal assays, the final concentration of NaCl was adjusted to 120 mM unless otherwise described.
Electron Microscopy
For electron microscopic analysis of in vitro–polymerized MTs, samples were prefixed with paraformaldehyde in PBS containing 10 µM taxol, fixed with glutaraldehyde, and negatively stained with 1% uranyl acetate. For transmission electron microscopy (TEM) analysis of cultured cells, cells were treated with medium containing 10 µM taxol for 3 min at 37°C and then prefixed with 4% paraformaldehyde in PBS containing 10 mM EGTA and 2 mM MgCl2 for 20 min at 37°C, followed by a conventional fixation procedure for TEM as described previously (Ogawa-Goto et al., 2002). Ultrathin sections were cut parallel to the substrate. MTs were identified by their characteristic tubular structure with a diameter of 24 nm.
Atomic Force Microscopy
Atomic force microscopic analyses were carried out using an SPM-9500J2 scanning probe microscope (Shimadzu, Kyoto, Japan) equipped with a piezoelectric scanner, of which the maximum widths in the x, y, and z scan range were 125, 125, and 8 µm, respectively. A rectangular cantilever with a tetrahedral silicon tip was used at a force constant of 42 N/m and a resonance frequency of 300 kHz (OMCL-AC160TS; Olympus). Both height and phase images were obtained simultaneously in a dynamic mode in air. The scan speed was generally maintained at 1 Hz. The MT bundles were fixed with 4% paraformaldehyde and deposited on glass coverslips. After extensive washing with PBS and distilled water, the samples were air-dried and processed for atomic force microscopic analyses.
Small Interfering RNA Oligonucleotides and Transfection
RNA duplexes (21 nucleotides) with symmetric 3'(2'-deoxy)thymidine overhangs (two nucleotides) corresponding to nucleotides 153–173 of human p180 were synthesized and annealed (Qiagen). Transfection of the small interfering RNAs (siRNAs) was carried out using Oligofectamine (Invitrogen, Rockville, MD) as previously described (Elbashir et al., 2001), and the cells were incubated for 2–4 d. For control experiments, fluorescein isothiocyanate–labeled nonsilencing oligonucleotides (Qiagen) were used.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; MacRae, 1997). Acetylated MTs were mainly detected in the perinuclear region of the control cell lines (Figure 1Aa). In contrast, the p180-overexpressing cell lines exhibited more organized acetylated MTs that extended to the distal region (Figure 1A, b and c). As shown in Figure 1B, Western blotting analysis revealed that acetylated tubulin was mildly increased in the transfectants (lanes 1 and 2) to 
1.71-fold the level in the parental CHO cells (lane 5), whereas acetylated tubulin in the control cell lines (lanes 3 and 4) was 1.14-fold the level in the parental cells. Electron microscopic analyses revealed that laterally coaligned and packed MT bundles were present in a wide area of the cytoplasm in the transfectants (Figure 1C), as has been shown for HEL cells (Ogawa-Goto et al., 2003). In these transfectants, p180 was found to associate with membranes (Figure 1D, lane 2), but was not detected in the soluble fraction (lane 3). Furthermore, enhanced acetylated MT arrays were observed after transient transfection in CHO cells (Figure 1E, left) and other cell types, including HeLa cells (data not shown). In the transiently p180-overexpressing cells, acetylated MT arrays were more markedly enhanced (Figure 1E, left) than in the stable transfectants (Figure 1A, b and c), probably because a higher level of p180 was expressed in each single cell that had been transiently transfected (Figure 1E, right).
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). At 3 d after transfection, the level of p180 protein was suppressed to <30% of the level in control cells (Figure 2A, top). Control cells transfected with a nonsilencing siRNA contained well-developed acetylated MT bundles (Figure 2Bb), whereas p180-depleted cells displayed immature acetylated MTs (Figure 2Ba). The number of cells possessing extended acetylated MTs was markedly reduced to about one-fourth of the number in control cells (Figure 2C). Consistently, decreased amounts of acetylated tubulin were also detected in p180-depleted cells by Western blotting analyses (Figure 2A, bottom). These results suggest a crucial role for p180 in the organization of acetylated MTs.
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), the next question to be addressed was whether endogenous p180 affected the overall MT organization and ER morphology. The MT network in p180-depleted cells stained for -tubulin appeared to change to a radial projection or unaligned random pattern (Figure 2Da2), whereas that in control cells showed laterally coaligned arrays (Figure 2Db2). The same results were observed for K-1034 cells (Figure 2D, c2 and d2), which are retinal pigment epithelial cells that also express a high level of p180. For both types of cells, the numbers of cells possessing a radial MT or nonparallel pattern were greatly increased among p180-depleted cells (Figure 2E). Simultaneously, targeted depletion of p180 severely affected the normal ER distribution, which retracted around the nucleus (Figure 2Da3). This ER retraction was also detected using other ER markers, such as PDI and Cnx (not shown), and differing degrees of p180 depletion seemed to be coincidentally associated with the severity of the ER retraction. Semiquantitative analyses revealed that the transfected cells exhibited less effective ER spreading than control cells (Figure 2F). Thus, depletion of endogenous p180 markedly affects the ER distribution and causes it to retract from the cell periphery, with a concomitant redistribution of MTs to a radial pattern in these cells.
Binding of Full-Length p180 to In Vitro–polymerized MTs
The rearrangement of MTs and the ER in p180-depleted cells suggested a specific interaction between p180 and MTs. To examine whether p180 binds directly to MTs in vitro, bacterially expressed full-length p180 was prepared and subjected to MT-binding assays. For these experiments, we used a splice variant of p180 carrying 24 repeats and the complete N- and C-terminal domains (24R-p180). 24R-p180 was recovered from bacterial lysates by protein A–conjugated magnetic beads using an anti-p180 antibody and incubated with in vitro–polymerized MTs (Figure 3A). The presence of MTs bound to p180 was detected using an anti-tubulin antibody. 24R-p180 exhibited effective interactions with MTs (lane 4), whereas beads with a control antibody did not (lane 6).
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Binding of p180-derived Polypeptides to MTs and Promotion of Bundle Formation In Vitro
To determine the domain responsible for p180 binding to MTs, a panel of GST-fusion proteins containing different portions of p180 was prepared as shown in Figure 4A. Based on its deduced primary sequence, p180 is predicted to be a membrane protein with a short luminal tail and to consist of a highly basic N-terminal region and an acidic C-terminal region comprising 15 segments of predicted coiled-coil domains (Wanker et al., 1995; Langley et al., 1998). MT cosedimentation assays (Figure 4B) revealed that a bacterially expressed N1 polypeptide effectively bound to taxol-stabilized MTs (lane 1), whereas other more COOH-terminal fragments (residues 853–1340; lane 9) and control GST (lane 5) did not. An NH2-terminal fragment, residues 25–157, which resides downstream of the predicted transmembrane domain, was also cosedimented (lane 13), indicating the presence of a second, but less effective, MT-binding domain. To test the interaction of N1 with MTs in more detail, truncated polypeptides containing only the repeat domain, N1
C (residues 623–737), or a flanking coiled-coil region, N1Rp (residues 738–944), were prepared. For this assay, we used (His)6-tagged polypeptides, because GST-tagged N1Rp appeared to be extremely unstable under the binding assay conditions. Although N1C failed to bind to MTs (Figure 4B, lane 21), the coiled-coil region N1Rp showed MT-binding ability (lane 17).
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Rp, suggesting bundle formation of the MTs. Electron microscopic analysis revealed that N1 induced dense and laterally aligned MT bundles within 1 min after mixing (Figure 4C, left). By atomic force microscopy, the diameter of the largest N1-induced bundle in Figure 4D (left) was estimated to be 109.5 ± 8.7 nm in height after fixation and dehydration, whereas that of a single filament after incubation with GST (Figure 4D, right) was 10.7 ± 0.49 nm, similar to a previously reported value for air-dried MTs (Vater et al., 1995). These results indicate that at least 10 MTs were assembled in the N1-induced bundle. (His)6-tagged N1 induced similar MT bundles to GST-N1 (Supplementary Figure S1A, c and d) and its bundling activity appeared to be much higher than that of N1Rp (Supplementary Figure S1B). Moreover, monitoring of the turbidity of the N1-induced bundles after calcium addition revealed a remarkable resistance toward calcium-dependent depolymerization (Figure 4E). On the basis of these results, we designated the domain at amino acids 623–944 as MTB-1 and the second potential MTB in the N-terminal region (amino acids 25–157) as MTB-2. Furthermore, in vitro sedimentation assays with full-length p180 (Figure 3C) revealed that pretreatment with an anti-N1 antibody (lane 5), but not with a control anti-GST antibody (lane 1), resulted in reduced p180 binding to MTs, suggesting the importance of this domain. Thus, p180 contains a unique MTB domain consisting of a predicted coiled-coil region and repeat domain that exhibits a direct binding capacity for MTs as well as a strong bundling activity in vitro.
Dimer Formation by the MTB-1 Domain
The MT-bundling capacity of the MTB-1 domain implied that it could form a dimer. To address this possibility, we assessed the dimer-forming abilities of (His)6-tagged N1 and other polypeptides in vitro by chemical cross-linking (Figure 5). After treatment with the cross-linker DMP, both His-N1 (Figure 5, lane 2) and His-N1Rp (lane 4) moved more slowly upon SDS-PAGE analysis and were detected at their expected dimer positions, whereas His-N1C remained unchanged (lane 6). These data suggest that MTB-1 forms a dimer naturally and that the responsible domain for dimer formation is the predicted coiled-coil region (N1Rp).
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). After transfection, Western blot analyses of the chimeras revealed that truncated proteins of the correct sizes were expressed (Figure 6B). For assessment of bundling activities, we examined the effect of each construct on MT organization by -tubulin staining. Overexpression of GFP-N1 produced a perinuclear fibrillar pattern that was coaligned with MTs (Figure 6Ca). Even at very low expression levels, GFP-N1 was colocalized with MTs (Supplementary Figure S2a). At very high expression levels, GFP-N1 induced extended and wavy MT bundles (Supplementary Figure S2b), which were also positive for acetylated tubulin (Supplementary Figure S3). A truncated construct, GFP-N1Rp, exhibited a similar, but less effective phenotype to GFP-N1 (Figure 6Cb), whereas the repeat domain alone, GFP-N1C, did not (Figure 6Cc). None of the other GFP-tagged proteins exerted MT-bundling activities, including GFP-1-157 and GFP-27-157 containing the second potential MTB. The phenotype of GFP-WT seemed to be somewhat similar to that of GFP-N1, although the MTs appeared to be crowded and to cross randomly (Figure 6Ce). These results indicate that the MTB-1 domain consisting of the repeat domain and coiled-coil region is capable of binding to MTs and promoting bundle formation in vivo when expressed alone. Again, a specific region of human p180 corresponding to amino acids 738–944 emerged as the minimum domain required for MT binding, although its bundling activity was less than that of MTB-1.
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12% of the cells expressing high levels of GFPer-WT (Figure 7D, top). In the peripheral region of these cells, the GFPer-WT signals showed a fibrillar pattern that was coaligned with an extended meshwork of acetylated MTs (Figure 7Ca). Deletion of the COOH-terminal region led to a striking phenotype. Specifically, the cells expressing high levels of GFPer-956 contained long and wavy bundles of acetylated MTs, which mostly overlapped with the GFPer-956 signals (Figure 7Cb). When stained for -tubulin, less clear, but distinct, bundling was observed in cells expressing GFPer-956 (Supplementary Figure S5b). In contrast, the MTB-1 deletion mutant (GFPer-623–956) failed to induce perinuclear acetylated MT bundles, even at very high levels of expression (Figure 7Cc). Because the MTs in some GFPer-WT–expressing cells were partially resistant to the MT depolymerization reagent nocodazole (data not shown), p180 overexpression seemed to moderately stabilize MTs, but less efficiently than has been reported for MAP2 or tau-transfected cells (Kanai et al., 1989; Lee and Rook, 1992). Thus, p180 led to enhanced acetylated MTs in an MTB-1–dependent manner.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). Although previous studies using heterogonous overexpression of canine p180 in yeast indicated its involvement in ER membrane proliferation and secretory pathways (Wanker et al., 1995; Becker et al., 1999), very little is known about its roles in mammalian cells. Here, we have demonstrated a new functional role for p180 in interactions between the ER and the MT cytoskeleton, which are mainly mediated by a newly identified domain designated MTB-1.
MT-Binding Properties of p180
We have presented data that MTB-1 promotes MT bundling both in vivo and in vitro when expressed alone, similar to other MAPs that are considered to induce MT assembly and stabilization in vivo (Mandelkow and Mandelkow, 1995). The bundling activity of MTB-1 was as strong as those of structural MAPs, although the primary sequence of the domain does not contain the typical repeat sequence found in MAPs (Lewis et al., 1988). Motif analyses previously predicted that the carboxyl half of p180 would form a rod-like coiled-coil structure (Leung et al., 1996; Langley et al., 1998). MTB-1 consists of this coiled-coil region and the repeat domain. Furthermore, the coiled-coil domain appeared to be the domain responsible for the dimeric conformation of p180, which is assumed to be a prerequisite for MT bundling, whereas the repeat region has been reported to act as the ribosome-binding domain (Wanker et al., 1995). MTB-1–induced MT bundles exhibited remarkable resistance toward calcium-induced depolymerization in vitro, although the underlying mechanism of this high resistance toward calcium remains to be clarified. Furthermore, full-length p180 was shown to bind directly to MTs in vitro using recombinant proteins. Because an anti-MTB-1 antibody abolished the MT-binding activity of p180, the binding is assumed to be domain-dependent, at least partially. Thus, full-length p180 seems likely to exhibit its MT-modulating activity mainly via MTB-1 and to take a dimeric or oligomeric conformation, although we cannot rule out the possibility that other regions could play roles in MT modulation in vivo. Indeed, we also identified the MTB-2 domain, which shows a lesser MT-bundling activity.
p180 Is an MT-Binding Protein on the ER
Overexpression of p180 led to increased acetylation of -tubulin, as has been reported for MAPs (Kanai et al., 1989). Consistently, p180 depletion resulted in reduced amounts of acetylated MTs. Because acetylated tubulin is a marker of nondynamic stable MTs, but absent from more dynamic MT structures (Bulinski and Gundersen, 1991; MacRae, 1997), the expression levels of p180 seem to affect the dynamic instability of the MT cytoskeleton. Moreover, our EM study clearly revealed MT bundles in p180-overexpressing transfectants, suggesting that MT bundling and stabilization may be coupled, as reported previously (Chapin et al., 1991). Overexpression of p180 led to the induction of acetylated MTs in an MTB-1–dependent manner. The MT-modulating activity of p180 was further verified by siRNA experiments showing that p180 is required for maintaining the overall parallel MT patterns and ER distribution. It is noteworthy that p180 at its normal expression level was sufficient to maintain the MTs and ER in both HEL (fibroblastic) and K-1034 (epithelial) cells. Moreover, our finding of ER retraction after p180 depletion appears to be reminiscent of the ER phenotype after kinesin suppression (Feiguin et al., 1994). These results imply roles for p180 in linking the ER and MTs and maintenance of the ER, as has been reported for CLIMP-63 (Klopfenstein et al., 1998), although the definitive role of p180 in ER morphology remains to be clarified in further studies. It is also unclear whether the elongated cell morphology is a direct consequence of p180 overexpression. Nevertheless, it is likely that high levels of p180 expression would be involved in maintaining the MT bundles and consequently affect the cell shape via an altered cytoskeleton network. This idea is consistent with our frequent observations that long cellular extensions formed after p180 overexpression were filled with acetylated MT bundles and p180. In cells expressing high levels of p180, effective mass transport of the ER may occur toward peripheral sites where the surface membrane is actively expanding.
Although the ER tubular network can be formed in vitro without an MT cytoskeleton (Dreier and Rapoport, 2000), it is clear that the ER is intimately associated with MTs as a framework (Baumann and Walz, 2001; Voeltz et al., 2002), and that a kinesin motor-dependent process is crucial for ER membrane motility in some mammalian cells (Cole and Lippincott-Schwartz, 1995). Recently, several nonmotor MT-binding proteins have been reported (Vedrenne and Hauri, 2006), and these are either integrated into the ER membrane (Klopfenstein et al., 1998) or associated with the membrane surface like p22 (Andrade et al., 2004). Among these, CLIMP-63 was first identified as a class of direct linkers between the ER and MTs that contribute to the positioning of the ER (Klopfenstein et al., 1998). Based on our data, p180 would be included in this class of protein. The ER phenotype after overexpression of p180 appears to be similar to that after overexpression of CLIMP-63, although their primary sequences show no obvious homology. In particular, the sizes of their cytoplasmic domains differ greatly (106 amino acids for CLIMP-63 vs. more than 1500 amino acids for p180), implying that the two proteins have distinct modes of binding.
The Cytoplasmic Region of p180 Acts as a Multifunctional Domain
Several factors have been reported to bind to the huge cytoplasmic domain of p180, which is comprised of tandem repeats and a coiled-coil rod (Kim et al., 2002; Ogawa-Goto et al., 2002; Diefenbach et al., 2004). Importantly, a potential kinesin-binding domain has been reported downstream of MTB-1 (Diefenbach et al., 2004), although an in vivo interaction with kinesin has not been reported. Indeed, we did not detect kinesin in our samples immunoprecipitated with p180 in HEL cells (data not shown). It remains to be clarified whether p180 interacts efficiently with kinesin in vivo using other samples containing abundant kinesin.
Interestingly, p180 shares highly homologous regions with kinectin (Leung et al., 1996; Langley et al., 1998), an ER membrane protein that binds to kinesin motors (Toyoshima et al., 1992). We examined whether the conserved domain binds to MTs in vivo and in vitro, but failed to detect any interactions. Because the primary sequence of p180 predicts many, as-yet unidentified, functional domains, further studies on p180 may provide new insights into our understanding of the association between the ER and the MT network.
Weak Bundling Activities of WT p180 in Cells
The MT-bundling activity of the C-terminal truncation mutant GFPer-956 was extremely strong and was similar to that of MTB-1. Compared with the mutant, the effects of WT p180 overexpression were unexpectedly low, suggesting the presence of an inhibitory effect of its own C-terminal region. Similarly, the MT-stabilizing effect of GFPer-956 was apparently higher than that of GFPer-WT (data not shown), although both are less efficient than the reported effects of neuronal MAPs, such as MAP2 and tau (Kanai et al., 1989; Lee and Rook, 1992). The highly stable and rigid MT bundles induced by the neuronal MAPs are assumed to play roles in neuronal functions. However, in the case of fibroblastic or epithelial cells, the formation of highly stable MT bundles may rather impede their normal cellular processes, and the MAP-like activity of p180 may be negatively regulated in order to avoid the formation of MT bundles that are too rigid. Although the underlying mechanism remains to be elucidated, one possibility is that an inhibitory factor that binds to the C-terminal coiled-coil region may negatively control the MT-bundling activity of MTB-1. Alternatively, some modifications, such as phosphorylation, may be involved in the exhibition of its full activity in vivo, as has been demonstrated for several structural MAPs (Mandelkow and Mandelkow, 1995).
p180 and ER Morphology
Remarkable proliferation of rough ER membranes has been reported in yeast cells overexpressing canine p180 in a repeat domain-dependent manner (Becker et al., 1999). However, our electron microscopic studies revealed that the ER phenotypes of p180 transfectants varied among cell lines of different origins (unpublished data). The major findings included proliferation of anastomosing smooth ER along the MT bundles. Occasionally, a mild increase and redistribution of the rough ER were observed. The reason for the variety of phenotypes is unknown at present, but one reason may be possible in-frame splice variants that possess different lengths of repeats (Wanker et al., 1995; Langley et al., 1998). Further studies are currently being undertaken to elucidate the definitive effects of p180 on ER morphology. Interestingly, an early EM study on chondroblasts strongly suggested a close relationship between the rough ER and MTs (Tokunaka et al., 1983). They have clearly demonstrated that ribosomes have been selectively displaced by MTs on the rough ER membrane in chondroblasts after taxol treatment. Our identification of the MT-binding features of p180, which also has a ribosome-binding domain (Wanker et al., 1995), may provide further insights into the intimate interactions between the rough ER and MTs.
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ACKNOWLEDGMENTS |
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Footnotes |
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The online version of this article contains supplemental material at MBC Online (http://www.molbiolcell.org). 
Address correspondence to: Kiyoko Ogawa-Goto (kgoto{at}nippi-inc.co.jp)
Abbreviations used: Cnx, calnexin; Crt, calreticulin; CLIMP-63, cytoskeleton-linking membrane protein of 63-kDa; CLIP, cytoskeleton-linking protein; DMP, dimethyl pimelimidate; ER, endoplasmic reticulum; HCMV, human cytomegalovirus; HEL, human embryonic lung; MAP, microtubule-associated protein; MT, microtubule; PDI, protein disulfide isomerase; TEM, transmission electron microscopy; WT, wild type.
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REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Andrade, J., Zhao, H., Titus, B., Timm Pearce, S., and Barroso, M. (2004). The EF-hand Ca2+-binding protein p22 plays a role in microtubule and endoplasmic reticulum organization and dynamics with distinct Ca2+-binding requirements. Mol. Biol. Cell 15, 481–496. Epub 2003 Dec 2002.
Baumann, O., and Walz, B. (2001). Endoplasmic reticulum of animal cells and its organization into structural and functional domains. Int. Rev. Cytol 205, 149–214.[Medline]
Becker, F., Block-Alper, L., Nakamura, G., Harada, J., Wittrup, K. D., and Meyer, D. I. (1999). Expression of the 180-kD ribosome receptor induces membrane proliferation and increased secretory activity in yeast. J. Cell Biol 146, 273–284.
Bulinski, J. C., and Gundersen, G. G. (1991). Stabilization of post-translational modification of microtubules during cellular morphogenesis. Bioessays 13, 285–293.[CrossRef][Medline]
Chapin, S. J., Bulinski, J. C., and Gundersen, G. (1991). Microtubule bundling in cells. Nature 349, 24.[Medline]
Charrasse, S., Schroeder, M., Gauthier-Rouviere, C., Ango, F., Cassimeris, L., Gard, D. L., and Larroque, C. (1998). The TOGp protein is a new human microtubule-associated protein homologous to the Xenopus XMAP215. J. Cell Sci 111, 1371–1383.[Abstract]
Cole, N. B., and Lippincott-Schwartz, J. (1995). Organization of organelles and membrane traffic by microtubules. Curr. Opin. Cell Biol 7, 55–64.[CrossRef][Medline]
Diefenbach, R. J., Diefenbach, E., Douglas, M. W., and Cunningham, A. L. (2004). The ribosome receptor, p180, interacts with kinesin heavy chain, KIF5B. Biochem. Biophys. Res. Commun 319, 987–992.[CrossRef][Medline]
Dreier, L., and Rapoport, T. A. (2000). In vitro formation of the endoplasmic reticulum occurs independently of microtubules by a controlled fusion reaction. J. Cell Biol 148, 883–898.
Elbashir, S. M., Harborth, J., Lendeckel, W., Yalcin, A., Weber, K., and Tuschl, T. (2001). Duplexes of 21-nucleotide RNAs mediate RNA interference in cultured mammalian cells. Nature 411, 494–498.[CrossRef][Medline]
Feiguin, F., Ferreira, A., Kosik, K. S., and Caceres, A. (1994). Kinesin-mediated organelle translocation revealed by specific cellular manipulations. J. Cell Biol 127, 1021–1039.
Infante, C., Ramos-Morales, F., Fedriani, C., Bornens, M., and Rios, R. M. (1999). GMAP-210, A cis-Golgi network-associated protein, is a minus end microtubule-binding protein. J. Cell Biol 145, 83–98.
Kanai, Y., Takemura, R., Oshima, T., Mori, H., Ihara, Y., Yanagisawa, M., Masaki, T., and Hirokawa, N. (1989). Expression of multiple tau isoforms and microtubule bundle formation in fibroblasts transfected with a single tau cDNA. J. Cell Biol 109, 1173–1184.
Karcher, R. L., Deacon, S. W., and Gelfand, V. I. (2002). Motor-cargo interactions: the key to transport specificity. Trends Cell Biol 12, 21–27.[CrossRef][Medline]
Kim, M., Ogawa, H., Kohu, K., Ichikawa, M., Satoh, K., Ishidao, T., Nada, S., and Akiyama, T. (2002). Binding of the mammalian homolog of the Drosophila discs large tumor suppressor protein to the ribosome receptor. Biochem. Biophys. Res. Commun 294, 1151–1154.[CrossRef][Medline]
Klopfenstein, D. R., Kappeler, F., and Hauri, H. P. (1998). A novel direct interaction of endoplasmic reticulum with microtubules. EMBO J 17, 6168–6177.[CrossRef][Medline]
Langley, R. et al. (1998). Identification of multiple forms of 180-kDa ribosome receptor in human cells. DNA Cell Biol 17, 449–460.[Medline]
Lee, C., and Chen, L. B. (1988). Dynamic behavior of endoplasmic reticulum in living cells. Cell 54, 37–46.[CrossRef][Medline]
Lee, G., and Rook, S. L. (1992). Expression of tau protein in non-neuronal cells: microtubule binding and stabilization. J. Cell Sci 102, 227–237.
Leung, E., Print, C. G., Parry, D. A., Closey, D. N., Lockhart, P. J., Skinner, S. J., Batchelor, D. C., and Krissansen, G. W. (1996). Cloning of novel kinectin splice variants with alternative C-termini: structure, distribution and evolution of mouse kinectin. Immunol. Cell Biol 74, 421–433.[Medline]
Lewis, S. A., Wang, D. H., and Cowan, N. J. (1988). Microtubule-associated protein MAP2 shares a microtubule binding motif with tau protein. Science 242, 936–939.
MacRae, T. H. (1997). Tubulin post-translational modifications—enzymes and their mechanisms of action. Eur. J. Biochem 244, 265–278.[Medline]
Mandelkow, E., and Mandelkow, E. M. (1995). Microtubules and microtubule-associated proteins. Curr. Opin. Cell Biol 7, 72–81.[CrossRef][Medline]
Ogawa-Goto, K., Irie, S., Omori, A., Miura, Y., Katano, H., Hasegawa, H., Kurata, T., Sata, T., and Arao, Y. (2002). An endoplasmic reticulum protein, p180, is highly expressed in human cytomegalovirus-permissive cells and interacts with the tegument protein encoded by UL48. J. Virol 76, 2350–2362.
Ogawa-Goto, K., Tanaka, K., Gibson, W., Moriishi, E., Miura, Y., Kurata, T., Irie, S., and Sata, T. (2003). Microtubule network facilitates nuclear targeting of human cytomegalovirus capsid. J. Virol 77, 8541–8547.
Pierre, P., Scheel, J., Rickard, J. E., and Kreis, T. E. (1992). CLIP-170 links endocytic vesicles to microtubules. Cell 70, 887–900.[CrossRef][Medline]
Rezaee, M., Isokawa, K., Halligan, N., Markwald, R. R., and Krug, E. L. (1993). Identification of an extracellular 130-kDa protein involved in early cardiac morphogenesis. J. Biol. Chem 268, 14404–14411.
Savitz, A. J., and Meyer, D. I. (1990). Identification of a ribosome receptor in the rough endoplasmic reticulum. Nature 346, 540–544.[CrossRef][Medline]
Sheetz, M. P. (1999). Motor and cargo interactions. Eur. J. Biochem 262, 19–25.[Medline]
Terasaki, M., Chen, L. B., and Fujiwara, K. (1986). Microtubules and the endoplasmic reticulum are highly interdependent structures. J. Cell Biol 103, 1557–1568.
Tokunaka, S., Friedman, T. M., Toyama, Y., Pacifici, M., and Holtzer, H. (1983). Taxol induces microtubule-rough endoplasmic reticulum complexes and microtubule-bundles in cultured chondroblasts. Differentiation 24, 39–47.[CrossRef][Medline]
Toyoshima, I., Yu, H., Steuer, E. R., and Sheetz, M. P. (1992). Kinectin, a major kinesin-binding protein on ER. J. Cell Biol 118, 1121–1131.
Vater, W., Fritzsche, W., Schaper, A., Bohm, K. J., Unger, E., and Jovin, T. M. (1995). Scanning force microscopy of microtubules and polymorphic tubulin assemblies in air and in liquid. J. Cell Sci 108, 1063–1069.[Abstract]
Vedrenne, C., and Hauri, H. P. (2006). Morphogenesis of the endoplasmic reticulum: beyond active membrane expansion. Traffic 7, 639–646.[CrossRef][Medline]
Voeltz, G. K., Rolls, M. M., and Rapoport, T. A. (2002). Structural organization of the endoplasmic reticulum. EMBO Rep 3, 944–950.[CrossRef][Medline]
Wanker, E. E., Sun, Y., Savitz, A. J., and Meyer, D. I. (1995). Functional characterization of the 180-kD ribosome receptor in vivo. J. Cell Biol 130, 29–39.
Zacharias, A. D., Violin, J. D., Newton, A. C., and Tsien, R. Y. (2002). Partitioning of lipid-modified monomeric GFPs into membrane microdomains of live cells. Science 296, 913–916.
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-actin. The transfectants contain 
3-fold longer than the length of the nuclear long axis. Data represent the means ± SD of three separate experiments. (D) Control (b and d) or siRNA-transfected (a and c) cells were fixed at 72 h after transfection and stained for 
) or bundled MTs (
) were counted. A typical example of cells displaying circular MTs is shown in Figure S2c, whereas a typical example of bundled MTs is shown in Figure S2b. Data are expressed as percentages of the total number of GFP-expressing cells and represent the means of two separate experiments. (E) COS-1 cells were transfected with GFP-N1, fixed at 16 h after transfection, and subjected to TEM analysis. In the perinuclear region of these cells, laterally coaligned MTs (arrows in top panel) are observed, which overlap with undefined high-density structures. Loose and twisted MT bundles can also be seen around the structures (bottom). Bars, 250 nm.